Ecophysiological Studies on Dinophysis and Its Food Chain, And

THESE DE DOCTORAT DE

L'UNIVERSITE DE NANTES

ECOLE DOCTORALE N° 598 Sciences de la Mer et du littoral Spécialité : Biologie marine

Par Sylvain GAILLARD Études écophysiologiques sur Dinophysis et sa chaîne alimentaire, et effets in vitro du dinoflagellé et de ses toxines sur les premiers stades de vie de deux modèles d'animaux marins (l’huître et le poisson) “Ecophysiological studies on Dinophysis and its food chain, and in vitro effects of the dinoflagellate and its toxins on early life stages of two models of marine animals (oyster and fish)” Thèse présentée et soutenue à Nantes, le 27 novembre 2020 Unité de recherche : Laboratoire Phycotoxines (PHYC), IFREMER Centre Atlantique de Nantes

Rapporteurs avant soutenance :

Beatriz REGUERA Directrice de recherche, Instituto Español de Oceanografia - Centro Oceanográfico de Vigo Per Juel HANSEN Professeur des universités, University of Copenhagen - Department of Biology

Composition du Jury :

Rapporteure : Beatriz REGUERA Directrice de recherche, Instituto Español de Oceanografia - Centro Oceanográfico de Vigo Rapporteur : Per Juel HANSEN Professeur des universités, University of Copenhagen - Department of Biology

Président : Laurent BARILLÉ Professeur des universités, Université de Nantes - MMS Examinatrice : Claudia WIEGAND Professeure des universités, Université de Rennes 1 - CNRS - ECOBIO Directeur de thèse : Philipp HESS Docteur, Cadre de recherche, Laboratoire Phycotoxines, IFREMER Nantes Co-directrice de thèse : Hélène HÉGARET Docteure, Chargée de recherche, CNRS - LEMAR, Plouzané

Invités : Véronique SÉCHET Docteure, Cadre de recherche, Laboratoire Phycotoxines, IFREMER Nantes Damien RÉVEILLON Docteur, Cadre de recherche, Laboratoire Phycotoxines, IFREMER Nantes Wolfgang VOGELBEIN Directeur de recherche, Virginia Institute of Marine Science, USA

Acknowledgements

I first would like to gratefully acknowledge the members of my committee and the examiners of my PhD thesis, Beatriz Reguera, Per Juel Hansen, Claudia Wiegand and Laurent Barillé.

Ma thèse s’est déroulée au laboratoire Phycotoxines de l’IFREMER de Nantes, je remercie donc Zouher Amzil son directeur, et a été financée grâce au projet CoCliME.

Merci à mes deux directeurs de thèse, Philipp Hess et Hélène Hégaret ainsi qu’à mes deux co- encadrants, Véronique Séchet et Damien Réveillon, pour m’avoir permis de réaliser cette thèse dans les meilleures conditions possibles, pour m’avoir fait confiance, pour votre aide et vos encouragements. Pour ces raisons, et pour tout le reste je vous serai toujours reconnaissant.

I would like to extend my gratitude to Wolfgang Vogelbein for welcoming me to your laboratory in Virginia and for teaching me a few of your knowledge.

Je remercie les membres de mon comité de thèse, Rodolphe Lemée, Thomas Lacour, Manoella Sibat et Marc Sourisseau pour pour vos conseils et encouragements.

Merci à tous mes collègues de Phyc, Mano, Fabienne, Michelle, Véro, Liliane, Charline, Énora, Georges et Korian pour tous ces moments partagés, au travail et en dehors. Un grand merci à mes collaborateurs au labo, mais aussi à PBA, Aurélie, Thomas et Gaël, et au LEMAR à Brest, Nelly, Caroline, Myrina, Justine, Lucas, Marc, pour votre aide pendant les manips et pour la rédaction.

Merci Nour, Solène, Marin, Thomas, Seb, Florent, Élise, Estelle, Cyrielle, François et Max on aura bien rigolé ! Également, merci à ma famille et mes ami(e)s.

Table of contents

Acknowledgements ...... 3

Table of contents ...... 4

List of figures ...... 11

List of tables ...... 16

List of abbreviations ...... 21

Contexte et objectifs de la thèse ...... 24

Context and aims of the thesis ...... 34

Chapter 1: State of the art - Literature review ...... 43

I-1) General characteristics of the dinoflagellates ...... 44 I-2) Morphology and taxonomy of Dinophysis ...... 44 I-3) Dinophysis food chain ...... 47 I-3-1) First tier of the food chain: T. amphioxeia ...... 47 I-3-2) Mixotrophy of M. rubrum ...... 48 I-3-3) Mixotrophy of Dinophysis ...... 50 I-4) Culture of the three-tier food chain of Dinophysis...... 52 I-4-1) Culture of T. amphioxeia ...... 53 I-4-2) Culture of M. rubrum ...... 54 I-4-3) Culture of Dinophysis ...... 55 I-5) Ecophysiological experiments with Dinophysis species ...... 57 I-5-1) Temperature ...... 57 I-5-2) Irradiance ...... 59 I-5-3) Salinity ...... 62 I-5-4) Nutrients ...... 62 I-5-5) Geographical origin ...... 64 I-5-6) Prey ...... 65 I-5-7) Growth phase and cell cycle ...... 66 I-5-8) Conclusion of the ecophysiological experiments on Dinophysis ...... 67 I-6) Dinophysis Toxins ...... 68

I-6-1) OA-group toxins ...... 68 I-6-1-1) Chemical structure of OA-group toxins ...... 68 I-6-1-2) Toxicity of OA-group toxins by intraperitoneal injection and oral administration in rodents ...... 69 I-6-1-3) Elucidation of action mode of OA-group toxins...... 70 I-6-2) PTXs ...... 71 I-6-2-1) Chemical structure of PTXs ...... 71 I-6-2-2) Toxicity of PTXs by intraperitoneal injection and oral administration in rodents ...... 72 I-6-2-3) Elucidation of action mode of PTXs ...... 73 I-6-3) Effects of OA-group toxins and PTXs on humans ...... 74 I-6-4) OA-group toxin and PTX measurements ...... 74 I-6-4-1) Functional methods ...... 75 I-6-4-2) Immunological methods ...... 75 I-6-4-3) Chemical analytical methods ...... 75 I-7) Accumulation of Dinophysis, OA-group toxins and PTXs in marine organisms ...... 76 I-7-1) Accumulation in crustaceans, echinoderms, gastropods and cephalopods ...... 76 I-7-2) Accumulation in filter-feeding bivalves ...... 77 I-7-3) Accumulation in fish ...... 78 I-7-4) Accumulation in turtles ...... 78 I-7-5) Accumulation in birds ...... 78 I-7-6) Accumulation in marine mammals ...... 79 I-8) Effects of Dinophysis, OA-group toxins and PTXs on marine organisms ...... 79 I-8-1) Allelopathic effects ...... 80 I-8-2) Effects of filter-feeding bivalves exposure to Dinophysis or DSTs ...... 81 I-8-2-1) Exposure to Dinophysis ...... 81 I-8-2-2) Exposure to DSTs ...... 83 I-8-3) Effects of fish exposure to Dinophysis or DSTs ...... 85 I-8-3-1) Exposure to Dinophysis ...... 85 I-8-3-2) Exposure to DSTs ...... 86 I-9) Biology of the Pacific oyster C. gigas and fish sheepshead minnow C. variegatus ...... 88 I-9-1) C. gigas ...... 88 I-9-1) C. variegatus ...... 90 I-10) DSTs distribution ...... 91 I-10-1) Global distribution ...... 92 I-10-2) Regional distribution ...... 95

I-10-2-1) French case study ...... 95 I-10-2-2) Northern Europe ...... 100 I-10-2-3) Great Britain and Republic of Ireland ...... 102 I-10-2-4) Spain and Portugal ...... 104 I-10-2-5) Morocco, Tunisia, Italia, Croatia, Greece and Russia (Black Sea) ...... 106 I-10-2-6) Nigeria, Angole and South Africa ...... 108 I-10-2-7) North America ...... 109 I-10-2-8) Central and South America ...... 111 I-10-2-9) Japan and Republic of Korea ...... 113 I-10-2-10) East Asia ...... 115 I-10-2-11) India and the Arabian Sea ...... 117 I-10-2-12) South Asia and North Oceania ...... 118 I-10-2-13) Oceania ...... 118 I-10-3) Conclusion of the DSTs distribution ...... 121 Chapter 2: Ecophysiological experiments Part 1: Factorial design on Teleaulax and predator feeding ...... 122

II-1) Combined effects of temperature, irradiance and pH on Teleaulax amphioxeia (Cryptophyceae) physiology and feeding ratio for its predator Mesodinium rubrum (Ciliophoran) ...... 123 II-1-1) Graphical abstract ...... 123 II-1-2) Résumé ...... 124 II-1-3) Abstract ...... 125 II-1-4) Key words ...... 125 II-1-5) Introduction ...... 126 II-1-6) Materials and methods ...... 127 II-1-6-1) Full factorial design experiment on T. amphioxeia ...... 127 II-1-6-1-1) Culture of T. amphioxeia...... 127 II-1-6-1-2) Factorial design ...... 127 II-1-6-2) Effect of the prey on M. rubrum ...... 128 II-1-6-2-1) Photoacclimation of T. amphioxeia ...... 128 II-1-6-2-2) Feeding experiment ...... 129 II-1-6-3) Experimental set-up ...... 129 II-1-6-3-1) Counting and growth rate ...... 129

II-1-6-3-2) The maximum quantum yield of the photosystem II (Fv/Fm) ...... 130 II-1-6-3-3) Pigment analysis ...... 130

II-1-6-3-4) Statistical analyses ...... 130 II-1-7) Results...... 131 II-1-7-1) Direct effects, interactions and optimum of temperature, irradiance and pH on the physiology of T. amphioxeia ...... 131 II-1-7-1-1) Effect on the maximum growth rate ...... 131 II-1-7-1-2) Effect on the Fv/Fm and the pigment content ...... 132 II-1-7-2) Light acclimation of T. amphioxeia ...... 133 II-1-7-3) Feeding experiment ...... 134 II-1-8) Discussion ...... 136 II-1-8-1) Full factorial design experiment on T. amphioxeia and photoacclimation in semi- continuous culture ...... 136 II-1-8-2) Feeding experiment of M. rubrum ...... 139 II-1-9) Conclusion ...... 141 II-1-10) Acknowledgements ...... 141 II-1-11) Supplementary materials...... 142 Chapter 2: Ecophysiological experiments Part 2: Salinity stress on D. sacculus ...... 145

II-2) Effect of a 24 h salinity stress on the growth, biovolume, toxins, osmolytes and metabolite profiles on three strains of the Dinophysis acuminata-complex (“D. sacculus”) ...... 146 II-2-1) Graphical abstract ...... 146 II-2-2) Résumé ...... 147 II-2-3) Abstract ...... 148 II-2-4) Key words ...... 148 II-2-5) Introduction ...... 149 II-2-6) Materials and methods ...... 152 II-2-6-1) Culture maintenance ...... 152 II-2-6-2) Experimental design ...... 152 II-2-6-3) Growth rate calculation and biovolume measurement ...... 154 II-2-6-4) LC-LRMS/MS and LC-HRMS ...... 154 II-2-6-4-1) Toxin analysis ...... 155 II-2-6-4-2) Proline, GBT and DMSP analysis ...... 155 II-2-6-4-3) Metabolomics ...... 155 II-2-6-5) Statistical analyses ...... 156 II-2-7) Results...... 157 II-2-7-1) Growth rate and biovolume ...... 157 II-2-7-2) Toxins ...... 157

II-2-7-3) Proline, GBT and DMSP ...... 159 II-2-7-4) Metabolomics ...... 160 II-2-8) Discussion ...... 161 II-2-9) Acknowledgements ...... 165 II-2-10) Supplementary materials...... 166 Chapter 3: Effects on marine organisms Part 1: Dinophysis and its toxins towards oyster gametes ...... 170

III-1) Cultures of Dinophysis sacculus, D. acuminata, and pectenotoxin 2 affect gametes and fertilization success of the Pacific oyster, Crassostrea gigas ...... 171 III-1-1) Graphical abstract ...... 171 III-1-2) Résumé ...... 172 III-1-3) Abstract ...... 173 III-1-4) Key words ...... 173 III-1-5) Introduction ...... 174 III-1-6) Materials and methods ...... 176 III-1-6-1) Microalgal cultures...... 176 III-1-6-2) Experimental design ...... 176 III-1-6-2-1) Experiment 1 (Exp. 1) - Effect of whole culture of D. sacculus upon gametes ..... 177 III-1-6-2-2) Experiment 2 (Exp. 2) - Effect of resuspended cells and culture filtrate of D. sacculus upon gametes ...... 177 III-1-6-2-3) Experiment 3 (Exp. 3) - Effect the whole culture of D. acuminata upon gametes 178 III-1-6-2-4) Experiment 4 (Exp. 4) - Effect of OA PTX2 standards on gametes ...... 178 III-1-6-3) Toxin analyses of Dinophysis spp. strains ...... 178 III-1-6-4) Sampling and maintenance of oysters ...... 179 III-1-6-5) Collection of gametes ...... 179 III-1-6-6) Flow-cytometry analysis - morphology, viability and ROS production ...... 180 III-1-6-7) Microscopy analysis - fertilization success assessment ...... 180 III-1-6-8) Statistial analysis ...... 181 III-1-7) Results ...... 181 III-1-7-1) Toxin contents of Dinophysis spp. cultures ...... 181 III-1-7-2) Exp. 1 - Effect of the whole culture of D. sacculus on gametes ...... 183 III-1-7-3) Exp. 2 - Effect of resuspended cells and culture filtrate of D. sacculus on gametes .... 185 III-1-7-4) Exp. 3 - Effect the whole culture of D. acuminata on gametes ...... 186 III-1-7-5) Exp. 4 - Effect of OA and PTX2 standards on gametes ...... 187 III-1-8) Discussion ...... 188

III-1-9) Conclusion ...... 193 III-1-10) Acknowledgements ...... 194 III-1-11) Supplementary materials ...... 195 Chapter 3: Effects on marine organisms Part 2: D. acuminata and PTX2 towards fish ...... 200

III-2) Mortality and histopathological features of sheepshead minnow (Cyprinodon variegatus) larvae exposed to Dinophysis acuminata and pectenotoxin 2 during a 96 h bioassay ...... 201 III-2-1) Graphical abstract ...... 201 III-2-2) Résumé ...... 202 III-2-3) Abstract ...... 203 III-2-4) Key words ...... 204 III-2-5) Introduction ...... 204 III-2-6) Materials and methods ...... 207 III-2-6-1) Microalgal cultures...... 207 III-2-6-2) Fish larvae ...... 208 III-2-6-3) Experimental design ...... 208 III-2-6-3-1) 96 h larval fish bioassay...... 208 III-2-6-3-2) Exposure of D. acuminata to C. variegatus ...... 208 III-2-6-3-3) Exposure of PTX2 to C. variegatus ...... 209 III-2-6-4) Histology: light microscopy and TEM techniques ...... 209 III-2-6-5) Toxin analyses ...... 210 III-2-6-6) Statistical analyses ...... 211 III-2-7) Results ...... 211 III-2-7-1) Toxins of D. acuminata...... 211 III-2-7-2) Cumulative mortalities ...... 212 III-2-7-3) Histology ...... 214 III-2-8) Discussion ...... 216 III-2-9) Acknowledgements ...... 220 III-2-10) Supplementary materials ...... 221 General discussion ...... 222

The effect of key factors of global change on T. amphioxeia ...... 223 The effect of contrasting physiological status and culture density of the prey, T. amphioxeia, on its predator M. rubrum ...... 224 Dinophysis in future climate conditions ...... 225 The osmoregulation process in D. sacculus ...... 229

Intraspecific variability in D. sacculus ...... 230 Roles and implications of the OA-group toxins and PTXs ...... 231 Effects of Dinophysis and its toxins on marine organisms ...... 234 Environmental risks posed by PTX2-producers ...... 239 Conclusion ...... 242

Scientific publications and communications ...... 245

Appendices ...... 247

References ...... 260

List of figures

Figure 1: Scanning electron microscopy image of (A) D. acuta (holotype) strain IFR-DAC-03Ke. Parameters H, height; L, large diameter; s, small diameter are dimensions measured to estimate equivalent cellular biovolumes according to Olenina et al. (2006). (B) D. caudata strain IFR-DCA- 04Tr, (C) D. tripos strain IFR-DTR-01Ar, (D) D. acuminata strain IFR-DAU-01Es and (E) D. sacculus strain IFR-DSA-01Lt modified from Séchet et al., 2020 (pictured by N. Chomérat), and (F) D. ovum from Gulf of Mexico modified from Wolny et al.,. (2020). (D), (E) and (F) belong to the D. acuminata-complex. Scale bar: 10 µm ...... 47

Figure 2: Acquisition of the cryptophyte plastids in M. rubrum and Dinophysis. From Wisecaver and Hackett (2010) ...... 50

Figure 3: Chemical structure of OA and its analogues. Modified from Boundy et al. (2020) ...... 69

Figure 4: Chemical structure of PTXs and their analogues. Modified from Boundy et al. (2020) 72

Figure 5: Life cycle of the Pacific oyster, Crassostrea gigas, modified from (Vogeler et al., 2016). hpf, hours post fertilization; dpf, days post fertilization; mpf, months post fertilization ...... 90

Figure 6: Global distribution of DSTs (black dots: n = 461) combining reports of toxins in Dinophysis species (in vivo and in vitro) (n = 233), marine animals (n = 209) and SPATT (n = 19)...... 93

Figure 7: Global distribution of DSTs measurements (white dots) in SPATT (n = 19) ...... 94

Figure 8: France. Distribution of DST measurements in Dinophysis species in Northern Bay of Biscay: A. Brittany and B. Vilaine and Loire Estuaries; C. Southern Bay of Biscay; D. English Channel and E. French Mediterranean coastal waters, and F. in marine animals ...... 97

Figure 9: Northern Europe. Distribution of DST measurements A. in Dinophysis species and B. marine animals, and C. in marine animals in Western Russia ...... 101

Figure 10: Great Britain and Republic of Ireland. Distribution of DST measurements A. in Dinophysis species and B. in marine animals, and C. in marine animals in Southern Scotland . 104

Figure 11: Spain and Portugal. Distribution of DST measurements A. and B. in Dinophysis species in Western and Southern Iberia and Western Spain, respectively and C. in marine animals ...... 105

Figure 12: Morocco, Tunisia, Italia, Croatia, Greece and Russia (Black Sea). Distribution of DST measurements A. in Dinophysis species in Italia and B. in marine animals from North Africa to the Russian Black Sea coastal waters...... 107

Figure 13: Nigeria, Angola and South Africa. Distribution of DST measurements A. in marine animals in Central-West Africa and B. and C. in Dinophysis species and marine animals, respectively, in South Africa ...... 108

Figure 14: USA and Canada. Distribution of DST measurements A. in marine animals in USA and Canada and in Dinophysis species in B. West USA, C. North East USA and South East Canada and D. South USA (Gulf of Mexico) ...... 110

Figure 15: Central and South America. Distribution of DST measurements in marine animals in A. North West, B. North East, C. South and D. South East South America, and in Dinophysis species for the same locations respectively: E., F., G. and H ...... 112

Figure 16: Japan and Republic of Korea. Distribution of DST measurements in Dinophysis species in A. North (Hokkaido), B. North (Aomori and Akita), C. North East, D. North West of Japan and E. South West of Japan and Southern Republic of Korea ...... 113

Figure 17: Japan and Republic of Korea. Distribution of DST measurements in marine animals in A. Southern Republic of Korea and B. North East of Japan ...... 115

Figure 18: East Asia. Distribution of DST measurements A. in Dinophysis species and B. in marine animals in China and Hong Kong ...... 116

Figure 19: West Asia. Distribution of DST measurements in Dinophysis species and marine animals in India and Qatar ...... 117

Figure 20: South Asia and North Oceania. Distribution of DST measurements in Dinophysis species in Republic of Singapore and Indonesia ...... 118

Figure 21: Oceania. Distribution of DST measurements in Dinophysis species in A. Australia and B. New Zealand, and in marine animals respectively for the same locations C. and D ...... 120

Figure 22: (A) Standard Pareto chart the model of the maximum growth rate. Linear and quadratic effects of factors on growth are represented by single or double parameters, respectively. (B) Direct effect of T, pH and I on growth rate of T. amphioxeia. (C) Interaction plots of growth rate; + and - correspond to the maximum and minimum values of the second factor. (D) Surface plot of the

modeled growth rate. T = temperature (°C) and I = irradiance (µmol photons · m-2 · s-1). Significant effects are marked with an asterisk ...... 132

Figure 23: Standard Pareto charts for (A) the model of maximum quantum yield of the photosystem II, (B) total chlorophyll a and (C) chlorophyll c. Linear and quadratic effects of factors on growth are represented by single or double parameters, respectively. T = temperature (°C) and I = irradiance (µmol photons · m-2 · s-1) ...... 142

Figure 24: Box plot of the sea water salinity at the three locations where the studied strains of “D. sacculus” were isolated ...... 154

Figure 25: Total (sum of intracellular and extracellular in pg cell-1) of A. okadaic acid (OA) and B. pectenotoxin 2 eq (PTX2eq) in the three strains of “D. sacculus” after 24 h of salinity stresses at 25 and 42 and for the control. Values are expressed as mean ± SD (n = 3). Treatments with asterisks were significantly different ...... 158

Figure 27: Principal component analysis (PCA) score plot obtained from LC-HRMS profiles of the three “D. sacculus” strains exposed to the three salinities (S) ...... 161

Figure 28: A. Adenosine (feature M268T51) and B. eicosapentaenoic acid (EPA) (feature M301T600) areas at the three salinities for the “D. sacculus” strain from Thau lagoon (Dsa-Th)...... 169

Figure 29: Effect of a gradient of concentration of D. sacculus on gamete fertilization (Exp. 1): Mean percentage of fertilized eggs with A. oocytes exposed, B. spermatozoa exposed, C. both gametes exposed and D. fertilization in presence of D. sacculus (without gametes exposure) from 0.5 to 500 cell mL-1 and to a sea water control. Values are expressed as mean ± SD (n = 8 - 11). Treatments with different superscript letters were significantly different...... 185

Figure 30: Effect of whole culture (500 cells mL-1), resuspended cells (500 cells mL-1) and culture filtrate (eq 500 cells mL-1) of D. sacculus on gamete fertilization (Exp. 2). Mean percentage of fertilized eggs (%) with A. oocytes exposed and B. spermatozoa exposed. Values are expressed as mean ± SD (n = 5). Treatments with different superscript letters were significantly different. .. 186

Figure 31: Mean percentage of fertilized eggs (%, Exp. 3) with exposure of both oocytes and spermatozoa to a gradient of concentration of D. acuminata from 0.5 to 500 cell mL-1 and to a sea water control. Values are expressed as mean ± SD (n = 4). Treatments with different superscript letter were significantly different...... 187

Figure 32: Effect a gradient of concentration of okadaic acid (OA) or Pectenotoxin 2 (PTX2) on gamete fertilization (Exp. 4): Mean percentage of fertilized eggs (%) with A. oocytes exposed, B spermatozoa exposed, C. both gametes exposed and D. fertilization in presence of OA or PTX2 (without gametes exposure) from 5 to 50 nM, including sea water and methanol controls (MeOH). Values are expressed as mean ± SD (n = 3 - 4). Treatments with different superscript letters were significantly different...... 188

Figure 33: Experimental design Chapter 3-Part 1 ...... 196

Figure 35: Cumulative mortality (%) over 96 h in C. variegatus larvae exposed to 50, 250, 500, 750, 1000, 2000, 4000 nM of pectenotoxin-2 (PTX2), filtered sea water and ethanol (0.8 % final) controls (n = 38 for each condition). *significantly different (Pearson’s Chi-square test and post hoc test on residuals) ...... 213

epithelium. Some evidence of cell necrosis (X60). E. Gill lamellae of fish exposed to 5500 cells mL-1 of D. acuminata culture sampled at 96 h and exhibiting largely intact, presumably functional lamellar tissue architecture. (G1) Primary gill filament (X40). F. Gill lamellae of fish exposed to 5500 cells mL-1 of D. acuminata culture sampled at 96 h. Partial occlusion of interlamellar spaces by hypertrophied and necrotic chloride cells (Cl) (X20)...... 216

Figure 37: Effect of abiotic factors of climate change (temperature, irradiance, pH, salinity and nitrogen) and biotic factors (prey concentration, competition) on the growth of the three organisms of the Dinophysis food chain. Blue arrows represent factors investigated in this thesis (Chapter 2- Part 1 and Chapter 2-Part 2) and grey arrows represent abiotic factors tested in the literature. Dotted arrows indicate putative effects. Arrowheads with “+” mean growth enhancement. Factors preceded by “±” mean tolerance to increase and decrease. The arrows size is not informative. . 227

List of tables

Table 1: Culture conditions for the main cultivated strains of T. amphioxeia from 2006 to 2020 53

Table 2: Culture conditions for the main cultivated strains of M. rubrum from 2006 to 2020 ...... 54

Table 3: Example of culture conditions used for D. acuminata and D. acuta from 2006 to 2020 56

Table 4: In vitro effect of temperature on Dinophysis growth rate, cellular concentration and toxin contents ...... 58

Table 5: In vitro effect of irradiance (intensity and quality) on Dinophysis growth rate, cellular concentration, cellular toxin content and photosynthetic parameters...... 60

Table 6: In vitro effect of salinity on Dinophysis growth rate ...... 62

Table 7: In vitro effect of nutrients on Dinophysis growth rate and toxin contents...... 63

Table 8: Effects of geographical origin of Dinophysis and/or its food chain on growth rate and toxin contents ...... 64

Table 9: In vitro effect of prey type and concentration on Dinophysis growth rate and toxin content...... 65

Table 10: Effects of the population growth phase and cell cycle stages on Dinophysis growth rate and toxin content ...... 67

Table 11: Effect of mollusks exposure to Dinophysis ...... 82

Table 12: Effect of mollusks exposure to Dinophysis toxins...... 84

Table 13: Effect of fish exposure to Dinophysis ...... 85

Table 14: Effect of fish exposure to Dinophysis toxins ...... 87

Table 15: Review of global distribution of Dinophysis shellfish toxin (DST) measurements in marine animals, in Dinophysis (culture and field sampling) and in solid phase adsorption toxin tracking (SPATT), available online at: https://doi.org/10.17882/76433 ...... 92

Table 16: Literature review of French DST measurements in marine animals and in field populations of Dinophysis species ...... 97

Table 18: Factor levels in the factorial design experiment, where α is the axial distance between star points and the center of the experimental domain ...... 128

Table 19: Comparison between observed values of maximum growth rate (µmax, d-1), maximum -1 quantum yield of the photosystem 2 (Fv/Fm), total chlorophyll a (TChl a, pg · cell ), chlorophyll c (Chl c, pg · cell-1) and total carotenoids (TCarotenoids, pg · cell-1) of T. amphioxeia acclimated to low light (LL) and high light (HL) conditions, and modeled values (marked with th) according to the factorial design experiment. Values are expressed as mean ± SD ...... 134

Table 20: Comparison between maximum growth rate (µmax, d-1), maximum cellular -1 concentration (cells · mL ), maximum quantum yield of the photosystem 2 (Fv/Fm), chlorophyll a total (TChl a, pg · cell-1), chlorophyll c (Chl c, pg · cell-1) and carotenoids total (TCarotenoids, pg · cell-1) of M. rubrum fed at different prey: predator ratios; high fed and low fed of T. amphioxeia itself acclimated to low light (LL) and high light (HL) conditions, and M. rubrum and T. amphioxeia controls. Values are expressed as ± SD. No significant differences were found when LL and HL feeding were compared two by two among each nutrition ratio...... 135

Table 21: Culture conditions of strains used in this study ...... 143

Table 22: Regression coefficients for the models of maximum growth rate (µmax th), maximum quantum yield of the photosystem II (Fv/Fm th), total chlorophyll a (TChl a th) and chlorophyll c

(Chl c th), where β0 is the model error, 1 is for temperature, 2 for pH and 3 for irradiance ...... 143

Table 23: Maximum growth rate (µmax), maximum quantum yield of the photosystem II (Fv/Fm), total chlorophyll a (TChl a), chlorophyll c (Chl c th) and total carotenoids (TCarotenoids) for the different conditions in the factorial design experiment ...... 144

Table 24: Growth rate (µ, d-1) and biovolume (x103 µm3) of the 3 strains of “D. sacculus” after 24 h of salinity stresses at 25 and 42 and for the control ...... 157

Table 25: Origin and culture conditions of the strains used in this study ...... 166

Table 26: Proportion of intracellular okadaic acid (OA intra) and pectenotoxin 2 eq (PTX2eq intra) contents for the three strains of “D. sacculus” after 24 h of salinity stresses at 25 and 42 and for the control ...... 166

Table 27: Lists (m/z, retention time (rt) and acquisition mode) of the significantly different features obtained from LC-HRMS profiles of the three “D. sacculus” strains A. Dsa-Me, B. Dsa-Lo and C. Dsa-Th exposed to the three salinities. Only two replicates were available for Dsa-Th S25 and Dsa- Th S35...... 167

Table 28: Mean intracellular (intra, pg cell-1), extracellular (extra, eq pg cell-1), total (sum of intracellular and extracellular, pg cell-1 and nM corresponding to 500 cells mL-1) of okadaic acid (OA) and pectenotoxin 2 eq (PTX2eq) for D. sacculus (Exp. 1 and Exp. 2) and D. acuminata (Exp. 3). Values are expressed as mean ± SD (n = 4 for Exp.1 and Exp. 2 and n = 1 for Exp. 3). Treatments with different superscript letter were significantly different and absence of superscript letter means NS difference...... 182

Table 29: Forward scatter (FSC), side scatter (SSC), reactive oxygen species (ROS) production (a.u., % of control) and mortality of oocytes and spermatozoa after exposure to a gradient of concentration from 0.5 to 500 cells mL-1 of D. sacculus (Exp. 1) or D. acuminata (Exp. 3) and to a gradient of concentration from 5 to 50 nM of okadaic acid (OA) or pectenotoxin 2 (PTX2) (Exp. 4) and to sea water and methanol (MeOH) controls. Values are expressed as mean ± SD (n = 4 - 5 for Exp 1, n = 3 - 4 for Exp. 3 and n = 5 for Exp. 4). Treatments with different superscript letter were significantly different. n.a. data not available ...... 183

Table 30: Culture conditions of strains used in this study ...... 195

Table 31: Mean total (intracellular and extracellular) concentration of okadaic acid (OA), dinophysistoxin 1 (DTX1) and pectenotoxin 2 (PTX2) in pg cell-1 and nM, corresponding to 5500 cell mL-1 (i.e. maximal concentration of D. acuminata used in this study) in D. acuminata strain DAVA01. Note that DAVA01 is largely a PTX2 producer. Values are mean ± SD, n = 3 ...... 212

-1 Table 32: LC50 (nM and ng mL ) of pectenotoxin-2 (PTX2) based on the Hill coefficient according to exposition time ...... 214

Table 33: Origins and culture conditions for the three organisms required to actively culture Dinophysis acuminata (strain DAVA01) used in this study ...... 221

Table 34: Cultivated strains of T. amphioxeia (and other TPG species) from 2006 to 2020: temperature (T °C), irradiance (I µmol photons m-2 s-1), pH, salinity and light: dark cycle (L: D). n.a. means that the information is not available...... 247

Table 35: Cultivated strains of M. rubrum from 2006 to 2020: temperature (T °C), irradiance (I µmol photons m-2 s-1), pH, salinity, light: dark cycle (L: D), predator: prey ratio and nutrition frequency. n.a. means that the information is not available...... 250

Table 36: Cultivated strains of D. acuminata from 2006 to 2020: temperature (T °C), irradiance (I µmol photons m-2 s-1), pH, salinity, light: dark cycle (L: D), predator prey ratio and nutrition frequency. n.a. means that the information is not available...... 253

Table 37: Cultivated strains of D. acuta from 2006 to 2020: temperature (T °C), irradiance (I µmol photons m-2 s-1), pH, salinity, light: dark cycle (L: D), predator: prey ratio and nutrition frequency. n.a. means that the information is not available...... 255

Table 38: Cultivated strains of Dinophysis sacculus from 2006 to 2020: temperature (T °C), irradiance (I µmol photons m-2 s-1), pH, salinity, light: dark cycle (L: D), predator prey ratio and nutrition frequency. n.a. means that the information is not available...... 256

Table 39: Cultivated strains of D. caudata, D. fortii, D. infundibulus, D. ovum and D. tripos from 2006 to 2020: temperature (T °C), irradiance (I µmol photons m-2 s-1), pH, salinity, light: dark cycle (L: D), predator prey ratio and nutrition frequency. n.a. means that the information is not available...... 257

List of abbreviations

DCFH-DA 2′7’-dichlorofluorescein FSW filtered sea water diacetate FSSW filter-sterilized sea water 7-epi-PTX2-sa 7-epi-pectenotoxin 2 seco acid FCM flow cytometry

ASP amnesic shellfish poisoning FSC forward scatter ACD at-column dilution IFREMER French Research Institute for Exploitation of the Sea AZP azaspiracid shellfish poisoning GBT glycine betaine BW body weight µ growth rate Chl c chlorophyll c HA harmful algae CP ciguatera poisoning HAB harmful algal bloom CoCliME Co-development of Climate HL high light services for adaptation to changing Marine Ecosystems HILIC Hydrophilic interaction liquid dpf days post fertilization chromatography DSP diarrheic shellfish poisoning hpf hours post fertilization

DST diarrheic shellfish toxin intra intracellular DMSP dimethylsulfoniopropionate I irradiance D Dinophysis L: D light: dark

DTXs dinophysistoxins LC liquid chromatography DIN Dissolved inorganic nitrogen LC-FLD liquid chromatography coupled to fluorescence detector DON Dissolved organic nitrogen LC-HRMS liquid chromatography EPA eicosapentaenoic acid coupled to high resolution mass spectrometry ESI electrospray ionization LC-LRMS/MS liquid chromatography ELISA enzyme-linked immunosorbent coupled to low resolution tandem mass assay spectrometry Caco-2 epithelial colorectal LC-MS liquid chromatography coupled adenocarcinoma cells to mass spectrometry EtOH ethanol LC-MS/MS liquid chromatography extra extracellular coupled to tandem mass spectrometry

LC-UV liquid chromatography coupled QTOF quadrupole-time of flight mass to UV detector spectrometer MS mass spectrometry RBA rat bioassay µmax maximum growth rate ROS reactive oxygen species Fv/Fm maximum quantum yield of the Ref. Reference photosystem II rETR Relative electron transport rate M Mesodinium SSC side scatter REPHYTOX Monitoring Network for Phycotoxins in marine organisms) SPATT solid phase adsorption toxin tracking mpf months post fertilization SPE solid phase extraction MBA mouse bioassay TPG Teleaulax/Plagioselmis/ NSP neurotoxic shellfish poisoning Geminifera NPQ Non photochemical quenching T temperature REPHY Observation and Surveillance Network for Phytoplankton and Tcarotenoids total carotenoids Hydrology in coastal waters TChl a total chlorophyll a OA okadaic acid TEM transmission electron microscopy PSP paralytic shellfish poisoning UHPLC-LRMS/MS ultra-high PTX2eq pectenotoxin 2 equivalent performance liquid chromatography coupled to low resolution tandem mass PTX2-sa pectenotoxin 2-seco acid spectrometry PTXs pectenotoxins

ΦPSII Photochemical yield of the photosystem II

PUFA polyunsaturated fatty acid

QC pool samples PI propidium iodide PP1 protein phosphatase 1

PP2A protein phosphatase 2A

PP protein phosphatases

Contexte et objectifs de la thèse

Par définition, le plancton est un organisme qui est adapté pour vivre en suspension dans l'eau de mer, les rivières, les lacs et les étangs (Reynolds, 2006). Le plancton regroupe des organismes vivants de la taille des virus (nanomètres) aux méduses (mètres), qui ne sont pas capables de contrôler leur position horizontale ou de nager à contre-courant (plancton dérivé du mot grec pour errant, vagabond). Parmi eux, les plantes unicellulaires - le phytoplancton - représentent les microorganismes photosynthétiques. Les organismes phytoplanctoniques sont à la base de la chaîne alimentaire aquatique et sont l'un des principaux producteurs primaires de carbone organique dans l’eau (Falkowski et Raven, 1997; Reynolds, 2006). En effet, grâce au phytoplancton, la majorité de la photosynthèse sur Terre se produit en milieu aquatique, où la consommation de dioxyde de carbone de l'atmosphère équivaut à celle des forêts et des autres plantes terrestres, et participe ainsi à la régulation du réchauffement climatique (Field et al., 1998). Les organismes phytoplanctoniques peuvent être classés en cinq groupes en fonction de la structure de leurs plastes (c.-à-d. des organites entourés de membranes), des arrangements membranaires et des pigments : les algues vertes, les algues bleues, les coccolithophores, les diatomées et les dinoflagellés (Falkowski et al., 2004; Pachiappan et al., 2019).

Certains organismes phytoplanctoniques peuvent être appelés algues nuisibles lorsqu'ils provoquent des effets délétères allant de simples nuisances, tel que le salissement des plages par échouage et l'anoxie, aux effets toxiques sur les organismes aquatiques tels que l'irritation cutanée ou la suffocation des poissons ainsi que l'intoxication alimentaire des humains causée par la consommation de crustacés, de poissons ou de mammifères par bioaccumulation des toxines dans les réseaux trophiques (Granéli et Turner, 2006; Smayda, 1997a). Ces phénomènes peuvent être renforcés lors des efflorescences (c.-à-d. des augmentations importantes et rapides de la population par rapport à la concentration classique) d'algues nuisibles ou HABs pour « harmful algal blooms » en anglais.

Une prolifération de phytoplanctons peut être déclenchée ou inhibée en fonction de facteurs à la fois biotiques (par exemple la nutrition, la compétition et le broutage) et/ou abiotiques (par exemple

Contexte et objectifs l’hydrologie et la météorologie). Pour le phytoplancton producteur de toxines, une prolifération peut également être définie comme une augmentation de la production de phycotoxines (c.-à-d. des toxines synthétisées par le phytoplancton), qui peuvent varier dans le temps, la durée et l'emplacement (Smayda, 1997a). Parmi les milliers d'espèces de phytoplancton marin décrites (Reynolds, 2006), environ 300 peuvent former des marées rouges (c.-à-d. une décoloration de l'eau, qui peut être nocive) et 141 sont capables de synthétiser des toxines (Moestrup et al., 2009). Parmi les producteurs de toxines, 50 à 75% sont des dinoflagellés (Hallegraeff, 1993; Moestrup et al., 2009; Smayda, 1997b).

Les toxines algales sont de petits métabolites, avec une diversité de composés, de structures chimiques et de voies de toxicité. Les toxines ont été classées en fonction de leurs effets toxiques et de leurs syndromes chez l'homme après la consommation de bivalves filtreurs contaminés, comme les huîtres, les moules ou les palourdes et des poissons. Les principales intoxications sont l'intoxication diarrhéique par les mollusques (DSP pour « diarrheic shellfish poisoning » en anglais), l'intoxication paralysante par les mollusques (PSP pour « paralytic shellfish poisoning » en anglais), l'intoxication par la ciguatera (CP pour « ciguatera poisoning » en anglais), l'intoxication amnésique par les mollusques (ASP pour « amnesic shellfish poisoning » en anglais), l'intoxication neurotoxique par les mollusques (NSP pour « neurotoxic shellfish poisoning » en anglais) ou l'intoxication par azaspiracide (AZP pour « azaspiracid shellfish poisoning ») (Hallegraeff, 2004; Lassus et al., 2016; Lawrence et al., 2011; Rossini et Hess, 2010; Young et al., 2020). Outre les effets sanitaires, des problèmes économiques et sociétaux se posent également fréquemment en raison des HABs (Glibert et al., 2014; Reguera et al., 2012). Par conséquent, et en raison de l'augmentation des intoxications humaines enregistrées par les phycotoxines dans la deuxième partie du siècle dernier (Hallegraeff, 2010; Lassus et al., 2016), les autorités ont mis en œuvre des programmes de détection et de surveillance des espèces de phytoplanctons toxiques et des bivalves filtreurs contaminés. Des seuils sanitaires spécifiques (de phytoplancton et/ou de phycotoxines) ont été promulgués selon plusieurs directives (par exemple des directives européennes, japonaises et américaines ; European Commissions 2011, DeGrasse et Martinez-Diaz 2012, Hess 2012, Suzuki et Watanabe 2012, NSSP 2017). Au-delà de ces seuils, les autorités (par exemple les préfectures en France) peuvent interdire la récolte des coquillages et les activités associées (y compris le transport, la vente et la distribution), ainsi que la récolte récréative pour des périodes allant de quelques jours à plusieurs mois (Belin et al., 2020). Les pertes économiques de

Contexte et objectifs ces interdictions sur les fermetures de fermes aquacoles et les effets sociétaux qui en découlent (par exemple la perte d'activité ou le chômage) ont atteint plus d'un milliard d’USD en 2006 en Europe et aux États-Unis, et pourraient même être plus élevées de nos jours (Granéli et Turner, 2006; Hoagland et al., 2020; Hoagland et Scatasta, 2006).

En France, le réseau de surveillance REPHY-REPHYTOX (appelé REPHY dans le manuscrit) a été créé en 1984 par l'Institut français de recherche pour l'exploitation de la mer (IFREMER). Ce réseau vise à acquérir des données sur les paramètres hydrologiques et sur l'abondance et la composition du phytoplancton ainsi qu'à surveiller les phycotoxines chez les bivalves filtreurs (Belin et al., 2020; REPHY, 2019). Des diatomées toxiques du genre Pseudo-nitzschia et des dinoflagellés des genres Alexandrium et Dinophysis et leurs toxines associées se retrouvent régulièrement dans l'eau et dans les coquillages des côtes de l'Atlantique, de la Méditerranée et de la Manche (Belin et al., 2020). Le premier événement d'intoxication répertorié en France a été enregistré en 1983 et était dû à l'accumulation de toxines DSP dans les moules provenant d'espèces de dinoflagellés toxiques du genre Dinophysis (Ehrenberg, 1841; Lassus et al., 1985), avec des symptômes d’une intoxication gastro-intestinale chez l'homme, comme des diarrhées, des nausées, des vomissements et des douleurs abdominales. Depuis cette épidémie qui a conduit à la mise en place du REPHY en 1984, Dinophysis est le genre le plus emblématique en France. En effet, des cellules de ce genre de phytoplancton et ces toxines se retrouvent chaque année dans l'eau et les mollusques et sont principalement responsables des interdictions, malgré des concentrations cellulaires généralement faibles (≤ 100 cellules L-1; Belin et al., 2020).

Plusieurs espèces du genre Dinophysis sont reconnues comme principalement responsables de DSP (Reguera et al., 2014). Parmi les 133 espèces du genre Dinophysis acceptées taxonomiquement (Gómez, 2005; Jensen et Daugbjerg, 2009), dix espèces de Dinophysis ont été liées à des évènements de DSP et/ou caractérisées comme productrices de toxines diarrhéiques par les mollusques (DST pour « diarrheic shellfish toxins » en anglais ) ainsi que deux espèces du genre Phalacroma (Nagai et al., 2020): D. acuminata (Claparède & Lachmann 1859), D. acuta (Ehrenberg, 1839), D. caudata (Saville Kent, 1881), D. fortii (Pavillard, 1923), D. infundibulum (Schiller, 1928), D. miles (Cleve, 1900), D. norvegica (Claparède and Lachmann, 1859) D. ovum (Schütt, 1895), D. sacculus (Stein 1883) et D. tripos (Gourret, 1883), et Phalacroma mitra (Schütt,

Contexte et objectifs

1895) (= D. mitra; Abé, 1967) et Phalacroma rotundatum (Kofoid and Michener, 1911) (= D. rotundata; Claparède and Lachmann, 1859).

Ces espèces toxiques produisent deux types de phycotoxines lipophiles, à savoir l'acide okadaïque (OA pour « okadaic acid » en anglais) qui est un polyéther et ces dérivés, les dinophysistoxines (DTXs pour « dinophysistoxins » en anglais), et les pecténotoxines (PTXs pour « pectenotoxins » en anglais qui sont des lactones macrocycliques. L’OA et les DTXs sont des toxines diarrhéiques, tandis que les PTXs, également souvent trouvées dans les coquillages en raison de la coproduction par Dinophysis, ne provoquent pas de symptômes diarrhéiques (Cohen et al., 1990; Reguera et Pizarro, 2008). Il faut noter que si la toxicité de l’OA, des DTXs et des PTXs a été bien décrite sur des modèles de rongeurs, leurs effets (et ceux dérivés directement des cellules de Dinophysis) sur les organismes marins, comme les mollusques ou les poissons qui sont régulièrement contaminés, et en particulier sur les premiers stades de vie qui sont des phases de développement critiques, sont encore mal connus.

Les contaminations DSP et les DST associées sont répertoriées dans le monde entier depuis la première intoxication massive survenue aux Pays-Bas en 1961 (Korringa et Roskam, 1961). Durant les 45 années suivantes, les données scientifiques ont été obtenues uniquement à partir d'études sur le terrain et/ou de cellules de plancton prélevées in situ (par exemple Yasumoto et al. (1978), Kumagai et al. (1986), Cembella (1989), Masselin et al. (1992), Vale et Sampayo (2002), Luisa Fernández et al. (2006)). La première tentative de culture de Dinophysis, dans des milieux de culture « classiques » (c. -à-d. de l’eau de mer enrichie en nutriments, en éléments trace métalliques et en vitamines) pour des organismes autotrophes date de 1935 (Reguera et al., 2012), mais fût sans succès. Après cela, plusieurs tentatives de culture de Dinophysis avec des proies (pico- nanoplancton) ou des bactéries ont également échouées (Ishimaru et al., 1988; Maestrini et al., 1995). Ces expériences infructueuses étaient en fait dues au mode de nutrition de Dinophysis.

Les protistes présentent différents types d'interactions trophiques, comme la mixotrophie, qui est la combinaison de la phototrophie et de l'hétérotrophie. Le mode de nutrition mixotrophique permet aux protistes d'obtenir de l'énergie, du carbone et des nutriments à partir de la photosynthèse et également à partir du carbone organique dissous et/ou de la phagotrophie (= photo-phago- mixotroph; Burkholder et al., 2008; Hansen et al., 2019). La mixotrophie est répandue dans plusieurs taxons de phytoplancton, tels que les radiolaires, les foraminifères, les ciliés et les

Contexte et objectifs dinoflagellés (Hansen et al., 2019; Stoecker, 1998). Les observations microscopiques traditionnelles suggéraient le mode de nutrition mixotrophique de Dinophysis. En effet, les plastes de Dinophysis ont été décrits comme dérivant de cryptophytes (Schnepf et Elbrachter, 1988), et des vacuoles alimentaires et des pédoncules alimentaires ont été observés au sein d'espèces toxiques du genre (Hansen, 1991; Jacobson et Andersen, 1994). L'origine cryptophytique des plastes a ensuite été prouvée par des analyses moléculaires (Janson et Granéli, 2003). Le cryptophyte Teleaulax amphioxeia (Conrad) Hill (1992) a été identifié comme le donneur des plastes (Janson, 2004) grâce à un vecteur cilié, à savoir Mesodinium rubrum (Lohmann 1908) (Yih et al., 2004) par un processus de séquestration des plastes, c.-à-d. la kleptoplastidie (Schnepf et Elbrächter, 1999; Takishita et al., 2002). Les souches polaires de M. rubrum quant à elles se nourrissent en culture sur un autre cryptophyte, Geminigera cryophila (Johnson et al., 2007).

La première culture permanente d'une espèce de Dinophysis en laboratoire a été établie par Park et al. (2006) et était basée sur une chaîne trophique à trois membres : Teleaulax sp. en bas de la chaîne, Mesodinium rubrum et D. acuminata comme prédateur supérieur. Depuis 2006, plusieurs cultures d'espèces toxiques de Dinophysis ont été établies, permettant ainsi aux chercheurs de mener des expériences écophysiologiques in vitro. Ces études ont donné un aperçu de l'impact des facteurs biotiques ou abiotiques sur par exemple la croissance et la teneur en toxines de Dinophysis, en particulier dans un contexte de changement global.

Des préoccupations ont été soulevées par la communauté scientifique sur l'augmentation régionale de l'intensité et de la fréquence des HAB, expliquée par (i) une augmentation des programmes de surveillance due à une meilleure prise de conscience, (ii) l'intensification de l'eutrophisation et aussi (iii) le changement global. Par la suite, des projets nationaux et internationaux ont vu le jour, comme le récent projet européen CoCliME (Co-development of Climate services for adaptation to change Marine Ecosystems). Ce projet de 3 ans (2017-2020) vise au développement et à la production de services climatiques régionaux pour faire face à l'impact du changement climatique sur les écosystèmes côtiers à travers l'Europe et par la suite sur la santé humaine, l'aquaculture, la pêche et tourisme (https://www.coclime.eu/About). Pour ce faire, le consortium CoCliME rassemble des spécialistes des sciences naturelles et sociales, des décideurs et des utilisateurs finaux des services climatiques.

Contexte et objectifs

Pour ce projet, le laboratoire Phycotoxines de l'IFREMER s'est concentré sur les espèces toxiques du genre Dinophysis. En effet, après plusieurs essais, la culture de Dinophysis a été implantée avec succès pour la première fois en France dans ce laboratoire en 2015. La thèse s'est focalisée sur plusieurs facteurs clés du changement global qui peuvent moduler, d'un point de vue écophysiologique, la chaîne trophique et Dinophysis. De plus, la thèse visait à caractériser les effets de Dinophysis et de ses toxines sur deux composants de l'écosystème avec des niveaux trophiques plus élevés, à savoir les bivalves filtreurs et les poissons.

Les données issues de nos expériences en laboratoire pourraient aider les modélisateurs environnementaux à développer des modèles, ainsi qu'à améliorer les connaissances sur Dinophysis et sa chaîne trophique dans les conditions environnementales futures.

Le projet de thèse (2017-2020) fait donc partie du projet CoCliME et est intitulé :

« Études écophysiologiques sur Dinophysis et sa chaîne alimentaire, et effets in vitro du dinoflagellé et de ses toxines sur les premiers stades de vie de deux modèles d'animaux marins (l’huître et le poisson) ».

Le manuscrit est composé d'un premier chapitre qui est une introduction avec un état de l'art. Ensuite, le deuxième chapitre présente un article accepté et un article soumis qui portent sur les expériences écophysiologiques sur la chaîne trophique et Dinophysis. Enfin, le chapitre trois présente un article accepté et un article en préparation pour soumission, qui correspondent aux études des effets de Dinophysis et leurs toxines envers les organismes marins.

La section suivante décrit les 3 chapitres de ce manuscrit, comprenant une brève introduction et une description du contenu, des principaux objectifs et des questions.

Le Chapitre 1 est une revue de la littérature, qui commence par une description générale des dinoflagellés et un focus sur le genre Dinophysis. Ensuite, une description de la mixotrophie et de la culture de la chaîne trophique de Dinophysis est présentée, et une revue des expériences écophysiologiques réalisées sur Dinophysis est faite. Une description du syndrome DSP et des toxines associées est présentée, avec un accent sur l'effet sur les organismes marins. Enfin, une représentation des mesures DST dans le monde et au niveau régional est présentée.

Contexte et objectifs

Le Chapitre 2 est consacré aux expériences écophysiologiques menées dans le cadre de cette thèse.

Le Chapitre 2-Partie 1 correspond à l'étude des effets des facteurs clés du changement global sur la physiologie de Teleaulax amphioxeia et l’effet du ratio de nutrition sur son prédateur Mesodinium rubrum. Bien que T. amphioxeia est le principal donneur de plastes dans la chaîne trophique de Dinophysis et qu’il est distribué dans le monde entier, les études en laboratoire axées sur l'influence des facteurs abiotiques sur la physiologie de T. amphioxeia restent rares. Les effets de la température (13 à 25° C), de la lumière (40 à 800 µmol photons m-2 s-1) et du pH (6,5 à 8,6) sur la croissance, la teneur en pigment et le rendement quantique maximal du photosystème II de T. amphioxeia ont été évalués à l'aide d’un plan factoriel, suivi d'une tentative de déterminer l'effet de différentes photoacclimatations de T. amphioxeia sur M. rubrum.

- Comment T. amphioxeia est-il influencé par trois facteurs clés du changement global ?

- Quelles sont les conditions de culture optimales pour la croissance de T. amphioxeia ?

- Existe-t-il un lien entre la croissance, la teneur en pigment et le rendement quantique maximal du photosystème II de T. amphioxeia et de M. rubrum?

- Comment les connaissances sur la physiologie de T. amphioxeia peuvent-elles expliquer la dynamique environnementale de M. rubrum et Dinophysis dans un contexte de changement global ?

Ce Chapitre 2-Partie 1, a été publié dans « Journal of Phycology » (Facteur d’impact 2019 : 2.33) en janvier 2020 :

Gaillard, S., Charrier, A., Malo, F., Carpentier, L., Bougaran, G., Hégaret, H., Réveillon, D., Hess, P., Séchet, V., 2020. Combined effects of temperature, irradiance, and pH on Teleaulax amphioxeia (Cryptophyceae) physiology and feeding ratio for its predator Mesodinium rubrum (Ciliophora). J. Phycol. 56, 775-783. https://doi.org/10.1111/jpy.12977. Open access: https://hal.archives- ouvertes.fr/hal-02880016

Contexte et objectifs

Le Chapitre 2-Partie 2 examine l'effet d'un stress de salinité à court terme (conditions hypo- hyperosmotiques à la salinité 25 et 42, respectivement) sur trois souches françaises de D. sacculus isolées de différentes régions. Bien que des études sur le terrain suggèrent un lien entre les occurrences de salinité et de Dinophysis, aucune expérience in vitro n'a été réalisée sur l'effet d'un stress de salinité sur la physiologie de D. sacculus, les toxines, les osmolytes et les profils métaboliques.

- Comment D. sacculus peut-il tolérer ces variations osmotiques ?

- Quelle est la réponse des osmolytes, des toxines et des autres métabolites (métabolomes) de D. sacculus aux stress de salinité ?

- Les teneurs en phycotoxines sont-elles modifiées et agissent-elles comme des osmolytes ?

- Y a-t-il un impact de l'origine de la souche de D. sacculus et de ses habitats sur la tolérance à la salinité ?

Ce Chapitre 2-Partie 2, est en révision dans le numéro spécial CoCliME dans « Harmful Algae » (Facteur d’impact 2019 : 3.71) depuis septembre 2020.

Contexte et objectifs

Le Chapitre 3 étudie les effets de Dinophysis et leurs toxines sur deux organismes marins.

Le Chapitre 3-Partie 1 décrit les effets de l'exposition de D. sacculus, D. acuminata et de leurs toxines sur les gamètes d’une espèce d’intérêt commercial, l'huître creuse du Pacifique (Crassostrea gigas) avec un focus sur la physiologie des gamètes et sur le succès de fécondation. Les effets toxiques indirects de Dinophysis et ses toxines sur l'homme par la consommation de mollusques contaminés sont bien connus, mais l’effet direct sur les bivalves est mal étudié. Cette étude examine les effets de deux espèces toxiques du genre Dinophysis (D. acuminata et D. sacculus), et de deux de leurs toxines (OA et PTX2) sur la physiologie des gamètes et le succès de la fécondation de C. gigas, une espèce d'intérêt commercial :

- Y a-t-il un effet de Dinophysis sur les gamètes et par la suite, sur le succès de fécondation de l'huître ? Quels compartiments (culture, cellules remises en suspension ou filtrat) ont un effet sur le succès de la fécondation ? L'effet est-il spécifique à l'espèce ?

- Y a-t-il un effet des toxines de Dinophysis sur les gamètes et par la suite, sur le succès de la fécondation de l'huître ?

- Les ovocytes et les spermatozoïdes sont-ils affectés différemment ? L'effet est-il cumulatif sur le succès de la fécondation ?

- Les Dinophysis et leurs toxines sont-ils délétères au recrutement des huîtres ? Qu'en est-il de la cooccurrence environnementale de Dinophysis et des larves d'huîtres ?

Ce Chapitre 3-Partie 1 a été publié dans Environmental Pollution (Facteur d’impact 2019 : 6.8) en mai 2020 :

Gaillard, S., Le Goïc, N., Malo, F., Boulais, M., Fabioux, C., Zaccagnini, L., Carpentier, L., Sibat, M., Réveillon, D., Séchet, V., Hess, P., Hégaret, H., 2020b. Cultures of Dinophysis sacculus, D. acuminata, and pectenotoxin 2 affect gametes and fertilization success of the Pacific oyster, Crassostrea gigas. Environ. Pollut. 265, 114840. https://doi.org/10.1016/j.envpol.2020.114840. Open access: https://hal.archives-ouvertes.fr/hal-02880050/

Contexte et objectifs

Le Chapitre 3-Partie 2 étudie la mortalité et l'histopathologie des larves d’un téléostéen, la fondule tête de mouton (Cyprinodon variegatus) après une exposition de 96 h à Dinophysis acuminata et à la PTX2. Peu d'études ont évalué l'effet des DST, de Dinophysis, et encore moins de la PTX2 sur les animaux marins et en particulier sur les premiers stades de vie des poissons. Ce travail a été réalisé en collaboration avec Wolfgang Vogelbein et son équipe du Virginia Institute of Marine Science (VIMS, Gloucester Point, Virginie, USA) de février à mai 2020. Cette section est présentée comme un article en préparation pour soumission, cependant certaines observations histopathologiques sont en cours d’analyse, mais des résultats préliminaires sont fournis.

- Y a-t-il un effet de D. acuminata sur la mortalité des larves ? Quels compartiments (culture, cellules remises en suspension, filtrat ou lysat) sont responsables du ou des effets observés ?

- Y a-t-il un effet de la PTX2 sur la mortalité larvaire ?

- Quels sont les effets histopathologiques ?

- Est-ce que les pathologies observées sont connues et comparables aux effets induits par d'autres toxines (par exemple la gonodiomine A, une autre lactone macrocyclique produite par Alexandrium spp.) ?

- Est-ce que Dinophysis et la PTX2 sont des menaces environnementales pour les poissons et en particulier dans la baie de Chesapeake ?

Context and aims of the thesis

A basic definition of the term plankton is an organism, which is adapted to live in suspension in sea water, rivers, lakes and ponds (Reynolds, 2006). The plankton regroups living organisms from the size of viruses (nanometers) to jellyfish (meters), which are not able to control their horizontal position or to swim against currents (plankton derived from the Greek word for errant, wanderer). Among them, the single-celled plants -the phytoplankton- represents the photosynthetic microorganisms. Phytoplankton organisms are at the basis of the aquatic food-chain and a major primary producers of organic carbon in waters (Falkowski and Raven, 1997; Reynolds, 2006). Indeed, thanks to the phytoplankton, the majority of photosynthesis on Earth occurs in aquatic environment, where the consumption of carbon dioxide from the atmosphere is equivalent to forests and other land plants and thus, participate in control of global warming (Field et al., 1998). Phytoplankton organisms can be classified according to the structure of their plastids (= membrane bound organelles), membrane arrangements and pigments into five groups: green algae, blue-green algae, coccolithophores, diatoms and dinoflagellates (Falkowski et al., 2004; Pachiappan et al., 2019).

Some phytoplankton can be named harmful algae when causing deleterious effects from simple nuisances, such as beach fouling and anoxia, to the toxic effects on aquatic organisms as skin irritation or suffocation of fish, and food-poisoning of humans caused by consumption of shellfish, fish or mammals through bioaccumulation of toxins within the food webs (Granéli and Turner, 2006; Smayda, 1997a). These phenomena can be enhanced during blooms (i.e. significant and rapid increases of the population compared to the classic concentration) of harmful algae, known as harmful algal blooms (HABs).

A phytoplankton bloom can be triggered or inhibited depending on both biotic (e.g. nutrition, competition, grazing) and abiotic (e.g. hydrology, meteorology) factors. For toxin-producing phytoplankton, a bloom can also be defined as an increase of measurable algal toxin (i.e. toxin synthesized by phytoplankton) levels, which can vary in time, duration and location (Smayda, 1997a). Among the thousands described species of marine phytoplankton (Reynolds, 2006) around 300 can form red-tides (i.e. discoloration of water, which can be harmful) and 141 are able to

Context and aims synthesize toxins (Moestrup et al., 2009). Among the toxin producers, 50 to 75% are dinoflagellates (Hallegraeff, 1993; Moestrup et al., 2009; Smayda, 1997b).

Algal toxins are small metabolites, with a diversity of compounds, chemical structures and toxicity routes. Toxins have been classified according to their toxic effects and their syndromes in humans after the consumption of contaminated filter-feeding bivalves, including oysters, mussels or clams, and fish. The major intoxications are diarrheic shellfish poisoning (DSP), paralytic shellfish poisoning (PSP), ciguatera poisoning (CP), amnesic shellfish poisoning (ASP), neurotoxic shellfish poisoning (NSP) or azaspiracid shellfish poisoning (AZP) (Hallegraeff, 2004; Lassus et al., 2016; Lawrence et al., 2011; Rossini and Hess, 2010; Young et al., 2020). In addition to the sanitary effects, economic and societal issues also frequently arise as a consequence of HABs (Glibert et al., 2014; Reguera et al., 2012). Consequently and because of the increase of recorded human intoxication by algal toxins in the second part of the last century (Hallegraeff, 2010; Lassus et al., 2016), authorities have implemented detection and monitoring programs of toxic phytoplankton species and contaminated filter-feeding bivalves. Sanitary specific thresholds (of phytoplankton and/or algal toxins) have been promulgated according to several directives (e.g. European, Japanese and American directives; EU.Commission 2011, DeGrasse and Martinez-Diaz 2012, Hess 2012, Suzuki and Watanabe 2012, NSSP 2017). Above these thresholds, the authorities (e.g. prefectures in France) can ban shellfish harvesting and associated activities (including transportation, sales and distribution), as well as recreational harvesting, for periods going from days to months (Belin et al., 2020). The economic losses of these bans on shellfish farming closures and subsequent societal effects (e.g. loss of activity or unemployment) reached more than 1 billion USD in 2006 in Europe and USA, which may even be higher nowadays (Granéli and Turner, 2006; Hoagland et al., 2020; Hoagland and Scatasta, 2006).

In France, the REPHY-REPHYTOX (Observation and Surveillance Network for Phytoplankton and Hydrology in coastal waters; referred to as REPHY in the manuscript) monitoring network was created in 1984 by the French Research Institute for Exploitation of the Sea (IFREMER). This ongoing network aims to acquire data on hydrological parameters and on the abundance and composition of phytoplankton as well as to monitor algal toxins in filter-feeding bivalves (Belin et al., 2020; REPHY, 2019). Toxic diatoms of the genus Pseudo-nitzschia and dinoflagellates of the genera Alexandrium and Dinophysis and their associated toxins are regularly found in water and in

Context and aims shellfish from the French Atlantic Ocean, Mediterranean Sea and English Channel coasts (Belin et al., 2020). The first listed event of intoxication in France was recorded in 1983 and was due to the accumulation of DSP toxins in mussels from toxic dinoflagellate species of the genus Dinophysis (Ehrenberg, 1841; Lassus et al., 1985), with symptoms in humans of gastrointestinal intoxication such diarrhea, nausea, vomiting and abdominal pain. Since this outbreak that led to the establishment of the REPHY in 1984, Dinophysis is the most emblematic genus in France, as cells and toxins are found every year in water and mollusks and are mainly responsible for bans despite usually low cell concentrations (≤ 100 cells L-1; Belin et al., 2020).

Several species of the genus Dinophysis are recognized as the main causative agent of DSP (Reguera et al., 2014). Among the 133 species of genus Dinophysis accepted taxonomically (Guiry and Guiry, 2020), ten species of Dinophysis have been related to DSP outbreaks and/or characterized as diarrheic shellfish toxin (DST) producers as well as two species of the genus Phalacroma (Nagai et al., 2020): D. acuminata (Claparède & Lachmann 1859), D. acuta (Ehrenberg, 1839), D. caudata (Saville Kent, 1881), D. fortii (Pavillard, 1923), D. infundibulum (Schiller, 1928), D. miles (Cleve, 1900), D. norvegica (Claparède and Lachmann, 1859) D. ovum (Schütt, 1895), D. sacculus (Stein 1883) and D. tripos (Gourret, 1883), and Phalacroma mitra (Schütt, 1895) (= D. mitra; Abé, 1967) and Phalacroma rotundatum (Kofoid and Michener, 1911) (= D. rotundata; Claparède and Lachmann, 1859).

These toxic species synthesize two types of lipophilic algal toxins, i.e. okadaic acid (OA; polyether) and its derivates, the dinophysistoxins (DTXs), which together form the OA-group of toxins, and the pectenotoxins (PTXs; macrocyclic polyether lactones). OA-group contained diarrheic toxins, while PTXs, also often found in shellfish due to the co-production by Dinophysis, do not cause diarrhea (Cohen et al., 1990; Reguera and Pizarro, 2008). It is noteworthy that while the toxicity of OA-group and PTXs has been well described on rodent models, their effects (and the ones derived directly from the Dinophysis cells) on marine organisms e.g. mollusks or fish, and especially on their early life stages which are critical development phases, are still poorly known and yet they are regularly contaminated.

DSP outbreaks and associated DSTs have been listed worldwide since the first mass intoxication that happened in the Netherlands in 1961 (Korringa and Roskam, 1961). For the next 45 years, scientific data were obtained only from field study and/or materials (i.e. picked plankton cells)

Context and aims collected in situ (e.g. Yasumoto et al. 1978, Kumagai et al. 1986, Cembella 1989, Masselin et al. 1992, Vale and Sampayo 2002, Luisa Fernández et al. 2006). The first attempt to cultivate Dinophysis, in « classic » culture media (i.e. seawater enriched with nutrients, trace metals and vitamins) for autotroph organisms dates from 1935 (Reguera et al., 2012) but was unsuccessful. After that, several attempts to cultivate Dinophysis with prey (pico- nanoplankton) or bacteria were also unsuccessful (Ishimaru et al., 1988; Maestrini et al., 1995). These fruitless experiments were in fact due to the mode of nutrition of Dinophysis.

Protists exhibit different kind of trophic interactions, including mixotrophy, which is the combination of both phototrophy and heterotrophy. The mixotrophic mode of nutrition allows protists to obtain energy, carbon and nutrients from photosynthesis and from dissolved organic carbon and/or phagotrophy (=photo-phago-mixotroph; Burkholder et al., 2008; Hansen et al., 2019). Mixotrophy is prevalent in several phytoplankton taxa, such as radiolaria, foraminifera, ciliates and dinoflagellates (Hansen et al., 2019; Stoecker, 1998).

Traditional microscopic observations suggested the mixotrophic nutrition mode of Dinophysis. Indeed, plastids of Dinophysis have been described deriving from cryptophytes (Schnepf and Elbrachter, 1988), and food vacuoles and feeding peduncle have been observed within toxic species of the genus (Hansen, 1991; Jacobson and Andersen, 1994). The cryptophyte origin of plastids was then proved through molecular analysis (Janson and Granéli, 2003). The cryptophyte Teleaulax amphioxeia (Conrad) Hill (1992) was identified as the donor of the plastids (Janson, 2004) through a ciliate vector, i.e. Mesodinium rubrum (Lohmann 1908) (Yih et al., 2004) by a process of sequestration of the plastids, i.e. kleptoplastidy (Schnepf and Elbrächter, 1999; Takishita et al., 2002). Polar strain of M. rubrum fed in culture on another cryptophyte, Geminigera cryophila (Johnson et al., 2007).

The first permanent culture of a Dinophysis species in the laboratory was established by Park et al. (2006) based on the three-member trophic (Teleaulax sp. at the bottom of the chain, Mesodinium rubrum and D. acuminata as the superior predator). Importantly, since 2006, several cultures of toxic species of Dinophysis have been established, thereby allowing researchers to conduct ecophysiological in vitro experiments. These studies gave insights into the impact of biotic or abiotic factors on e.g. growth and toxin contents of Dinophysis, especially in a context of global change.

Context and aims

Concerns have been raised by the scientific community on the regional increase of intensity and frequency of HABs, explained by (i) an increase of monitoring programs due to a better awareness, (ii) the intensification of eutrophication and also (iii) global change. Subsequently, national and international projects emerged, such as the recent European project CoCliME (Co-development of Climate services for adaptation to changing Marine Ecosystems). This 3-year project (2017-2020), aims at the development and the production of regional climate services to address the impact of climate change on coastal ecosystems and subsequently on human health, aquaculture, fisheries and tourism across Europe (https://www.coclime.eu/About). To do this, the CoCliME consortium brings together natural and social scientists, decision makers and end users of climate services.

For this project, the Phycotoxins laboratory of IFREMER focused on toxic species of the genus Dinophysis. Indeed, after several trials, the culture of Dinophysis was successfully established for the first time in France in this laboratory in 2015. The thesis focused on several key factors of the global change which can modulate, from an ecophysiological point of view, the food chain and Dinophysis. Moreover, the thesis aimed to characterize the effects of Dinophysis and its toxins on two components of the ecosystem with higher trophic levels i.e. sessile filter-feeding bivalves and fish.

Data obtained from our laboratory experiments could help environmental modelers to the development of models as well as improve knowledge on Dinophysis and its food chain in future environmental conditions.

The thesis project (2017-2020) is therefore a part of the CoCliME project and is entitled:

“Ecophysiological studies on Dinophysis and its food chain, and in vitro effects of the dinoflagellate and its toxins on early life stages of two models of marine animals (oyster and fish)”.

The manuscript is composed of a first chapter which is an introduction with a state of the art. Then the second chapter presents one accepted and one submitted article which focus on the ecophysiological experiments on the food chain and Dinophysis. Finally, the chapter three presents one accepted article and an article in preparation for submission which correspond to the studies of the effects of Dinophysis and their toxins towards marine organisms.

Context and aims

The following section describes the 3 chapters of this manuscript, including a brief introduction and description of the content, main objectives and questions.

Chapter 1 is a literature review, which starts with a general description of the dinoflagellates and a focus on the genus Dinophysis. Then, a description of the mixotrophy and the culture of the food chain of Dinophysis is presented, and a review of the ecophysiological experiments performed of Dinophysis is made. A description of the syndrome DSP and the associated toxins is reported, with a focus on the effect on marine organisms. Finally, a representation of DST measurements worldwide and regionally is presented.

Chapter 2 is dedicated to the ecophysiological experiments conducted as part of this thesis.

Chapter 2-Part 1 corresponds to the study of the effects of key factors of global change on the physiology of Teleaulax amphioxeia and the ratio of nutrition for its predator Mesodinium rubrum. While T. amphioxeia is the main donor of plastids in the food chain of Dinophysis and is globally distributed, laboratory studies focusing on the influence of abiotic factors on T. amphioxeia physiology remain scarce. Effects of temperature (13 to 25 °C), light (40 to 800 µmol photons m-2 s-1) and pH (6.5 to 8.6) on growth, pigment content and maximum quantum yield of the photosystem II of T. amphioxeia were assessed using a factorial design approach, followed by an attempt to determine the effect of contrasted photoacclimation of T. amphioxeia on M. rubrum.

- How is T. amphioxeia influenced by the three studied key factors of global change?

- What are the optimal culture conditions for T. amphioxeia growth?

- Is there a link between the growth, pigment contents and the maximum quantum yield of the photosystem II of both T. amphioxeia and M. rubrum?

- How can knowledge on the physiology of T. amphioxeia explain environmental dynamics of M. rubrum and Dinophysis in a context of global change?

Context and aims

This Chapter 2-Part 1, has been published in Journal of Phycology (IF 2019: 2.33) in January 2020:

Chapter 2-Part 2 examines the effect of a short-term salinity stress (hypo- hyperosmotic conditions at salinities 25 and 42, respectively) on three French strains of D. sacculus isolated from different regions. While field studies suggested a link between salinity and Dinophysis occurrences, no in vitro experiments have been performed on the effect of a quick salinity stress on D. sacculus physiology, toxins, osmolytes and metabolite profiles.

- How can D. sacculus tolerate these osmotic variations?

- What is the response of known osmolytes, toxins and other metabolites (metabolomes) of D. sacculus to the salinity stresses?

- Are algal toxin contents modified and are they acting like osmolytes?

- Is there an impact of the origin of the strain of D. sacculus and its habitats on the salinity tolerance?

This Chapter 2-Part 2, is in revision in the special issue CoCliME in Harmful Algae (IF 2019: 3.71) since September 2020.

Context and aims

Chapter 3 studies the effects of Dinophysis and their toxins on two marine organisms.

Chapter 3-Part 1 describes the effects of the exposure of gametes of a species of commercial interest, the Pacific oyster (Crassostrea gigas), to D. sacculus, D. acuminata and their toxins with a focus on gametes physiology and fertilization success. The indirect toxic effects of Dinophysis and its toxins on humans through the consumption of contaminated shellfish is well known, but the direct effect on the bivalves is poorly studied.

- Is there an effect of Dinophysis on gametes and subsequently, on the fertilization success of oyster? Which compartments (culture, resuspended cells or filtrate) have an effect on fertilization success? Is the effect species-specific?

- Is there an effect of Dinophysis toxins on gametes and subsequently, on the fertilization success of oyster?

- Are oocytes and spermatozoa differentially affected? Is the effect cumulative on fertilization success?

- Are Dinophysis and their toxins detrimental for oyster recruitment? What about the environmental co-occurrence of Dinophysis and oyster larvae?

This Chapter 3-Part 1, has been published in Environmental Pollution (IF 2019: 6.8) in May 2020:

Context and aims

Chapter 3-Part 2 investigates mortality and histopathology of teleost fish larvae, sheepshead minnow (Cyprinodon variegatus) following a 96 h exposure to Dinophysis acuminata and PTX2. Few studies assessed the effect of DSTs/Dinophysis, and even less of PTX2 on marine animals and especially on early life stages of fish. This work was made in collaboration with Wolfgang Vogelbein and his team from the Virginia Institute of Marine Science (VIMS, Gloucester Point, Virginia, USA) in February to May 2020. This section is presented as an article in preparation for submission, however histopathological observations are ongoing but preliminary results are provided.

- Is there any effect of D. acuminata on larval mortality? Which compartments (culture, resuspended cells, filtrate or lysate) are responsible for the observed effect(s)?

- Is there any effect of PTX2 on larval mortality?

- What are the histopathological effects?

- Are the pathologies known comparable to the effects induced by other toxins (e.g. gonodiomin A, another macrocyclic lactone produced by Alexandrium spp.)?

- Are Dinophysis and PTX2 environmental threats for fish and especially in the Chesapeake Bay?

Chapter 1: State of the art - Literature review

Chapter 1: Literature review

I-1) General characteristics of the dinoflagellates

The dinoflagellates are one of the most important phytoplanktonic and phytobenthic group of microalgae in term of number of species (more than 2000; Falkowski et al., 2004). There are unicellular and eukaryotic dinoflagellates living in diverse habitat (e.g. pelagic, benthic), having several modes of nutrition (e.g. photosynthetic, heterotrophic, mixotrophic) and different morphologies (e.g. thecate, athecate, presence or absence of extrusomes) (Taylor et al., 2003). There are special features to all members of the dinoflagellate group e.g. (i) the spiraling movement, (ii) the presence of amphiesma and (iii) the dinokaryon (Spector, 1984). (i) The spiraling movement is due to the presence of two flagella, inserted orthogonally. Movements differ according to the type of dinoflagellates, i.e. desmokont (flagella not associated with furrows) or dinokont (flagella inserted in furrows). The transversal furrow is named cingulum and the longitudinal furrow is the sulcus. The cingulum divides the cell in epitheca and hypotheca (Figure 1 A) (Spector, 1984; Taylor, 1980). (ii) The amphiesma covers the cell and is composed of an external plasma membrane, an amphiesmal vesicle and an internal membrane. Amphiesmal vesicles can contain cellulosic materials (thecal plates) or not (athecate) (Höhfeld and Melkonian, 1992). Traditional morphological taxonomy uses the organization/position of the thecal plates on epitheca and/or hypotheca (see dinoflagellates tabulation, Kofoid System; Taylor et al. 2003) to differentiate between the dinoflagellate groups. (iii) The dinoflagellates nucleus is called dinokaryon; it contains chromosomes always condensed, no histones and usually a huge genome (e.g. the dinoflagellates Prorocentrum micans has a genome ca. 85 times greater than the human genome) (Fukuda and Suzaki, 2015; LaJeunesse et al., 2005).

I-2) Morphology and taxonomy of Dinophysis

Dinophysis (Chromalveolata, Myzozoa, Dinophyceae) (Ehrenberg, 1841) is a genus of thecate dinoflagellates, laterally compressed, dinokont type with a girdle and sulcal lists (Figure 1 A). These characters and the size and contour of the large hypothecal plates and their ornamentation can be used by taxonomists for Dinophysis species discrimination/identification (Jensen and Daugbjerg, 2009; Taylor et al., 2003). 44

Chapter 1: Literature review

The holotype species, D. acuta, is characterized by an outline with a wider depth under the median part, and a triangular shape at their posterior end (Figure 1 A) (Séchet et al., 2020). D. caudata has a prominent ventral projection of the hypotheca (Figure 1 B) and D. tripos has two antapical projections, a short (ventral) and a long (dorsal) (Figure 1 C) (Séchet et al., 2020).

As morphological differences are almost absent between D. acuminata, D. sacculus and D. ovum (Figure 1, D, E and F, respectively), which havemore or less oval cells, the term D. acuminata- complex has been introduced to group these three species (Lassus and Bardouil, 1991). Moreover, species of the D. acuminata-complex often co-occur. As phytoplankton identification in monitoring program is usually made by microscopic analyses (e.g. REPHY in France), identification is limited to the genus level of Dinophysis. Initially, molecular taxonomy for identification of Dinophysis presented problems of genetic delineation with the use of several genes (i.e. rDNA subunit or ITS). Now, identification is based on cytochrome C oxidase I gene (cox1) (Raho et al., 2008; Wolny et al., 2020), but the problem of identification with the D. acuminata- complex is not solved (Séchet et al., 2020).

Chapter 1: Literature review

Epitheca

Anterior cingular list

Posterior cingular list Sulcal list

Hypotheca

Chapter 1: Literature review

I-3) Dinophysis food chain

After several unsuccessful attempts to cultivate Dinophysis in the laboratory (Ishimaru et al., 1988; Maestrini et al., 1995), Park et al. (2006) established for the first time a permanent culture of Dinophysis. This culture was based on the three-tier food chain Teleaulax sp. - Mesodinium rubrum - Dinophysis acuminata (Figure 2). Toxic species of the genus Dinophysis and M. rubrum are both mixotrophic species (i.e. here combination of photosynthesis and heterotrophy via phagotrophy): M. rubrum feeds on cryptophytes (e.g. T. amphioxeia) and Dinophysis feed on M. rubrum.

I-3-1) First tier of the food chain: T. amphioxeia

Cryptophytes and especially species belonging to the clade Teleaulax/Plagioselmis/Geminifera (TPG) (Hoef-Emden, 2008) are commonly found in marine coastal environments (Adolf et al., 2006; Cerino and Zingone, 2007). For instance, in the French Atlantic, Mediterranean Sea and English Channel coasts, the taxon cryptophyte is one of the most observed in water samples (Belin et al., 2020; REPHY, 2019).

Cryptophytes have a high nutritional value (Lee et al., 2019), thus they are important and common prey organisms of several protists, including M. rubrum (Peterson et al., 2013; Yih et al., 2004) (see section “I-3-2) Mixotrophy of Mesodinium rubrum”). Teleaulax amphioxeia (Chromista, Cryptophyta, Geminigeraceae) (Conrad) Hill (1992) has been widely used and referenced as prey of M. rubrum in culture (Park et al., 2007; Smith et al., 2018) (see section “I-4-1) Culture of Teleaulax amphioxeia”). Moreover, environmental studies established a link between the initiation

Chapter 1: Literature review of M. rubrum bloom and the decline of Teleaulax abundance (Hamilton et al., 2017; Peterson et al., 2013).

Despite the importance of this taxon, there is a lack of data regarding laboratory studies on T. amphioxeia physiology. A recent study showed the intraspecific variability of the qualitative and quantitative nutritional composition (i.e. proteins, lipids and carbohydrates) of six mass-cultured strains of T. amphioxeia for biotechnological purposes (Lee et al., 2019). Fiorendino et al. (2020) observed a significant effect of temperature and irradiance on two strains (from USA and Denmark) of T. amphioxeia and maximum growth rate at high temperature (24-27 °C), irradiance (400 µmol photons m-2 s-1) and intermediate salinity 30. However, no study focused on the impact of abiotic factors on T. amphioxeia pigment contents and photosynthetic activity, and how these parameters may affect the higher levels (i.e. M. rubrum and Dinophysis) of the food chain.

I-3-2) Mixotrophy of M. rubrum

The photosynthetic ciliate Mesodinium rubrum (Chromista, Ciliophora, Cyclotrichiida) (Lohmann 1908) forms red tides in coastal ecosystems worldwide (Herfort et al., 2011; Lindholm, 1985). In the 1970s, ultrastructural observations of M. rubrum revealed the cryptophytic origin of its plastids, mitochondria and nuclei (see references in Johnson and Stoecker 2005). In 2000, Gustafson et al. observed ingestion and uptake of plastids by M. rubrum from a cryptophyte (Teleaulax acuta) that was required for sustained growth. The first permanent culture of M. rubrum was established by Yih et al. (2004) with Teleaulax sp. as prey. Interestingly, two hypotheses exist on the degree of relationship between M. rubrum and its cryptophyte prey: is it an endosymbiosis (i.e the cryptophyte organelles are endosymbiont) or is it kleptoplastidy without harboring a permanent endosymbiont? While M. rubrum can starve in the light without feeding on prey, there is a loss of chloroplasts after successive divisions (Hansen and Fenchel, 2006; Johnson et al., 2006; Johnson and Stoecker, 2005). However, M. rubrum has the ability to synthesize chlorophyll a and divides and photoacclimates its plastids (Figure 2) (Hansen and Fenchel 2006, Johnson et al. 2006, Moeller et al. 2011). This regulation of plastids (i.e. photoacclimation and division) may be possible thanks to the uptake of prey nuclei (Figure 2) (Johnson, 2011; Kim et al., 2017; Wisecaver and Hackett, 2010), which remain transcriptionally active in M. rubrum (Johnson et al., 2007; Lasek-Nesselquist et al., 2015), but are lost after successive divisions (Kim et al., 2017). Even in the presence of prey,

Chapter 1: Literature review

M. rubrum cannot grow indefinitely in the dark: it is “an obligate mixotroph that depends on phototrophy”, with a predominance of phototrophy (only 1-2 % of carbon obtained from prey when fed 1 prey d-1) (Hansen et al., 2013; Smith and Hansen, 2007). Johnson et al. (2006) suggested that M. rubrum may be “in the transition to permanently acquired plastids”. As mentioned by Johnson et al. (2017), M. rubrum “do not fit into established boxes”, thus this complex and unique relation between M. rubrum and its cryptophyte prey can be named as “incorporation of ingested prey cells, which are reduced symbionts” (Hansen et al., 2013). M. rubrum has been found to preferably feed cryptophytes belonging to the TPG (Teleaulax/Plagioselmis/Geminifera) clade (Gustafson et al., 2000; Hansen et al., 2012; Park et al., 2007) and the temperate strains of M. rubrum especially fed the red tides forming species, T. amphioxeia (Yoo et al. 2017) (see sections “I-4-1) Culture of Teleaulax amphioxeia” and “I-4-2) Culture of Mesodinium rubrum”).

The prey capture mechanism has been well described by Yih et al. (2004). First a physical contact is made between Teleaulax sp. cells and the ciliate’s oral tentacles, followed by a fixation of the cells. Then, Teleaulax sp. prey reach a cytosome-like structure and is engulfed by M. rubrum. Prey are then successively transported to the equatorial cirri and to the aboral end. The duration range of the process is ca. 1 to 1.5 minutes (Yih et al., 2004). According to the prey concentration, M. rubrum may harbor between 6 and 36 chloroplasts (Hansen and Fenchel, 2006) and from 1 prey nucleus at low prey concentration to a maximum of 11 (in addition to a centered prey nucleus, see Kim et al., 2017).

Chapter 1: Literature review

Geminigera cryophila Mesodinium rubrum Dinophysis acuminata (or Teleaulax amphioxeia) (or Dinophysis spp.)

Figure 2: Acquisition of the cryptophyte plastids in M. rubrum and Dinophysis. From Wisecaver and Hackett (2010). The three organism drawings are not to scale. G. cryophila is the plastids donor of M. rubrum and Dinophysis from polar waters (appendices Table 34). Nucleus (A), mitochondria and plastids (B) of the cryptophytes are retained in M. rubrum. In Dinophysis there is a loss of the outer membranes of the plastids and nucleomorph (C). Cryptophyte nucleus (1), plastid (2), nucleomorph (3), mitochondrion (4) and nucleus and cytoplasms surrounded by M. rubrum membrane (5). Ciliate nucleus (6), plastid-mitochondrial complex surrounded by M. rubrum membrane (7) and mitochondrion (8). Dinophysis nucleus (9), kleptoplast (10) and mitochondrion (11)

I-3-3) Mixotrophy of Dinophysis

The cryptophytic origin of Dinophysis organelles was highlighted by both microscopic (Jacobson and Andersen, 1994; Schnepf and Elbrachter, 1988) and molecular analyses (Hackett et al., 2003; Janson, 2004; Janson and Granéli, 2003; Minnhagen and Janson, 2006; Takishita et al., 2002), which finally led to the establishment of its culture using a ciliate vector to get its plastids (Park et al., 2006). However, Dinophysis only retained chloroplasts but neither the mitochondria nor the nuclei, after ingestion (Figure 2) (Hansen et al., 2013). Similarly to M. rubrum, controversy about the transient (i.e. sequestered) or permanent nature (Garcia-Cuetos et al. 2010) of Dinophysis

Chapter 1: Literature review plastids have been raised. Plastidic modifications inside Dinophysis have been highlighted by Kim et al (2012a) who showed physiological changes in the stolen plastids (i.e. position of the pyrenoids) and a transformation (i.e. position of the pyrenoids and decrease in number of thylakoids) and transfer of these plastids. Dinophysis is able to photoregulate/photoacclimate its ingested plastids (García-Portela et al., 2018b; Hansen et al., 2016) and divide them (Rusterholz et al., 2017), and carbon uptake from prey accounts for 70-90 % when prey concentration is high (Kim et al., 2008; Riisgaard and Hansen, 2009) to 0-55 % when concentration of prey is low (Riisgaard and Hansen, 2009). Dinophysis has only five chloroplastic housekeeping genes to regulate the ingested plastids and only one from cryptophytic origin (Wisecaver and Hackett, 2010). These findings suggest that Dinophysis has kept chloroplast-regulating genes from an ancestor and/or these genes were transferred from a prey (Hansen et al., 2016; Rusterholz et al., 2017). Similarly to M. rubrum, Dinophysis may be in a transitional phase to a permanent plastids acquisition (Rusterholz et al., 2017).

A description of the prey capture mechanisms of M. rubrum by D. acuta and/or D. acuminata was made by Ojamäe et al. (2016) and Mafra et al. (2016). M. rubrum is known for its fast jumps (i.e. fastest inside ciliates group) (Fenchel and Hansen, 2006) while Dinophysis swims slowly. In the presence of Dinophysis, M. rubrum has been observed to be trapped in mucus, which resulted in a decrease of swimming activity and speed, and ultimately, immobilization (Ojamäe et al., 2016). Mafra et al., (2016) also observed a decrease in swimming frequency of M. rubrum in the presence of Dinophysis before being attached to the mucus traps, and suggested the effect of allelopathic compound(s) (see section “I-8-1) Allelopathic effects”). Mucus may be synthestised by Dinophysis or by M. rubrum due to the action of allelopathic compound(s) (Ojamäe et al., 2016). These authors observed in trapped M. rubrum a loss of cirri and changes in cell shape (becoming rounded). Dinophysis attached to M. rubrum by a feeding peduncle and ingested its contents during minutes to hours until prey emptying (Mafra et al., 2016; Ojamäe et al., 2016). Number of kleptochloroplasts inside Dinophysis varied according to prey concentration and time, and is around 28 for D. acuta and 20 for D. acuminata (well-fed cultures) (Rusterholz et al., 2017)

To conclude, both M. rubrum and Dinophysis are plastidic specialists (with prey preferences) (Hansen et al., 2012; Park et al., 2007; Smith et al., 2018) non-constitutive mixoplancton (phototrophy acquires from a prey) (Flynn et al., 2019; Hansen et al., 2019; Mitra et al., 2016). M.

Chapter 1: Literature review rubrum may regulate its ingested plastids by prey nuclei and/or nucleomorph (i.e cryptophyte reduced nucleus acquired from an earlier endosymbiont) and Dinophysis by ancient or transferred genes (Hansen et al., 2016). Observations suggest that both organisms may be in a transition to acquire permanent chloroplasts. Molecular studies are still needed to fully explain the kleptochloplast control by Dinophysis and nuclei by M. rubrum. Moreover, the donor of plastids at the basis of the food chain in most cases is Teleaulax amphioxeia. Thus, studies on this cryptophyte appear also fundamental to explain physiology of the predators.

I-4) Culture of the three-tier food chain of Dinophysis.

Culture of Dinophysis is time consuming and requires precaution, because three organisms with different growth rates need to be cultivated at the same time, and the fast-growing autotrophic organism (i.e. T. amphioxeia) can smother the ciliate M. rubrum and outcompete it (e.g. by increase of pH) (Smith and Hansen, 2007). It could be interesting to harmonize culture conditions between laboratory as well as found optimal culture conditions to maximize Dinophysis growth when performing experiments. The tables 34, 35 and 36-39 in appendices represent the culture conditions of cultivated species and strains of respectively, Teleaulax amphioxeia (and other species of the TPG clade), Mesodinium rubrum and Dinophysis since 2006.

The culture of the food chain of Dinophysis has successively been established in 9 countries and 11 laboratories: in Republic of Korea (Park et al., 2006), Denmark (Smith and Hansen, 2007), Japan (Nagai et al., 2008), USA-Massachusetts (Hackett et al., 2009), Spain (Rodríguez et al., 2012), USA-New York (Hattenrath-Lehmann et al., 2013), Brazil (Mafra et al., 2014), New- Zealand (Papiol et al., 2016), China (Gao et al., 2017), USA-Virginia (Smith et al., 2018) and France (Séchet et al., 2020). However, only three laboratories (in Japan, Republic of Korea and Denmark) used their own locally isolated strains of cryptophyte, ciliate and Dinophysis, and may explained the higher growth rate of Dinophysis fed with M. rubrum and T. amphioxeia from the same location (Hernández-Urcera et al., 2018).

Here the culture conditions for the main cultivated strains of T. amphioxeia (Table 1) and M. rubrum (Table 2) and example of cultures of D. acuminata and D. acuta (Table 3) are summarized.

Chapter 1: Literature review

Lists of all the cultivated species and strains of the food chain are presented in the appendices (Tables 34-39).

I-4-1) Culture of T. amphioxeia

As above-mentioned, species of the cryptophyte TPG clade, and especially T. amphioxeia, are the preferred food source for M. rubrum. Thereby, this latter organism is used as prey in every laboratory (appendices Table 34), except for polar strains of M. rubrum, which are fed Geminigera cryophila (Hackett et al., 2009) (appendices Table 34 and 35). The Japanese (AB364287 and TA- INO01), Spanish (AND-A0710) and Danish (K-0434) strains are the more widely cultivated (> 75 %) to sustain M. rubrum growth (Table 1). The average culture conditions for temperate strains of T. amphioxeia are 17 °C, 93 µmol photon m-2 s-1, salinity 31 and a nutrient, vitamin and trace metals rich culture medium (i.e. f/2 or L1 media) with a seawater pH of 8 (Table 1).

Table 1: Culture conditions for the main cultivated strains of T. amphioxeia from 2006 to 2020: temperature (T °C), irradiance (I µmol photons m-2 s-1), pH, salinity and light: dark cycle (L: D). n.a. means that the information is not available. Reference (Ref.): (A): Basti et al., 2015c, 2018; Gao et al., 2019; Hongo et al., 2019; Jiang et al., 2018; Kamiyama et al., 2010; Mafra et al., 2014, 2016; Nishitani et al., 2008a, 2008b; Smith et al., 2018; Tong et al., 2015a; Wolny et al., 2020. (B) Gaillard et al., 2020a, 2020b; Gao et al., 2018, 2017; García-Portela et al., 2018; Jia et al., 2019; Raho et al., 2013; Rial et al., 2013; Riobó et al., 2013; Rodríguez et al., 2012; Gaillard et al., 2020c; Séchet et al., 2020. (C) Fiorendino et al., 2020; Garcia-Cuetos et al., 2010; Hattenrath-Lehmann and Gobler, 2015; Nielsen et al., 2012, 2016, 2013, 2020; Riisgaard and Hansen, 2009; Rountos et al., 2019; Rusterholz et al., 2017. (D) Nagai et al., 2013, 2011, 2008; Papiol et al., 2016

I (µmol Culture Strain Origin T (°C) photons m-2 s- pH Salinity L: D (h) Ref. media 1)

AB364287 Japan f/2-f/6 15-22 20-150 n.a. 30-32 12:12-14:10 (A)

f/2-f/6 or L1- AND-A0710 Spain 15-25 50-150 n.a. 32-35 12:12-14:10 (B) L1/20

K-0434 Denmark f/2 or L1 15-21 70-130 8-8.2 25-32 12:12-16:8 (C)

TA-INO01 Japan f/2 18 70-150 n.a. 30 12:12 (D)

Chapter 1: Literature review

I-4-2) Culture of M. rubrum

Mesodinium rubrum is the only food source cultivated for Dinophysis (appendices Table 35). The strains mainly cultured and used (> 75%) were isolated from Japanese (AB364286), Spanish (AND-A0711), Antarctic (CCMP2563) and Danish (MBL-DK2009) waters and they display an important size variation (3.3-fold higher the Japanese compared to the Spanish strains), which could impact growth, ingestion rate and biomass of Dinophysis (Smith et al., 2018) (Table 2). Abiotic conditions are similar to the ones used for T. amphioxeia cultures (described above), but a common practice is to dilute the culture media to control T. amphioxeia outgrowth when fed to M. rubrum. Indeed, the cryptophyte grows faster than the ciliate (ca. 2-fold) (Fiorendino et al., 2020; Gaillard et al., 2020a) and would be favored in rich culture media. Moreover, T. amphioxeia is less affected by elevated pH than M. rubrum (Smith and Hansen, 2007). To further avoid the cryptophyte taking over, the predator: prey ratio tends to be balanced (i.e. usually 1: 1).

Table 2: Culture conditions for the main cultivated strains of M. rubrum from 2006 to 2020: temperature (T °C), irradiance (I µmol photons m-2 s-1), pH, salinity, light: dark cycle (L: D), predator: prey ratio and nutrition frequency. n.a. means that the information is not available. Reference (Ref.): (A) Basti et al., 2018, 2015c, 2015a; Gao et al., 2019, 2017; Hongo et al., 2019; Jiang et al., 2018; Kamiyama et al., 2010; Mafra et al., 2016; Nishitani et al., 2008b, 2008a; Smith et al., 2018; Tong et al., 2015; Wolny et al., 2020. (B) Gaillard et al., 2020a; Gao et al., 2017; García-portela et al., 2020; García-Portela et al., 2019, 2018; Jia et al., 2019; Raho et al., 2013; Rial et al., 2013; Riobó et al., 2013; Rodríguez et al., 2012; Smith et al., 2018. (C) Fux et al., 2011; Hackett et al., 2009; Smith et al., 2018; Tong et al., 2010, 2011, 2015b. (D) Fiorendino et al., 2020; Gaillard et al., 2020b, 2020c; Séchet et al., 2020; Garcia-Cuetos et al., 2010; Hansen et al., 2016; Hattenrath-Lehmann and Gobler, 2015; Nielsen et al., 2012, 2016, 2013, 2020; Ojamäe et al., 2016; Riisgaard and Hansen, 2009; Rountos et al., 2019; Rusterholz et al., 2017

I (µmol preda Prey Culture T photon Salinit L: D Strain Origin pH tor: feeding Ref. media (°C) s m-2 s- y (h) prey frequency 1) once/week- AB36428 12: 12- 1: 1- Japan f/2-f/12 15-20 20-150 n.a. 30-32 every two (A) 6 14: 10 1: 10 weeks AND- f/2-f/6 or 12: 12- once/week- Spain 15-25 50-270 8 30-35 1: 1 (B) A0711 L1- 16: 8 thrice/week

Chapter 1: Literature review

L1/20 or K or seawater CCMP25 1: 1- Antarctica f/2 3-6 50-100 n.a. n.a. 14: 10 n.a. (C) 63 3: 1 f/2 or L1 Twice/wee MBL- 8- 12: 12- 1: 1- Denmark or 15-20 70-130 25-32 k- (D) DK2009 8.2 16: 8 1: 10 seawater thrice/week

I-4-3) Culture of Dinophysis

Amongst the ten species of Dinophysis and two of Phalacroma found to contain toxins, eight have been successfully cultivated. The cosmopolitan species D. acuminata represents 45% of the published papers dealing with Dinophysis culture (Table 3) (appendices Table 36). D. acuta (Table 3 and Table 37 in appendices) and D. sacculus (appendices Table 38) represent 18 and 16% of cultivated species, 10% of D. caudata and 10% of D. fortii, D. ovum and D. infundibulus (Table 39). Up to date, there are no reports of established cultures of D. miles, D. mitra and D. rotundata and D. norvegica culture has been established recently in the US (Deeds et al., 2020). Dinophysis species are cultivated in the same conditions as described above, except for the cold-polar water strains of D. acuminata (e.g. DAEP01 strain; Hackett et al. 2009) (Table 3). D. sacculus, which belongs to the D. acuminata-complex has been mainly found in French Atlantic Ocean waters and French and Spanish Mediterranean Sea water.

Similarly to M. rubrum cultures, Dinophysis are cultivated either with classic nutrient-enriched culture media or with diluted culture media in order to prevent outgrowing of potentially remaining cells of T. amphioxeia and M. rubrum, which could be a problem for the experiments. The predator: prey used can be up to 20:1 (M. rubrum: Dinophysis). Using both this ratio and feeding the cultures twice a week leads to a good growth rate and biomass increase of Dinophysis (Table 3).

Since the first successful permanent culture of D. acuminata in 2006, several studies have focused on ecophysiological traits of different species and strains.

Chapter 1: Literature review

Table 3: Example of culture conditions used for D. acuminata and D. acuta from 2006 to 2020: temperature (T °C), irradiance (I µmol photons m-2 s-1), pH, salinity, light: dark cycle (L: D), predator prey ratio and prey feedingfrequency. n.a. means that the information is not available. Reference (Ref.): (A) Garcia-Cuetos et al., 2010; Riisgaard and Hansen, 2009; Rusterholz et al., 2017. (B) Fux et al., 2011; Gao et al., 2019; Hackett et al., 2009; Smith et al., 2018; Tong et al., 2015b, 2011, 2010. (C) Gao et al., 2019, 2017; Jia et al., 2019. (D) García-Portela et al., 2018, 2019, 2020

I (µmol L: pred Prey Culture T Species Strain Origin photons pH Salinity D ator: feeding Ref. media (°C) m-2 s-1) (h) prey frequency 14: Da- 1: 5- 15- 10- DK200 Denmark f/2-f/20 100 8.2 22-30 1: twice/week (A) 21 16: 7 10 10 f/2-f/6 1: 4- DAEP0 3- 14:

USA or 20-300 8 30 1: n.a. (B) 1 15 10 seawater 20 1: 5- DAYS 14:

D. acuminata China f/6 15 50-100 n.a. n.a. 1: twice/week (C) 01 10 10 12: VGO13 L1/20 or 15- 160- 12- 1: Spain 8 32 once/week (D) 49 K 19 270 16: 10 8 14: North 1: 5- DANA 100- 8- 10- Atlantic f/2 15 30-35 1: twice/week (E) -2010 130 8.2 16:

Ocean 10 8

12: D. acuta VGO10 L1/20 or 15- 160- 12- 1: Spain 8 32 once/week (F) 65 K 19 270 16: 10 8

Chapter 1: Literature review

I-5) Ecophysiological experiments with Dinophysis species

In the context of global change, the frequency of Dinophysis occurrences, and its intensity, abundance, diversity and distribution could be modified by eutrophication of coastal areas, by the introduction of new species or the modification of environmental conditions (GEOHAB, 2008; Hallegraeff, 2010, 1993). Such modifications can occur globally (ocean) or regionally (coastal waters) and concern different abiotic and biotic factors such as nutrient composition (Glibert, 2019), grazing, temperature, pH, light availability, stratification and salinity (Fu et al., 2012; Glibert et al., 2014; Wells et al., 2019, 2015). Notably, the effect of global change has been modeled for D. acuminata. A study showed an increasing trendof D. acuminata growth rate (+ 0.01 d-1) and bloom duration (+ 3 weeks) over the last 30 years in North Atlantic and Pacific regions when using models based on sea surface temperatures and growth rate of D. acuminata in cultures (Gobler et al., 2017).

Ecophysiological experiments performed with different species of Dinophysis in culture, from 2006 to 2020, are listed in Tables 4-10. Experiments have been carried out to study the effect of different (i) abiotic factors, such as temperature (Table 4), irradiance (intensity and quality) (Table 5), salinity (Table 6) and nutrient availability (Table 7); (ii) biotic factors, such as prey concentration and type (Table 9) on (iii) biological processes such as growth phase and cell cycle (Table 10) and (iv) origin of the species/strains of Dinophysis (Table 8). The aim of these experiments was to study how these factors affected the growth rate, photosynthetic activity (e.g. maximum quantum yield of the photosystem II, Fv/Fm), uptake (e.g. prey and/or C) and toxin production of Dinophysis species

I-5-1) Temperature

Global warming affects both the atmosphere and the ocean. Indeed, over the period 1971-2010, the temperature of the upper 75 m of the global ocean increased by 0.33 °C (0.11 °C per decade, 99- 100% of probability; IPCC 2013). Temperature is a key factor affecting several physiological parameters of microalgae (both HAB and non-HAB species) such as their growth rate, photosynthesis and cellular toxin concentrations as well as their seasonal patterns (e.g. bloom onset, duration) (Fiorendino et al., 2020; Gobler et al., 2017; Wells et al., 2019, 2015).

Chapter 1: Literature review

At least 6 laboratory experiments have assessed the effect of temperature on D. acuminata, D. caudata, D. tripos and D. ovum, especially from Japan (Basti et al., 2015c, 2018; Kamiyama et al., 2010), on growth and cellular toxin content (Table 4). According to their results, temperature above 18-20 °C led to increased growth rate and cellular concentration (cells mL-1) all cases except for D. tripos (Rodríguez et al., 2012), while high temperatures (i.e. > 29 °C) to decreased growth of Japanese D. acuminata and D. caudata (Basti et al., 2015c, 2018) (Table 4). Kamiyama et al. (2010) observed higher intracellular content of PTX2 in D. acuminata at 10 °C and similarly, Rodríguez et al. (2012) observed a higher PTX2 content in D. tripos at the lowest tested temperature (15 °C). However, some other studies showed balance intracellular toxin content, but an increase of toxin production linked to the growth rate (Basti et al., 2015c, 2018) (Table 4).

Table 4: In vitro effect of temperature on Dinophysis growth rate, cellular concentration and toxin contents. Okadaic acid, OA; DTX1, dinophysistoxin 1; PTX2, pectenotoxin 2

Temperature Species/Strain Origin Response(s) Reference (°C)   µ and cell concentration (Kamiyama et al., 2010) with  temperature D. acuminata Japan 10, 14, 18, 22  = OA and DTX1intra.   PTX2 intra. at 10   µ and cell concentration (Gao et al., 2017) D. acuminata with  temperature China 10, 15, 20 DAYS01  = OA, DTX1 and PTX2

intra.   µ and cell concentration at (Basti et al., 2018) D. acuminata 5, 8, 11, 14, 17, 20-26,  at 29-32 Japan DA0810HAR03 20, 23, 26, 29, 32   OA, DTX1 and PTX2

production at 17-29 D. acuminata   µ with  temperature, USA 12, 18, 24, 27 (Fiorendino et al., 2020) DAVA01 maximum at 24 D. ovum

DoSS3195 D. tripos  = µ Spain 15,19,25 (Rodríguez et al., 2012) VGO1062  + PTX2 at 15

Chapter 1: Literature review

  µ and cell concentration (Basti et al., 2015c) with  temperature,  at 32.5 15, 18, 21, 24, D. caudata Japan  Correlation PTX2 27, 30, 32.5 production and growth, except at 32.5

I-5-2) Irradiance

Similar to temperature, irradiance was found to affect phytoplankton growth, photosynthesis and toxin production of several HAB species (Fu et al., 2012). The hindcast showed an increase of solar irradiance and a change in solar UV radiation (IPCC 2013). However, trends depend on the scale (e.g. ocean vs. coastal waters or mixed vs. stratified layers) and latitude (e.g. cloud cover in high latitude vs. low latitude) of the study. Indeed, phytoplankton in coastal waters in some areas may receive less irradiance due to increased turbidity (phenomenon of “brownification”, see references in Wells et al. (2015)), while phytoplankton of more mixed layers will be exposed to higher light intensity and UV (Wells et al., 2019), which may be linked to a reduced cloud cover between 50° North and South (Wells et al., 2015).

The effects of light intensity (and to a lesser extent, quality) on D. acuminata and D. acuta and to a lesser extent D. caudata and D. ovum (Table 5) have been studied. Most of the studies shown an increased growth rate with increasing light intensity (García-Portela et al., 2018b; Kim et al., 2008; Nielsen et al., 2012, 2013; Rial et al., 2013), except for the Japanese strain of D. acuminata (DA0810HAR03) (Tong et al., 2011) for which growth was was not affected by light and D. ovum from Gulf of Mexico, which did not grow at light intensities > 100 µ mol photons m-2 s-1 (Fiorendino et al., 2020) (Table 5). Contrasting results regarding toxin production have been observed between studies and species/strains. Indeed, while Tong et al. (2011) and Nielsen et al. (2012, 2013) observed similar toxin production at different light intensities in two strains of D. acuminata, García-Portela et al. (2018b) showed a higher toxin production at 370 µmol photons m-2 s-1 (compared to 10) in D. acuminata (OA) and D. acuta (OA, DTX2 and PTX2), with strains of both species from Spain (Table 5).

Chapter 1: Literature review

Pigment concentrations were constant for D. acuminata and D. acuta under low light and high light conditions (García-Portela et al., 2018b), while photosynthetic parameters changed, a fact which suggested some kind of photoacclimatation of Dinophysis (Table 5).

However, Hansen et al. (2016) showed a decrease in chlorophyll a and increase in carotenoid contents (i.e. photoprotective pigments) (Roy et al., 2011) under high light for a strain of D. acuta without modification of the relative electron transport rate (rETR). Moreover, Rusterholz et al. (2017) observed by light microscopy divisions and changes in morphologies of D. acuminata and D. acuta chloroplasts. These observations suggested these two species of Dinophysiswere able to perform photoregulation (Table 5).

Table 5: In vitro effect of irradiance (intensity and quality) on Dinophysis growth rate, cellular concentration, cellular toxin content and photosynthetic parameters. Maximum quantum yield of the photosystem II, Fv/Fm; Non photochemical quenching, NPQ. Photochemical yield of the photosystem II, ΦPSII; Relative electron transport rate, rETR; Okadaic acid, OA; dinophysistoxin 2, DTX2; pectenotoxin 2, PTX2; Fv/Fm, ΦPSII and rETR are combined as photosynthetic activity

Species/Strain Origin µmol photons m-2 s-1 Response(s) Reference

  µ with  light intensity D. acuminata Republic of 10, 60, 120 (Kim et al., 2008) DA-MAL01 Korea   ingestion rate with  light intensity

D. acuminata  = µ Japan 0, 65, 145, 284, 302 (Tong et al., 2011) DA0810HAR03  = OA, DTX1 and PTX2

  µ with  light intensity

 = PTX2 (Nielsen et al., D. acuminata Denmark 7, 15, 30, 150 2012)   photosynthesis  light intensity

  division and  volume, D. acuminata (Rusterholz et al., Denmark 25, 50, 75, 100 numbers and centers of Da-DK2007 2017) kleptochloroplasts at 75-100

  µ and cell concentration at 370 D. acuminata 10, 370 (García-Portela et Spain   ingestion rate at 370 VGO1349 al., 2018b)  = pigments

 = rETR,  ΦPSII at 370

Chapter 1: Literature review

  OA at 370 intra.

White: 40, 75, 200,   µ at green light at 40 650  rETR, ΦPSII and F /F at Blue and green: 10, v m 650 40

D. acuminata 50, 100, 200, 300,  growth at 50-100 (Fiorendino et al., USA DAVA01 400 2020)

  µ at 200 D. caudata Spain 70, 200 (Rial et al., 2013) VGO1064   chlorophyll a at 200 and = carotenoid ratios

North D. acuta   µ at 7-15 (Hansen et al., Atlantic 7, 15, 30, 130 DANA-2010 2013) Ocean   photosynthesis at 7-15

D. acuta   µ at 200 VGO1065 Spain 70, 200 (Rial et al., 2013)  = carotenoid ratios at 200

  cell concentration at 100 North D. acuta   chlorophyll a and F /F (Hansen et al., Atlantic 25, 50, 75, 100 v m DANA-2010 and  alloxanthin at 100 2016) Ocean  = rETR

North   division and  volume, D. acuta (Rusterholz et al., Atlantic 25, 50, 75, 100 numbers and centers of DANA-2010 2017) Ocean kleptochloroplasts at 75-100

  µ and cell concentration at 370

D. acuta   ingestion rate at 370 10, 370 (García-Portela et VGO1065 Spain  = pigments al., 2018b)  =  rETR,  ΦPSII at 370

  OA, DTX2 and PTX2 at 370 intra.

  µ at green and blue light at White: 40, 75, 200, 40. 650 Blue and green: 10,  rETR, ΦPSII and Fv/Fm at 40 650

Chapter 1: Literature review

D. ovum 50, 100, 200, 300,  growth at 50-100 (Fiorendino et al., DoSS3195 USA 400 2020)

I-5-3) Salinity

In the future ocean, salinity may increase by 0.3 in temperate Atlantic Ocean and up to 0.4 in the Mediterranean Sea (Tester et al., 2020). These modifications could be more important in coastal areas, such as bays, lagoons and estuaries, due to run-off waters, precipitation, evaporation and extreme events related to global changes (IPCC, 2013; Wells et al., 2019). Several species of the genus Dinophysis, such as D. sacculus and D. acuminata have been found to grow in these coastal areas (see section “I-10) DSTs distribution”) and thus subjected to quick salinity fluctuations.

To date, only one recent study has focused on the effect of salinity acclimation (among other factors, i.e. irradiance and temperature) (Fiorendino et al., 2020) (Table 6). These authors showed that both American strains of D. acuminata and D. ovum grown with salinities ranging between 22 and 34, with an optimum at 22, seemed to have a good tolerance to salinity changes (Table 6).

Table 6: In vitro effect of salinity on Dinophysis growth rate

Species/Strain Origin Salinity Response(s) Reference D. acuminata  Growth between 22-34 with optimum at (Fiorendino et al., USA 22, 26, 30, 34 DAVA01 22 2020) D. ovum

DoSS319

I-5-4) Nutrients

Nutrient concentration, ratio and composition, especially in coastal waters, can be modified by different factors, e.g. modification of precipitation regime, increased stratification and extreme events impact and/or eutrophication (Fu et al. 2012, IPCC 2013, Glibert 2019, Wells et al. 2019). Growth, and ultimately blooms, of coastal species mainly depend on nutrients from water fluxes

Chapter 1: Literature review

(i.e. upwelling areas or run-off of nutrient rich waters). The growth of dinoflagellates, and especially mixotrophic species, is favored in stratified and oligotrophic waters (Fu et al., 2012; Hansen et al., 2019; Wells et al., 2019).

Nutrient uptake and preference of D. acuminata and to a lesser extent D. acuta, have been well documented based on both laboratory (Table 7) and field studies (not presented here) (e.g. Hattenrath-Lehmann et al., 2015; Seeyave et al., 2009).

Some studies showed that D. acuminata (from Spain and USA) was able to uptake ammonium (i.e. inorganic nitrogen), urea and/or glutamine (i.e. organic nitrogen) (García-Portela et al., 2020; Hattenrath-Lehmann and Gobler, 2015; Hattenrath-Lehmann et al., 2015) and to a lesser extent nitrate (Hattenrath-Lehmann et al., 2015). These observations thus suggested that D. acuminata and D. acuta had a preference for uptake of regenerated inorganic and organic nitrogen, and the uptake was enhanced by prey availability (García-Portela et al., 2020) (Table 7). Tong et al. (2015a) did not observe uptake of nitrate, ammonium and phosphate with an American strain of D. acuminata (DAMV01). The absence of uptake of nitrate observed in García-Portela et al. (2020) have been suggested to be related to the low number of nitrate-reductase transporters in the membrane of D. acuminata.

Toxin contents per cell (OA, DTX1 and PTX2) of two American strains showed a positive correlation with glutamine (Hattenrath-Lehmann et al., 2015) (Table 7).

Table 7: In vitro effect of nutrients on Dinophysis growth rate and toxin contents. Okadaic acid, OA; DTX2, dinophysistoxin 2; PTX2, pectenotoxin 2

Species/Strain Origin Range(s) Response(s) Reference Nitrate, ammonium, urea,   µ and cell concentration with nitrate, (Hattenrath- D. acuminata USA glutamine, phosphate and ammonium   OA, DTX1 Lehmann et al., phosphate, and PTX2 intra. with glutamine 2015) vitamin B12 Ammonium,   µ with ammonium, glutamine and (Hattenrath- nitrate, organic nitrogen from sewage with and D. acuminata USA Lehmann and glutamine, without prey Gobler, 2015) vitamin B12,   µ with nitrate with prey

Chapter 1: Literature review

organic nitrogen  Uptake of ammonium and urea and less from sewage nitrate

Nitrate,  = µ, no uptake of nitrate, ammonium, D. acuminata (Tong et al., USA ammonium, phosphate DAMV01 2015a) phosphate  = OA, DTX1 and PTX2

D. acuminata Ammonium, (García-Portela Spain  Uptake of ammonium and urea, no nitrate. VGO1349 nitrate, urea et al., 2020) D. acuta Spain VGO1065

I-5-5) Geographical origin

Intraspecific variations in growth rates and toxin profiles of Dinophysis were highlighted in two laboratory studies (Fux et al., 2011; Tong et al., 2015b) (Table 8). Such variation in toxin profiles was also reported in the section “I-10) DSTs distribution” Figure 8 A-C and Figure 14 D for instance between French and American of D. acuminata (OA and PTX2 producers vs. OA producers only). These intraspecific differences could be explained by different chemotype and should be further explored in other species of the genus.

Table 8: Effects of geographical origin of Dinophysis and/or its food chain on growth rate and toxin contents

Species/Strains Factor(s) Range(s) Response(s) Reference D. acuminata DAEP01, From Canada,  Different toxin profiles depending on the DAMV01, Geographical north USA, (Fux et al., geographical origin DABOF01, origin south USA 2011)  Similarities in isolates from North America DARE01 and and Chile DAPA01 D. acuminata From Canada DAEP0,  Similarities in growth rates and toxin profiles Geographical and north (Tong et al., DAMV01, and contents between isolates from the same origin USA and 2015b) DABOF01 and origins south USA DAPA01

Chapter 1: Literature review

I-5-6) Prey

As mentioned in section “I-3-3) Mixotrophy of Dinophysis”, Dinophysis is a mixotroph, which specifically feeds on M. rubrum. Thus, Dinophysis, which is usually a rare genus among the marine phytoplankton community, is classified as a “plastidic specialist non-constitutive mixotroph” (Mitra et al., 2016).

Several studies (Gao et al., 2019; Hattenrath-Lehmann and Gobler, 2015; Kim et al., 2008; Riisgaard and Hansen, 2009; Tong et al., 2015a) showed a positive effect of the prey concentration on Dinophysis growth rate (Table 9). Moreover, the origin of M. rubrum affected growth rate (Smith et al., 2018) and metabolite contents (García-Portela et al., 2018). Furthermore, when M. rubrum growth rate increased, growth and ingestion rate of D. acuminata increased (Riisgaard and Hansen, 2009) (Table 9).

While prey depletion was related to increased toxin concentration per cell in cultures of Spanish D. acuminata and D. acuta (García-Portela et al., 2018), increased prey concentration led to increased toxin contents in American strains of D. acuminata (Tong et al., 2015a) (Table 9).

Table 9: In vitro effect of prey type and concentration on Dinophysis growth rate and toxin content. Okadaic acid, OA; DTX1, dinophysistoxin 1; DTX2, dinophysistoxin 2; PTX2, pectenotoxin 2

Species/Strain Origin Range(s) Response(s) Reference D: M: 5: 10; 10: 50; 20:1 00; 50: D. acuminata Republic of 500; 100: 1000; (Kim et al.,   µ with  prey concentrations DA-MAL01 Korea 300: 3000; 500: 2008) 5000; 1000: 10000 D: M: 40: 1500,   µ with  prey concentrations 60: 1900, 34:   µ and ingestion rate with  growth rate D. acuminata 270, 4: 600, 10: (Riisgaard and Denmark of M. rubrum Da-DK2007 600, 35: 600, 4: Hansen, 2009)  C uptake is explained by food at high prey 100, 10: 100, 35: concentration 100 and 20: 20

Chapter 1: Literature review

(Hattenrath- Ratio D: M: 1: 2, D. acuminata USA   µ with  prey concentrations Lehmann and 1: 5 and 1: 10 Gobler, 2015)   µ with  prey concentrations D. acuminata   OA, DTX1 and PTX2 with  prey (Tong et al., USA DAMV01 concentrations 2015a)

M. rubrum and T. amphioxeia D. acuminata origins (+  µ and cell concentration impacted by M. (Smith et al., USA DAEP01 biovolume and rubrum origin and nutritional content 2018) nutritional content)   µ and cell concentration with  prey D. acuminata (Gao et al., USA lysate DAEP01 2019)   OA and DTX1 with prey lysate   OA and DTX2 prey-limited D. acuminata Prey origin and  Prey preference (García-Portela Spain VGO1349 food quantity  Difference in metabolomic profiles with et al., 2018a) prey origin and quantity   OA prey-limited D. acuta VGO  Prey preference Spain 1065  Difference in metabolomic profiles with prey origin and quantity D. acuminata (García-Portela Spain Limitation or not   of nitrogen uptake when well-fed VGO1349 et al., 2020) D. acuta VGO Spain 1065

I-5-7) Growth phase and cell cycle

Most HAB species seem to contain higher cellular concentrations of toxins at the end of the exponential growth phase and during the stationary phase than during the exponential growth phase, with a release when cultures are in senescent phase (e.g. field population of D. acuta (Pizarro et al., 2009) or culture of Azadinium spinosum (Jauffrais et al., 2013)).

Chapter 1: Literature review

Recently, Jia et al. (2019) showed that a significant amount of OA, DTX1 and PTX2 (e.g. from 30 to 50 pg cells-1 in 8 h) was produced by D. acuminata during a 24 h cycle (Table 10).

Table 10: Effects of the population growth phase and cell cycle stages on Dinophysis growth rate and toxin content. Okadaic acid, OA; DTX2, dinophysistoxin 2; PTX2, pectenotoxin 2

Growth /cell Species/Strain Origin Response(s) Reference cycle GP Exponential, D. acuminata   toxins by mid-plateau phase (Tong et al., Japan stationary and DA0810HAR03  of OA and DTX2 when culture aged 2011) aged culture D. fortii

DF0705TTR24 North Exponential vs. D. acuta DANA-  PTX2 content is linked to growth rate (Nielsen et al., Atlantic Stationary/declin 2010  toxins when cultures declined 2013) Ocean ed  toxin in G1 phase,  at the beginning of D. acuminata Cell G1, S, G2 and M (Jia et al., the S phase,  at the end of the S phase and DAYS01 cycle phases 2019)  at M phase

I-5-8) Conclusion of the ecophysiological experiments on Dinophysis

Overall, the increase of temperature, irradiance and regenerated form of nitrogen (i.e. abiotic factors) led to increased growth rate of Dinophysis species without any clear effect on cellular toxin contents or toxin production, as there are still a lot of discrepancies between studies. The differences may be explained by intra and/or interspecific variations, the nutritional state of Dinophysis (e.g. well-fed or not) and even the nutritional state of their prey.

Nonetheless, there are still too few ecophysiological experiments on Dinophysis. This lack of studies can be explained by the difficulties and time-consuming laboratory work to cultivate, maintain and conduct in vitro experiments with Dinophysis.

Chapter 1: Literature review

I-6) Dinophysis Toxins

Species of the genus Dinophysis can synthesize two types of lipophilic toxins, the okadaic acid (OA) and its analogues the dinophysistoxins (DTXs) (named OA-group toxins) (Figure 3), which are responsible for DSP, and the non-diarrheic pectenotoxins (PTXs) (Figure 4).

I-6-1) OA-group toxins

Toxins of the okadaic acid-group (OA-group toxins), which includes OA and DTXs, are linear polyethers. Over the 93 analogues of the esters of the OA-group toxins (Pan et al., 2017), only OA, DTX1, DTX2, DTX3 and diol esters have been found in Dinophysis (Reguera et al. 2014). The other analogues are products of transformation in shellfish (see examples in Figure 3) or come from toxic species of Prorocentrum (Pan et al., 2017), as hybrid diol-fatty acid esters (Torgersen et al., 2008).

The polyether OA was first isolated from the sponge Halichondria okadai, from which it takes its name (Tachibana et al., 1981). Dinophysistoxin 1 was isolated from mussels (Murata et al., 1982) and DTX2 was initially isolated from mussels from Ireland (Hu et al., 1992a) and simultaneously also identified in two Prorocentrum species (Hu et al., 1992b). Other DTX analogues were also isolated from Prorocentrum spp. (Suzuki, 2014).

I-6-1-1) Chemical structure of OA-group toxins

The analogues DTX1 and DTX2 differ by the addition or the position of one methyl group in the molecule, respectively (Figure 3). The group DTX3 (okadaates), acyl derivatives of OA, are the transformation products of OA in shellfish (Figure 3) (Suzuki and Mitsuya, 2001). Dinophysistoxin 4 and DTX5 are diol ester precursors (Figure 3).

Chapter 1: Literature review

A B C

Analogue R1 R2 R3 R4 R5 R6

OA CH3 H H H H

DTX1 CH3 CH3 H H H

DTXDTx22 H H CH3 H H

DTXDTx33 H or CH3 CH3 H Acyl H

Diol esters CH3 CH3 H H A OH

Sulfated DTX4/5 CH3 CH3 H H A B

DTX6 CH3 CH3 H H C

Figure 3: Chemical structure of OA and its analogues. Modified from Boundy et al. (2020)

I-6-1-2) Toxicity of OA-group toxins by intraperitoneal injection and oral

administration in rodents

Okadaic acid and DTX1 showed similar toxicity by intraperitoneal injections in mice and rats, leading to intestine and liver injuries (Berven et al., 2001; Ito and Terao, 1994; Terao et al., 1993) while DTX2 was, according to the assays, about 40% to 50% less toxic than OA and DTX1 (Aune et al., 2007). Oral administration of OA, DTX1 and DTX3 led to similar injuries (Terao et al., 1993; Tubaro et al., 2003).

Lethal toxicity of OA-group toxins is higher by oral administration than by intraperitoneal injection, e.g. 225 µg/kg vs. 2000 µg/kg body weight (BW) for intraperitoneal injection and oral administration of OA, respectively (Tubaro et al., 2003).

The toxicity of OA esters (Figure 3) and DTX esters is similar to the toxicity of the parent compounds (Torgersen et al., 2005).

Chapter 1: Literature review

I-6-1-3) Elucidation of action mode of OA-group toxins

Okadaic acid and DTXs are selective inhibitors of eukaryotic serine/threonine protein phosphatases (PP) (Bialojan and Takai, 1988; Valdiglesias et al., 2013), especially protein phosphatase 1 (PP1) and protein phosphatase 2A (PP2A) (Cruz et al., 2007; Fernandez et al., 2002). Thus, they interfere with proteins that control sodium secretion in intestinal cells that ultimately leads to diarrheic symptoms (Cohen et al., 1990).

Inhibition of these PP also induced diarrhea by degenerative changes in the absorptive epithelium of the intestine of mammals, including humans (Terao et al., 1986; Tripuraneni et al., 1997). The inhibition of PP2A impeded synthesis of an essential protein involved in initiation of DNA replication (Chou et al., 2002), PP1 inhibition promoted tumor and induced apoptosis in mammals (Long et al., 2002) and inhibition of both proteins could impede essential metabolic processes (e.g. growth, division) (Daranas et al., 2007).

Tumor promoting role of OA and DTX had been shown to be related to DNA adduct formation on hamster fibroblasts by Fessard et al. (1996), and is mainly due to the fact that they induced the increase of cytokine synthesis and secretion, especially TNF-α (del Campo et al., 2017; Fujiki et al., 2013). Indeed, TNF-α-deficient mice did not develop tumors when exposed to OA (Suganuma et al., 1999), contrary to regular mice (Feng et al., 2006; Fujiki and Suganuma, 1999; Suganuma et al., 1999; Thompson et al., 2002), that developed mainly skin cancers. For instance, application of pmol of DTX1 and OA on mouse ear skin induced irritation while nmol of these same toxins applied twice a week on back skin for 30 weeks induced tumor in ≥ 80% of the tested animals (Fujiki et al., 1988). Daily oral ingestion of 12 nmol of OA by rat for 1 year then 24 nmol for 4 months induced neoplasia in the stomach (i.e. hyperplasia and carcinoma) (Suganuma et al., 1992).

Immunotoxicity was observed in mice macrophages and dendritic cells exposed to around 100 nM of DTX1, stimulated by the secretion of several inflammatory factors (e.g. TNF-α) (del Campo et al., 2017)

Cytotoxic effects (e.g. cell loss, activation of apoptotic proteins such as caspases) were reported by Ferron et al. (2016) in human hepatocyte cell lines exposed to 4 to 42 nM of OA, DTX1,2 and PTX2. A similar cytotoxic effect was observed with OA and DTX1 and 2 on epithelial colorectal adenocarcinoma cells (Caco-2) (Ferron et al., 2014). Several authors highlighted the higher toxic 70

Chapter 1: Literature review effect of DTX1 toward cell line compared to other toxins of OA-group (del Campo et al., 2017; Ferron et al., 2014). In addition, a recent study by Alarcan et al. (2019) showed the cytotoxic effect of 75 nM of OA on Caco-2 cells, while 19 nM of the same toxin induced a release of interleukin 8 (cytokine) and caused genotoxicity by DNA strand breaks. Okadaic acid has also been shown to induce changes in F-actin filaments in several models of cell lines (see Vale and Botana (2008). Indeed, µM of OA led to e.g. cytoskeletal disruption after inhibition of PP in rat hepatocytes (Macías-Silva and García-Sáinz, 1994), decreased concentration of F-actin and altered microtubules in human platelets (Yano et al., 1995), and nM of OA decreased F-actin levels in human neuroblastoma (IC50 1 h of 100 nM) (Leira et al., 2001).

I-6-2) PTXs

Pectenotoxins are macrocyclic polyether lactones that contain more than 20 analogues (Boundy et al., 2020), mainly resulting from metabolic transformation of PTX2 in shellfish (Figure 4) (Allingham et al., 2007; Blanco, 2018; Miles et al., 2004b). In Dinophysis, only PTX2 (and sometimes seco acid products; Fernández Puente et al., 2004), PTX11 (Suzuki et al., 2004), PTX 12 (Miles et al., 2004b) and PTX13 and PTX14 (Miles et al., 2006a) have been found.

Pectenotoxin 1 and 2 were isolated from the scallop Patinopecten yessoensis and were named according to the genus name (Yasumoto et al., 1984, 1985).

I-6-2-1) Chemical structure of PTXs

In shellfish, PTX2 is metabolized in compounds such as pectenotoxin 2-seco acid (PTX2-sa), 7- epi-pectenotoxin 2 seco acid (7-epi-PTX2-sa), PTX1, PTX3 or PTX6 (Miles et al., 2006b; Suzuki et al., 2005). They differ from each other by the degree of oxidation of carbon C-43 (i.e. R3 Figure 4), arrangement or epimerization of the spiroketal ring system and the opening of the lactone ring C-1 - C-33 (Figure 4) (Allingham et al., 2007; Reguera et al., 2014).

Chapter 1: Literature review

Analogue Figure R1 R2 R3 C7

PTX1 A H H CH2OH R

PTX2 A H H CH3 R

PTX2b A H H CH3 S

PTX2c A H H CH3 S PTX3 A H H CHO R

PTX4 A H H CH2OH S PTX6 A H H COOH R PTX7 A H H COOH S

PTX8 B H H CH2OH S PTX9 B H H COOH S A PTX11 A OH H CH3 R PTX11b A OH H CH3 S

PTX11c B OH H CH3 S

PTX13 A H OH CH3 R PTX2sa C H H H R 7-epi -PTX2sa C H H H S 37-O-acyl PTX2sa C acyl H H R 33-O-acyl PTX2sa C H acyl H R 11-O-acyl PTX2sa C H H acyl R

B PTX12

C PTX14

Figure 4: Chemical structure of PTXs and their analogues. Modified from Boundy et al. (2020)

I-6-2-2) Toxicity of PTXs by intraperitoneal injection and oral administration

in rodents

Hepatoxicity was shown for PTX1, PTX2 and PTX6 by intraperitoneal injections in mice and rats, while no toxic effect was observed for PTX2-sa and 7-epi-PTX2-sa (Dominguez et al., 2010; Ito et al., 2008; Terao et al., 1986).

Chapter 1: Literature review

Intraperitoneal injections of PTX2 led to LD50 of 250 µg/kg BW while >5000 µg/kg BW of PTX2- sa did not induce death, but neither PTX2 nor PTX2-sa are toxic by oral administration (Miles et al., 2004a).

A study from 1988 demonstrated that oral ingestion of PTX2 induced diarrhea in mice (Ishige et al., 1988), whereas no effect was observed in recent studies (Ito et al., 2008; Miles et al., 2004a). Controversy exists about the degree of toxicity of PTXs, but it is now admitted that PTXs cannot be considered as DSTs (Boundy et al., 2020; Miles et al., 2004a).

I-6-2-3) Elucidation of action mode of PTXs

While PTXs are not considered as diarrheic toxins, their mode of action has been well described. Pectenotoxin 1 (58 to 582 nM - 0.5 to 2 h) caused disruption of actin bundles stress fibers, loss of arrangement of microtubules and accumulation of actin in chicks liver cells(Zhou et al., 1994). Pectenotoxin 2 and PTX6 depolymerized actin and disrupted F-actin in rat aorta cells (300 nM - ≤ 1.5 h (English abstract and figures in Hori et al., 1999) and human neuroblastoma cells, respectively (700 nM - 24 h (Leira et al., 2002). Also, sequestration of monomeric actin filaments with 20 nM of PTX2 was observed on actin by Spector et al. (1999). Ares et al. (2005) showed a decrease in F-actin filament in rabbit enterocytes exposed 4 h to 1000 nM of PTX6.

This interaction with actin was observed and well described by Allingham et al. (2007). Indeed, the observation of X-ray crystal structure of PTX2 bound to actin showed that the toxin inhibited actin depolymerization by capping the barbed-end of F- and G-actin (Allingham et al., 2007).

Apoptosis (caspase activation) was triggered by PTX2 in mice oocytes exposed 24-48 h to 116 nM (Chae et al., 2005). Fladmark et al. (1998) also observed apoptosis induced by 2000 nM of PTX1 in rat and salmon hepatocytes after 1 h of exposure.

Cytotoxicity towards several human cancer cell lines, including lung, colon and breast, was highlighted after exposition with 8 to 800 nM of PTX2 (Jung et al., 1995). In a recent study, Hori et al. (2018) observed inhibition of smooth muscle of rat contractions and inhibition of velocity/degree of actin polymerization after 2 h exposure at 300 nM of PTX2.

Genotoxicity was shown by DNA strand breaks in Caco-2 cells when exposed to 200 nM of PTX2 (Alarcan et al., 2019). Interestingly in this last publication, the authors reported an antagonistic 73

Chapter 1: Literature review effect of the combination of OA and PTX2 at low concentration while high concentration of both compounds resulted in additive and synergistic effects (Alarcan et al., 2019).

I-6-3) Effects of OA-group toxins and PTXs on humans

The lipophilic and diarrheic toxins from the OA-group toxins (mainly synthesized by Dinophysis and to a lesser extent by benthic species of Prorocentrum (Ehrenberg, 1832) are responsible for the DSP syndrome in the human consumers of filter-feeding bivalves contaminated with them. Indeed, OA-group toxins can be accumulated in digestive glands of mollusks (Van Dolah, 2000), as well as the non-diarrheic PTXs, and consumption of such mollusks lead to DSP symptoms: diarrhea, nausea and abdominal pain 3 to 12 h after ingestion (Yasumoto et al., 1978). The first listed events of “DSP” happened in the Netherlands in 1961 and in Chile in 1970 (Guzmán and Campodonico, 1975; Korringa and Roskam, 1961). The intoxication was named DSP after an outbreak in Japan in the late 1970s due to consumption of mussels and scallops contaminated by D. fortii (Murata et al., 1982; Yasumoto et al., 1978, 1980). The European Council (2004) sets a limitation of 160 µg of OA-group toxins per kilo of meat for bivalves.

In addition to the gastro-intestinal symptoms, little information is available on other toxic effects towards human consumers of bivalves contaminated with DSTs. According to studies of Cordier et al. (2000) and Manerio et al. (2008) high DST levels in contaminated filter-feeding bivalves consumed by humans could potentially increase the risks of digestive cancers (e.g. colorectal cancer). To date, no toxicity of PTX2 towards human consumers has been reported.

I-6-4) OA-group toxin and PTX measurements

The detection of lipophilic toxins was initially performed by mouse bioassays (MBAs) until 2004 (European Food Safety legislation 2004) according to the method of Yasumoto et al. (1984). This method consists of the injection of mollusk extracts (suspected to contain toxins) from the hepatopancreas or the whole body dissolved in a solvent (Yasumoto et al., 1978, 1984) by intraperitoneal injections in at least 3 mice (or rats, rat bioassays (RBAs)). A positive test corresponds to the death of 2 out of 3 mice/rats after 24 h. The MBA and RBA tests are simple and no complex equipment is required (for instance compared to chemical measurements) and the main

Chapter 1: Literature review advantage is the potential detection of unknown toxins due to unusual symptoms. However, these biological assays are not specific: it is not possible to identify the toxin(s) responsible for the effects, unreliable (both false positive and negative results have been reported) and especially they cause animal suffering (Alexander et al., 2008; Hess et al., 2006, 2003; Lawrence et al., 2011). Therefore, non-animal detection methods have been recommended, such as (1) functional methods, (2) immunological methods or (3) chemical analytical methods.

I-6-4-1) Functional methods

Functional methods are based on the selective interaction and recognition of toxins on a cellular component (biological receptor), leading to the transformation of the chemical information of the toxin into a defined effect. For OA-group toxins, the assay consists of the quantification by fluorimetry/colorimetry of the PP2A inhibition or fragmentation (Hess et al., 2006; Simon and Vemoux, 1994). This method is fast, sensitive and the sample processing is easy to operate, but identification of kind of toxins involved is not possible and false positives are easily generated (Alexander et al., 2008; Lawrence et al., 2011).

I-6-4-2) Immunological methods

Similarly, the immunological assays consist of the binding of the toxin to an antibody, e.g. the colorimetric detection after an enzyme-linked immunosorbent assay (ELISA). For OA-group toxins, an ELISA test is available since 1988 (Uda and Yasumoto, 1988), with a poor limit of detection and cross reactivity of the antibody (Hess et al., 2006).

I-6-4-3) Chemical analytical methods

The chemical analytical methods consist of the separation of the toxins in the extract by liquid chromatography (LC), and the detection by UV (LC-UV), fluorescence (LC-FLD) or mass spectrometry (LC-MS).

Nowadays, OA-group toxins are monitored in Europe using instrumental detection methods, i.e. liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS) (EU.Commission,

Chapter 1: Literature review

2011; Hess, 2011), which also allows the detection of the PTXs. Moreover almost all the DSTs are detected, but the detection becomes more complex when samples contain a biological matrix (e.g. matrix effects, increase of noise, ion suppression) (Lawrence et al., 2011; Suzuki and Quilliam, 2011).

In scientific publications, compared to routine monitoring, the detection of DSTs in phytoplankton and shellfish was mostly performed by chemical analytical methods (see Table 15 available online at: https://doi.org/10.17882/76433). Otherwise, studies combined MBA (or RBA) with ELISA or with chemical analytical methods. Only few studies performed only MBA (or RBA) or ELISA analysis, thus the harmonization of method (i.e. the use of LC-MS method) makes comparison between studies more accurate.

I-7) Accumulation of Dinophysis, OA-group toxins and PTXs in marine organisms

Marine organisms can be directly exposed to intracellular or extracellular toxins through the consumption of toxic microalgae (e.g. planktonic organisms, filter-feeding bivalves or planktivorous fish), or indirectly by the consumption of contaminated organisms, which have bioaccumulated, biotransformed or biomagnified toxins.

Many marine organisms have been described as accumulating DSTs, however, very often the effects on these organisms are not assessed. We intend to review here all the organisms or groups, from crustaceans to marine mammals, which have been recorded to accumulate DSTs.

I-7-1) Accumulation in crustaceans, echinoderms, gastropods and cephalopods

Okadaic-acid group toxins have been recorded in small crustacean zooplankton, crabs, echinoderms and gastropods. Indeed, a DSP outbreak happened in Norway in 2002 after the consumption of contaminated brown crabs (Cancer pagarus) by OA-group toxins (i.e. OA, DTX1 and DTX2) (Torgersen et al., 2005). The amount of OA-group toxins found in crabs was below the

Chapter 1: Literature review regulatory limit, but esterified forms of OA-group toxins were also found in crab meats and could be responsible for the observed toxicity. This DSP event may be due to the consumption of contaminated mussels by crabs in the first place (Torgersen et al., 2005).

In 2015, OA, above the regulation limit, and DTX1 were found in two carnivorous species of gastropods (sea snails) from Southern Chile (García et al., 2015). In a massive bloom dominated by the D. acuminata-complex species, Mafra et al. (2019) found OA at a concentration < 160 µg kg-1 of whole body tissue, in small crustacean zooplankton, echinoderms and gastropods. Mafra et al. (2015) reported the presence, below the regulation limit of OA and DTX1 in octopuses which fed contaminated mollusks. This cephalopod is a food item for humans and a top predator marine animal (Mafra et al., 2015).

However, in these studies, the authors only provided information regarding toxin accumulation and did not describe the potential effects on the animals.

I-7-2) Accumulation in filter-feeding bivalves

Toxic phytoplankton can be retained by filter-feeding bivalves by filtration through the gills. This filtration may explain why filter-feeding bivalves are usually contaminated when exposed to HAB of toxic species, especially in coastal areas where shellfish farming activities are located (Bauder et al., 2001; Blanco, 2018; Simões et al., 2015). Accumulation of DSTs has been observed in numerous species of e.g. mussels, oysters, clams and scallops (Reguera et al., 2014). Among them, mussels seem to accumulate more DSTs than other bivalve species, which can be due to their lower rates of metabolization and higher filtration rate (Blanco et al., 2018; Lindegarth et al., 2009; Vale, 2006) (Table 15 available online at: https://doi.org/10.17882/76433). Indeed, since the 1970s, mussels are the bivalves in which DSTs have been mainly found. Thus, Mafra et al. (2019) observed in whole body of mussels (Perna perna) 7693 µg kg-1 of OA, which is the highest concentration ever reported. Thus, mussels are used as sentinel species in monitoring programs (Trainer et al., 2013; Vale, 2004) since they get contaminated with high concentration of toxins earlier than other bivalve shellfish species (Belin et al., 2020)

Chapter 1: Literature review

I-7-3) Accumulation in fish

Accumulation in suspensivorous, as well as predator fish, which are at a higher trophic level, may happen in the environment. Indeed, the presence of OA (44 µg kg-1 of liver and digestive tract tissues) in a filter-feeding fish (Cetengraulis edentulous) and in a mullet (Mugil liza) gizzard (at a concentration closed to the regulation limit) was reported (Mafra et al., 2019, 2014). Moreover, OA was found in the liver of flounders (Platichtys flesus) which had fed blue mussels, at a concentration of 222 µg kg-1 (Sipiä et al., 2000).

These reports suggested that accumulation and contamination of fish with OA and derivatives should be taken into account, as these food items appeared to be vectors of DSP.

I-7-4) Accumulation in turtles

Only one occurrence of OA accumulation in a turtle was reported. Indeed, Landsberg et al. (1999) hypothesized an association between green turtles (Chelonia mydas) fibropapillomatosis in Hawaiian Islands and Prorocentrum spp. (and thus OA). Sampling of the green turtles showed that 2 out of 3 of the animals were infected by fibropapillomatosis, which is a neoplastic disease, characterized by tumor in liver and kidneys (Landsberg et al., 1999). In these islands, benthic species of Prorocentrum genus (producer of OA) were found on seaweed on which the green turtles fed. Moreover, OA was found in turtle tissues by PP inhibition assay, even though this assay is not very specific as microcystins also inhibit PPs.

I-7-5) Accumulation in birds

Accumulation of DSTs in birds has been reported twice in the literature, without direct evidence of pathologies and/or mortalities. During the early 1990s, sea bird mortalities were observed in several taxa, including penguins, in New Zealand. Mortalities co-occurred with several syndromes -NSP, PSP, ASP and DSP- in human consumers and blooms of the associated toxic genus/species, but these events were not directly linked either to HAB toxins, or to DSTs (reviewed in Shumway et al. 2003). Similarly, mortality of sea birds (Black-legged Kittiwakes) contaminated with DSTs (presence in the liver) was observed in UK in late 1990s (Shumway et al., 2003).

Further studies should focus on sea bird mortalities in a context of HAB of Dinophysis species.

Chapter 1: Literature review

I-7-6) Accumulation in marine mammals

Marine mammals are the top predators in marine ecosystems, and thus may accumulate toxins as DSTs. In 2008, an important mortality of bottlenose dolphins (Tursiops truncatus) was observed in coastal water of Texas, Gulf of Mexico (Fire et al., 2011). These stranding dolphins co-occurred with an important bloom of Dinophysis ovum (up to 3.3 million cells L-1) and OA was found in the faeces and the gastric intestinal content of the animals (Fire et al., 2011). The observed mortalities may be linked to a cocktail of toxins, because brevetoxins and domoic acid were also found (Fire et al., 2011).

In another study, sampling of urine, feces, intestinal and stomach contents of live and healthy Peruvian fur seals (Arctocephalus australis) and South American sea lions (Otaria byronia) in South Peru in 2012 revealed the presence of OA (0.5 to 36 ng g-1) in the faeces (Fire et al., 2017). The animals may have been exposed to toxic blooms of Dinophysis species (60,000 to 150,000 cells L-1) weeks to months before sampling (Fire et al., 2017).

Intoxications of marine mammals with DST’s are scarce but acute or chronic effects should be more intensively studied due to a high risk of exposition (Broadwater et al., 2018; Mafra et al., 2019).

I-8) Effects of Dinophysis, OA-group toxins and PTXs on marine organisms

Dinophysis exposure and DST accumulations can have deleterious effects on marine organisms. As mentioned previously, an important mortality of bottlenose dolphins (Tursiops truncatus) was observed on the coasts of Texas, Gulf of Mexico in 2008 (Fire et al., 2011). This is the only report about effects of DST on marine mammals. However, in vitro effects on other phytoplankton species, mollusk bivalves and fish have been reported and are described below.

Chapter 1: Literature review

I-8-1) Allelopathic effects

The release of toxins and/or other bioactive compounds (metabolites) can induce adverse effects on bacteria, grazers and other competitors. These metabolites are defined as allelochemicals (Granéli and Hansen, 2006; Spector, 1984; Stoecker et al., 2006; Tillmann et al., 2007). For instance, allelochemicals with negative allelopathic effects released by species of the genus Alexandrium and Karenia may deter grazers, inhibit growth of other phytoplankton species and thus constitute a competitive advantage for the producer (Long et al., 2018b; Spector, 1984; Targett and Ward, 1991). Allelochemicals such as karlotoxines produced by toxic species of the genus Karlodinium (REF) or unknown bioactive substances produced by Alexandrium (Blossom et al., 2012) have been shown to play a role in prey capture.

Several authors suggested that DSTs could also have an negative allelopathic role (Sugg and VanDolah, 1999; Windust et al., 1996). Growth of the diatom Thalassiosira weissfogii was inhibited when grown in culture media with micromolar concentrations of OA (i.e. 3-6 µM) or DTX1 (3 µM), without significant synergistic or antagonistic effect (Windust et al., 1996, 1997). In contrast, Windust et al. (1996) did not observe any inhibition of growth when the toxic species P. lima (DSTs producer) was exposed to high doses of OA (until 500 µg disk-1 with disk-diffusion bioassay and 12 µM in culture media) and only a higher concentration of OA (i.e. 21 µM) in the culture media inhibited the growth of a non-toxic P. minimum, thus suggesting that this genus harbors specific self-defense adaptations against DSTs. Sugg and VanDolah (1999) observed sensitivity of protein phosphatases (i.e. target of OA) of several phytoplankton species (i.e. Amphidinium klebsii, Coolia monotis, Gambierdiscus toxicus and Ostreopsis lenticularis) when exposed to 1 µM of OA and to culture media containing exudates of a toxic strain of P. lima. However, as the P. lima medium contained only traces of OA (i.e. 5 nM) the authors suggested that the OA concentration was not sufficient to play a role in growth inhibition and that the effect was due to an unknown - different to OA - bioactive compound (Sugg and VanDolah, 1999).

In a recent study of the capture mechanism of M. rubrum by Dinophysis, Mafra et al., (2016) suggested the role of an unknownbioactive compounds in the immobilization of prey cells, different to DSTs, because addition of OA, DTX1 and PTX2 on Mesodinium cultures did not show any effect. Mafra et al., (2016) suggested that eicosapentaenoic acid (EPA), a polyunsaturated fatty acid (PUFA), at a concentration of ca. 77 ng mL-1 in their cultures, could be responsible for the

Chapter 1: Literature review toxicity of Dinophysis to M. rubrum in their experiment. In a previous similar feeding experiment, Ojamäe et al., (2016) hypothesized the role of allelochemicals (and possiblythe DSTs) involved in the loss of cirri and/or secretion of mucus by M. rubrum.

I-8-2) Effects of filter-feeding bivalves exposure to Dinophysis or DSTs

I-8-2-1) Exposure to Dinophysis

Most bivalve exposure experiments have been performed with the other genus responsible for DSP - Prorocentrum - and especially P. lima. Briefly, important findings are: genotoxic effects (DNA damage) towards mussel (P. perna and M. galloprovincialis) and clam (R. decussatus) hemocytes (Carvalho Pinto-Silva et al., 2005; Florez-Barros et al., 2011; Prego-Faraldo et al., 2016), modification of the expression of genes involved in immunological response in oyster (C. gigas) ( Romero-Geraldo et al., 2014) and physiological effects on mussel (P. perna) (e.g. shell-valve closure) (Neves et al. 2019).

Two studies examined the effect of natural blooms of D. acuminata on marine bivalves (Table 11). Exposure of adult oysters, mussels and clams to a highly concentrated bloom (up to ca. 17,000 cells L-1) of D. acuminata for months led to cytotoxic and immunotoxic effect in hemocytes, such as modifications in number and type (i.e. hyaline hemocytes vs. granular hemocytes) and of phenoloxydase activity (involved in stress response) (Mello et al., 2010) (Table 11). Similar immunotoxicity was noticed by Simões et al. (2015) in oyster and mussel hemocytes, when adults were exposed to a D. acuminata bloom (< 5,000 cells L-1) for a week, as well as an increase in total protein and number of circulating hemocytes. These last authors also observed infiltration of hemocytes in intestines and stomach (Table 11).

While the negative effects of Dinophysis towards marine bivalves have been suggested, only three experiments assessing the effect of Dinophysis cultures on adult bivalves have been carried out. In two recent feeding experiments of mussels (M. edulis) with cultures of D. acuta (OA, DTX1 and PTX2 producer) Nielsen and al. (2016) and Nielsen et al. (2020) observed a decrease in clearance rate but no mortality was observed. Importantly, in 2015, Basti et al. observed for the first time a direct mortality of mollusks exposed to Dinophysis (Table 11). Japanese (Patinopecten yessoensis) and noble (Mimachlamys nobilis) scallops were fed every day for one week with 20,000 cells of

Chapter 1: Literature review

D. caudata (PTX2 producer). After 24 h, hypersecretion of mucus and pseudo feces, retraction of the mantles and tentacles and reduction or even suppression of escape response, compared to the control, were observed. After 3 days, all P. yessoensis were paralyzed. After 7 days, mortality of P. yessoensis and M. nobilis reached 73% and 40%, respectively while necrosis was shown in the hepatopancreas (Table 11). However, the authors mentioned the role of unknown toxins/bioactive compound(s) (Basti et al., 2015b).

Table 11: Effect of mollusks exposure to Dinophysis

Concentration Animal Cell type Species Time Effect Reference (cells L-1) Oyster  Cytotoxicity: (Crassostrea modification in number gigas), mussel D. and type of hemocytes (Mello et (Perna perna) and Hemocytes acuminata up to 17,600 Months  Immunotoxicity: al., 2010) clam bloom modification in (Anomalocardia phenoloxydase activity brasiliana)  Cytotoxicity:  total proteins and number of Oyster (C. gigas) D. hemocytes (Simões et and mussel (P. Hemocytes acuminata 2950-4145 6-7 d  Immunotoxicity: al., 2015) perna) bloom modification in phenoloxydase activity  Infiltration of Histology hemocytes in stomach and intestine  Hypersecretion of Scallops mucus, pseudo feces, (Patinopecten retraction of mantles D. (Basti et yessoensis and Animal 20,000 24 h and tentacles and caudata al., 2015b) Mimachlamys reduction and nobilis) suppression of escape response 7 d  Mortality  Necrosis in Histology 7 d hepatopancreas

Chapter 1: Literature review

I-8-2-2) Exposure to DSTs

As for the effect of Dinophysis toxins on mice, rats and cell lines, cytotoxic and genotoxic effects have been observed on cell-lines of marine filter-feeding bivalves as well as immunological effects. Indeed, almost every author focused the effect of OA on mollusks hemocytes which are circulating cells in the hemolymph involved in the response against environmental stresses (including diseases and parasites) (Hégaret et al., 2003a) (Table 12).

Cytotoxicity was noticed by Prado-Alvarez et al. (2013, 2012) and Svensson et al. (2003) in mussel and clam. Hemocytes of these bivalves exposed to nM from µM levels of OA from hours to days, showed a reduction of viability and apoptosis (Table 12). In heart cell lines of oyster, Talarmin et al. (2008) reported cytotoxic activity towards a specific kinase (i.e. MAP kinase) (Liu and Hofmann, 2004) (Table 12).

Genotoxicity in hemocytes was observed in a time and dose dependent manner (Table 12). Indeed, McCarthy et al. (2014) and Prego-Faraldo et al. (2015) showed DNA fragmentation, damage and necrosis in mussel and oyster hemocytes exposed to nM of OA for hours to days, while Carvalho Pinto-Silva et al. (2003) reported formation of micronuclei in mussel hemocytes exposed to µM of OA for 24 h (Table 12).

Immunological effects have been shown by increased phagocytic activity of mussels hemocytes when exposed to OA (Malagoli et al., 2008). Two recent studies (Chi et al., 2017, 2016) with scallops (A. irradians) exposed to OA highlighted the negative effect on hemocytes antioxidant response (e.g. effect on super oxide dismutase enzyme), increase in ROS production. In addition, a high effect on immune response led to: increased damage of hemocytes and decrease in total number of circulating hemocytes, and modification of gene expression related to immune response (Table 12).

To the authors’ knowledge, there is no study investigating the effect of PTXs on marine mollusks. This lower interest could be explained by the fact that PTXs toxins do not have diarrheic effects on humans and because they are rapidly transformed/metabolized by mollusks, into nontoxic (even for mice and rats) forms (i.e. PTX2 transformed to seco acid derivatives such as PTX2-sa or 7-epi- PTX2-sa; Blanco, 2018; Nielsen et al., 2016). Moreover, most studies focused on the effect on

Chapter 1: Literature review mollusk hemocytes, while experiments on other cell types, such as early life stages (i.e. gametes or larvae), juvenile or adults, remained unexplored.

Table 12: Effect of mollusks exposure to Dinophysis toxins. OA, okadaic acid

Concentration Time Animal Cell type Toxin Effect Reference (nM) (h) (Svensson Mussel (Mytilus  Cytotoxicity:  of 50 % of Hemocytes OA 1000 72 et al., edulis) viability 2003) Mussel (Mytilus (Prado- galloprovincialis)  Cytotoxicity: damage and Alvarez et and clam Hemocytes OA 100 4 apoptosis al., 2013, (Ruditapes 2012) decussatus) Oyster (Talarmin  Cytotoxicity: effects on (Crassostrea Hearth cells OA 1000 24 et al., protein kinase gigas) 2008) (Carvalho Mussel (Perna  Genotoxicity:  number Pinto-Silva Hemocytes OA 30,000 24 perna) micronucleus et al., 2003) Mussel (M. 24 to (Mccarthy  Genotoxicyty: DNA edulis) and oyster Hemocytes OA 2.5 and 1.2 7 et al., fragmentation (C. gigas) days 2014) (Prego- Mussel (M.  Genotoxicity/cytotoxicity: Hemocytes OA 10 and 500 2 Faraldo et galloprovincialis) DNA damages and necrosis al., 2015) (Malagoli Mussel (M.  Immunotoxicity:  of Hemocytes OA 100 0.25 et al., galloprovincialis) phagocytic activity 2008)  Immunotoxicity: Scallop modification antioxidant (Chi et al., (Argopecten Hemocytes OA 50 6 response and  ROS 2017) irradians) production

Chapter 1: Literature review

Immunotoxicity/genotoxicit Scallop (A. 3 to y: damage and  number (Chi et al., Hemocytes OA 50 and 100 irradians) 12 hemocytes; modification of 2016) gene expressions

I-8-3) Effects of fish exposure to Dinophysis or DSTs

I-8-3-1) Exposure to Dinophysis

In a 1967 publication, Okaichi reported fish mortalities in the Gulf of Thailand during a bloom of the PTX-producer D. caudata. In summer 1993 in India, Santhanam and Srinivasan (1996) observed a decline in the number of fish but also diatoms and the absence of zooplankton during a bloom (1.5 x 103 cells mL-1) of D. caudata. Since then, to the authors’ knowledge, no mortality of fish due to Dinophysis has been reported in the literature whereas several laboratory experiments highlighted detrimental effects of DSTs and/or Dinophysis on fish.

To date, only one study focused on the in vitro effect of Dinophysis cultures on fish larvae (Rountos et al., 2019) (Table 13). These authors exposed larvae of two estuarine fish, Mediname beryllina and Cyprinodon variegatus to toxic strains of D. acuminata and A. catenella, and a mixture of the two species, at a concentration of 1000 cells mL-1 (Rountos et al., 2019). During a 96-h incubation, no effects on time to death were observed in fish larvae exposed to cultures of D. acuminata. However, D. acuminata had a sub-lethal effect, as it caused a growth rate decline of C. variegatus (Rountos et al., 2019) (Table 13).

Table 13: Effect of fish exposure to Dinophysis

Cell Concentration Animal Species Time Effect Reference type (cells mL-1) Inland silverside (Mediname (Rountos beryllina) and sheepshead Dinophysis   growth Animal 1000 4 days et al., minnow (Cyprinodon variegatus) acuminata rate 2019) larvae

Chapter 1: Literature review

Unlike with mollusks, experiments on fish exposure to Dinophysis, focused on early life stages, larvae and to a much lesser extent on adults. However, there are no previous studies exploring the effect of the non-diarrheic PTXs on mollusks and fish. Most authors observed histopathological features on exposed fish (e.g. Souid et al., 2018). The only reported experiment with Dinophysis (Rountos et al., 2019) observed harmful effect (i.e. growth rate decline), but did not perform any histological observations.

I-8-3-2) Exposure to DSTs

Some studies focused on the effects of DSTs on several fish life stages, such as embryos, larvae and adults (Table 14). Escoffier et al. (2007) exposed embryos of Medaka fish (Oryzias latipes) to nM of OA or to crude extracts of the OA producer Prorocentrum arenarium for 1 day and observed increased liver and digestive tract, as well as global body and vitellus areas when exposed to crude extract. The authors suggested the presence of additional substances in P. arenarium crude extract due to its more toxic effect and lower EC50 (Escoffier et al., 2007) (Table 14). Exposition of Longfin yellowtail (Seriola rivoliana) embryos to nM of OA and DTX1 for 28 h led to mortality of 60 and 100 % of the eggs, respectively (Le Du et al., 2017). Moreover, the authors reported a decrease in egg hatching, protein phosphatase activities and a significant modification of the expression of genes involved in bone development and in DNA replication (Le Du et al., 2017) (Table 14).

Exposure to ca. µM of OA for hours on larvae of gilt-head bream (Sparus aurata) showed an increase of peroxidation products and antioxidant enzymes (Souid et al., 2018). In the same study, histopathological observations revealed damages in hepatocytes, such as vacuolization, proliferation of fibroblasts, change of size and membrane degradations, and in gills with hypertrophy and fusion in the secondary lamellae (enlargement of tissues), dealing with necrosis (Souid et al., 2018) (Table 14).

Figueroa et al. (2020) exposed Zebrafish (Dania rerio) larvae to µM of OA and DTX1 for hours and reported a decrease in the fish growth rate of after 6 to 24 h of exposure, an inhibition of the activity of antioxidant enzymes (e.g. super oxide dismutase) and oxidative damages to proteins. Moreover, in the same study, OA and especially DTX1 led to pericardial edema and development

Chapter 1: Literature review disorders, as cyclopia, shortening of the anterior-posterior axis and delay (Figueroa et al., 2020) (Table 14).

Zhang et al., (2014) exposed Zebrafish (Dania rerio) adults µM of OA for 6 h by intraperitoneal injections and noticed modification of the expression of genes involved in the cell defense system (Table 14).

Table 14: Effect of fish exposure to Dinophysis toxins. OA, okadaic acid; BW, body weight

Concentration Time Animal Cell type Toxin Effect Reference (nM) (h) Medaka fish (Escoffier (Oryzias   liver and digestive tract Histology OA 520 (EC50) 24 et al., latipes) areas 2007) embryos OA from   liver and digestive tract Histology or P. 170 (EC50) areas,  body size and vitellus animal arenarium area  Mortality   egg hatching Longfin   protein phosphatase yellowtail activities (Le Du et (Seriola Animal OA 149 28  Modification of expression al., 2017) rivoliana) of genes involved in bone embryos development and DNA replication DTX1 217 Gilt-head  Modification in antioxidant bream responses:  of peroxidation (Souid et (Sparus Animal OA 9316 hours products and antioxidant al., 2018) aurata) enzymes larvae

Chapter 1: Literature review

 Hepatocytes: vacuolization, proliferation of fibroblasts, change of size and membrane Histology degradations  Gills: hypertrophy, fusion in secondary lamellae and necrosis   growth rate  Modification in antioxidant responses: inhibition of super oxide dismutase and damage Zebrafish (Figueroa Animal or to proteins (Dania OA 12,422 (LD50) hours et al., morphology  Pericardial edema rerio) larvae 2020)  Development disorders: cyclopia, shortening of the anterior-posterior axis and delay of developments

DTX1 8,588 (LD50) Zebrafish Animal  Modification of expression 176 to 1760 (Zhang et (D. rerio) (intraperitoneal OA 6 h of genes involved in cell g-1 BW al., 2014) adults injections) defense system

I-9) Biology of the Pacific oyster C. gigas and fish sheepshead minnow C. variegatus

The thesis focused on two marine animals, the Pacific oyster, Crassostrea gigas (Thunberg, 1793) and the sheepshead minnow (Cyprinodon variegatus) (Lacépède, 1803). The next two sections describe the biology of these two species with a focus on reproduction.

I-9-1) C. gigas

The Pacific oyster (Mollusca, Bivalvia, Ostreidae) is a sessile organism that feeds on suspended particles, mainly phytoplankton (Bayne, 2017). This oyster is a cosmopolitan species found in

Chapter 1: Literature review coastal and estuarine areas, which was imported from Japan in more than 70 countries by the second half of the 19th century in order to establish new oyster fisheries (Bayne, 2017). The Pacific oyster has a high commercial value and is used as a model organism for scientific purposes (Bayne, 2017). Shellfish farming activities are an important economic sector in France. According to FranceAgriMer (2020) (an institution affiliated to the French ministry of agriculture), in 2017 shellfish sales in France represented a total of 551 millions of euros, 402 of which corresponded to oysters, and the farming sector, in 2017, provided 14900 jobs in 2645 farms.

The Pacific oyster has an annual life cycle composed of two stages: (i) a planktonic stage of embryos and larvae and (ii) a sessile stage, after metamorphosis and fixation of larvae, of juveniles (i.e. spat) and adults (Gosling, 2015) (see example of the life cycle of the Pacific oyster Figure 5) (Vogeler et al., 2016).

Chapter 1: Literature review

Figure 5: Life cycle of the Pacific oyster, Crassostrea gigas, modified from (Vogeler et al., 2016). hpf, hours post fertilization; dpf, days post fertilization; mpf, months post fertilization

The fertilization of the C. gigas is external, with a release of female (oocytes) and male (spermatozoa) gametes in the surrounding environment (i.e. seawater) (Figure 5). Importantly, in the first hours of bivalve life (6-12 h for the Pacific oyster; Figure 5), their protection is limited due to various factor, such as the absence of shell (Hégaret et al., 2007).

I-9-1) C. variegatus

The sheepshead minnow (Actinopterygii, Teleostei, Cyprinodontidae) is a teleost- ray-finned fish, a group composed of > 23,500 species (Burton and Burton, 2017; Volff, 2005). This omnivorous and benthic fish is found from fresh to salt waters at a wide range of temperature, from the East coast of North America to the East coast of Brazil (Bennett and Beitinger, 1997; Froese and Pauly,

Chapter 1: Literature review

2019; Zappler, 2000). C. variegatus is ecologically and environmentally important as prey of fish species of commercial value (Gervasi et al., 2019; Hall et al., 1994; Rosenfield et al., 2004). In addition, it is a valuable resource for its use as forage fish in aquaculture and is also as a experimental model fish orgamism because of its rapid development in aquarium and its tolerance to laboratory conditions (Schnitzler et al., 2017).

Reproduction of the teleost fish C. variegatus occurs in spring-summer. Females spawning in a nest is followed by males fertilization (Wootton and Smith, 2014). Eggs hatchafter ca. 1-2 weeks and larvae development lasts for ca. 28 days (Schnitzler et al., 2017). These phases are sensitive and critical, because, for instance due to their low, or even null mobility, they cannot escape from waters contaminated by toxic phytoplankton and phycotoxins (Wiegand et al., 1999).

I-10) DSTs distribution

The following maps represent the global and regional distribution of DST measurements in marine animals, in Dinophysis cells from field samples and cultures, as well as extracellular DST in seawater (collected with solid phase adsorption toxin tracking, SPATT). The data encompass the locations, species and toxin profiles of Dinophysis species or taxa when marine animals were analyzed. Data were obtained and updated from Lee et al. (Lee et al., 1989), Blanco et al. (2005), Reguera and Pizarro (2008), Reguera et al. (2012) and Reguera et al. (2014). Scientific publications on Dinophysis occurrences but without associated toxin profiles or quantifications were not included. Toxin contents of Dinophysis species were reported on a per cell basis (pg cell-1) or in a few cases per volume (ng mL-1), when the concentration in a per cell basis was not available in the publications, while presence or absence of the toxins in marine animals were shown. Marine animals are grouped by taxa (e.g. C. gigas and C. virginica = oysters). The global (worldwide) distributions were represented in WSG84 EPSG 3857 system and regional maps in WSG84 EPSG4326 system. A complete description of DST measurements in publications is represented for France whereas a brief overview was made for the other regions. Similarly, monitoring programs of phytoplankton and toxins are established in several countries (e.g. North America (NSSP 2017) or Spain, see references in Díaz et al. (2016)), but only a description of the French monitoring network (i.e. REPHY) is presented.

Chapter 1: Literature review

Data and additional information (e.g. toxin quantities, coordinates, method of analysis) were compiled in a file available online:

I-10-1) Global distribution

This work is based on an important database of 461 records (from 1978 to July 2020) from 163 scientific publications, conference papers, reports and conference abstracts (Table 15).

The proportion of DST measurements in Dinophysis species and in animals is similar. Also, the proportion of measurements in Dinophysis species did not increase since 2006 compared to the period pre-2006, although laboratory cultures were established. This can be explained by (i) the lack of laboratory studies due to the difficulties in cultivating Dinophysis (see section “I-4) Culture of the three-tier food chain of Dinophysis”), (ii) measurements in marine animals are an efficient way to prevent/study DSTs and DSP and (iii) an important part of DSTs is found in marine animals i.e. after being exposed to Dinophysis blooms.

Thereby, this section adds supplementary information to the work of Reguera et al. (2014) (i.e. addition of 187 toxin measurements from 44 new scientific publications) and corroborates their interpretation of the species/toxin distributions. However, some data reported in Reguera et al. (2014) are missing in this work because of the lack of toxicity information or toxin measurements, or because data were not accessible (not found or not in English).

The genus Dinophysis has a worldwide distribution from cold, temperate, tropical to warm waters (Nagai et al., 2020; Reguera et al., 2012) (Figure 6). Importantly, new reports of DST measurements (mainly SPATT measurements) (Figure 7) were found in Africa, Antarctica, Gulf of Oman and Argentina which strengthen the worldwide and cosmopolitan distribution of several species of the genus Dinophysis. Records exist mainly in areas where shellfish farming is an important economic sector and where monitoring programs are implemented i.e. Western Europe, Japan and South America (especially Chile).

Chapter 1: Literature review

Figure 6: Global distribution of DSTs (black dots: n = 461) combining reports of toxins in Dinophysis species (in vivo and in vitro) (n = 233), marine animals (n = 209) and SPATT (n = 19). Dots can be superposed

Chapter 1: Literature review

Figure 7: Global distribution of DSTs measurements (white dots) in SPATT (n = 19). Dots can be superposed

Chapter 1: Literature review

I-10-2) Regional distribution

I-10-2-1) French case study

In summer 1983, more than 3000 people in southern Brittany (France) were intoxicated after the consumption of blue mussels (Mytilus edulis) contaminated with DSTs. The causative species was identified as D. acuminata, and synthesized OA (Alzieu et al., 1983; Kumagai et al., 1986; Lassus et al., 1985; Lee et al., 1987) (Figure 8 A-B; Table 16).

Masselin et al. (1992) and Marcaillou et al. (2005) determined OA concentrations of 7-14 pg cell- 1 and DTX1 of 1 pg cell-1 in isolated (or picked) cells of D. sacculus and a mean concentration of 79 pg cell-1 of OA in D. acuminata, in samples from southern Brittany (Figure 8 B; Table 16). Differences in toxin profile of the D. sacculus between Masselin et al. (1992) and Séchet et al. (2020) (Tables 16 and 17) could be explained by differences in the analytical systems and by the fact that PTX2 was not monitored in France in the 1990’s..

Amzil et al. (2007) found PTX2, associated with PTX2sa and 7-epi-PTX2sa, and OA group toxins in mussels, clams and oysters from the French Mediterranean, southern Bay of Biscay and southern Brittany coasts (Figure 8 F) (Table 16).

Since the establishment of Dinophysis cultures in the Phycotoxins laboratory in France, the toxin profiles of French strains is now possible (Table 17). D. acuminata was found along the Atlantic and English Channel coasts (Figure 8 A-E) and produced only OA, from 11 to 96 pg cell-1 (Table 17). D. sacculus was isolated from the French Mediterranean (Figure 8 E) and southern Bay of Biscay to Brittany coasts (Figure 8 A-C). This species synthesized 7 times more PTX2 than OA (Table 17). D. acuta was found in the same places as D. sacculus (Figure 8 A-C, E), but it produced mainly diarrheic toxins, OA (10 to 98 pg cell-1) and DTX2 (7 to 30 pg cell-1), and PTX2 (6 to 82 pg cell-1) (Table 17). D. fortii and D. caudata were sometimes found all along the Bay of Biscay (Figure 8 A-C) and were high producers of PTX2 (up to 200 pg cell-1 for D. caudata) (Table 17).

In France, D. sacculus and D. acuminata exhibited two different toxin profiles, constituted by OA and PTX2, and only OA, respectively (Sechet et al. 2020) (Table 17), while such differentiation by the chemotype is not possible in other regions of the world (Table 15 available online at: https://doi.org/10.17882/76433). This characteristic is interesting and could be a criteria of

Chapter 1: Literature review differentiation between these species (or ecotypes) (Séchet et al., 2020) which exhibit small morphological and genetic differences (Lassus and Bardouil 1991, Zingone et al. 1998, Séchet et al., 2020) (see section “I-2) Morphology and taxonomy of Dinophysis”).

Chapter 1: Literature review

Table 16: Literature review of French DST measurements in marine animals and in field populations of Dinophysis species. Measurement from the French database (i.e. REPHY) are not presented in this table. Toxin quantities in Dinophysis species are mean values in pg cell-1, while the toxin in marine animals are qualitative profiles. Marine animals are regrouped by taxa. French coastlines of the Mediterranean Sea are in red, Atlantic Ocean in green and English Channel in red. Additional information (e.g. coordinated, method of analysis) are compiled in a file, Table 15 available online at https://doi.org/10.17882/76433. OA, okadaic acid; DTX1, dinophysistoxin 1, DTX2, dinophysistoxins 2; DTX3, dinophysistoxin 3; PTX2, pectenotoxin 2; PTX2sa, pectenotoxin 2-seco acid; 7-epi-PTX2sa, 7-epi-pectenotoxin 2-seco acid

7-epi- Species Place OA DTX1 DTX2 DTX3 PTX2 PTX2sa Reference PTX2sa

(Kumagai et In mussels Le Havre 1 al., 1986)

(Lee et al., In mussels Le Havre 1 1987)

(Masselin et D. sacculus Le Croisic 11 al., 1992) D. sacculus Morgat 8 D. sacculus Kervel 18 1 D. sacculus Pont-Aven 7 D. sacculus La Gachère 14

D. (Marcaillou Kervoyal 79 acuminata et al., 2005) (Amzil et al., In mussels Loscolo 1 1 2007) In clams Loscolo 1 1 1 1 1 1 In mussels Arcachon Bay 1 1 1 1 1 1 In oysters Arcachon Bay 1 1 1 In mussels Thau Lagoon 1 1 1 1 In oysters Thau Lagoon 1 1 1 1

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Table 17: Literature review of DST measurements in Dinophysis cultures from French coastal waters. Toxin quantities in Dinophysis are mean values in pg cell-1. French coastlines of the Mediterranean are in red, Atlantic in green and English Channel in red. Additional information (e.g. coordinated, method of analysis) is compiled in a file, Table 15 available online at: https://doi.org/10.17882/76433. OA, okadaic acid; DTX2, dinophysistoxin 2; PTX2, pectenotoxin 2

Species Strain Place OA DTX2 PTX2 Reference

D. sacculus IFR-DSA-01Lt La Teste, Arcachon 5 86 (Gaillard et al., 2020b) D. acuminata IFR-DAU-02Ar Bouée 7, Arcachon 87

D. sacculus IFR-DSA-01Lo Loscolo 2 18 (Gaillard et al., 2020c) D. sacculus IFR-DSA-01Me Arcachon 9 84 D. sacculus IFR-DSA-02Th Thau Lagoon 11 44 D. acuminata IFR-DAU-03An Antifer 39 (Séchet et al., 2020) D. acuminata IFR-DAU-01Es Saint Aubin - les Essarts 33 D. acuminata IFR-DAU-01Ca Cabourg 32 D. acuminata IFR-DAU-01Ke Kervel 81 D. acuminata IFR-DAU-01Du Dumet 11 D. acuminata IFR-DAU-02Bm Le Croisic 96 D. acuminata IFR-DAU-01Bo Noirmoutier 92 D. acuminata IFR-DAU-02Ar Arcachon 45 D. sacculus IFR-DSA-01Co Concarneau 7 21 D. sacculus IFR-DSA-01Lo Loscolo 3 22 D. sacculus IFR-DSA-01Me Arcachon 3 29 D. sacculus IFR-DSA-01Lt La Teste 3 75 D. sacculus IFR-DSA-01Th Thau Lagoon 5 33 D. sacculus IFR-DSA-02Th Thau Lagoon 7 36 D. sacculus IFR-DSA-03Th Thau Lagoon 9 34 D. sacculus IFR-DSA-01Vp Vic Lagoon 2 4 D. sacculus IFR-DSA-01Po Ponant Lagoon 25 89 D. sacculus IFR-DSA-01Ma Berre Lagoon 4 16 D. sacculus IFR-DSA-01Lj Berre Lagoon 9 69 D. tripos IFR-DTR-01Ar Arcachon 176

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D. caudata IFR-DCA-01Ke Kervel 168 D. caudata IFR-DCA-03Ke Kervel 200 D. caudata IFR-DCA-02Lo Loscolo 39 D. caudata IFR-DCA-04Tr Le Grand Travers 140 D. acuta IFR-DAC-03Ke Kervel 19 7 54 D. acuta IFR-DAC-02Lc Le Croisic 23 14 69 D. acuta IFR-DAC-03Lc Le Croisic 37 30 82 D. acuta IFR-DAC-04Lc Le Croisic 16 13 30 D. acuta IFR-DAC-01Ar Arcachon 38 21 64 D. acuta IFR-DAC-01Po Ponant Lagoon 10 8 6

The French monitoring network REPHY (Observation and Surveillance Network for Phytoplankton and Hydrology in coastal waters) was created in 1984 and the first available data in open access date from 1987 (Belin et al., 2020; REPHY, 2019). This long-term (33 years in 2020) time series of data display spatio-temporal information of phytoplankton concentrations (including toxic and non-toxic taxa), measurements of abiotic factors in the water (e.g. salinity, temperature) and phycotoxin concentration in bivalves. Dinophysis species is found along each French coastline (Mediterranean including Corsica, Bay of Biscay, Brittany and English Channel). Generally, when Dinophysis species are present, the concentration is around 100 cell L-1 (which is also the detection limit in the French monitooring system) and the annual maxima rarely exceed 10,000 cells L-1 (Belin et al., 2020) with a maximum of 803,000 cells L-1 in Calvados in 2006 (Belin et al., 2020). Dinophysis species are low biomass HABs, cause shellfish harvesting bans even at low concentrations. When compared to ASP and PSP, DSP affected most areas since 1987 (around 30 % of the monitored Atlantic areas were contaminated over the last 33 years). Since the beginning of DST measurements in filter-feeding bivalves, mussels (since 1987) and oysters (since 2001) were contaminated at least once a year and this contamination was mainly due to D. acuminata and D. sacculus (Belin et al., 2020).

Chapter 1: Literature review

I-10-2-2) Northern Europe

The most common species in Northern Europe are the cosmopolitan D. acuminata and the cold- temperate species D. acuta. Other rare species such as D. norvegica and D. rotundata (Phalacroma rotundatum), were also reported in these temperate and cold-temperate waters (Miles et al., 2004b; Reguera et al., 2014). These D. acuminata and D. acuta mainly synthesized OA and PTX2, up to 130 pg cell-1 of OA (from net hauls; Haamer et al., 1990) and 134 pg cell-1 of PTX2 (from culture; Nielsen et al., 2013) for D. acuta. Additionally, DTX1 (Godhe et al., 2002; Kuuppo et al., 2006; Lee et al., 1989; Nielsen et al., 2016, 2013) and PTX 12 (Miles et al., 2004b) were also recorded in these species (Figure 9 A) while low amounts of PTX2 and PTX12 were found in D. rotundata (P. rotundatum) (Miles et al., 2004b) (Figure 9 A).

Toxins were mainly found in mussels, but exceptionally in crabs, due to an earlier bloom period of D. acuta in 2002 (Torgersen et al., 2005), and consisted mainlyby diarrheic toxins and traces of pectenotoxins (Figure 9 B-C).

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Figure 9: Northern Europe. Distribution of DST measurements A. in Dinophysis species and B. marine animals, and C. in marine animals in Western Russia

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I-10-2-3) Great Britain and Republic of Ireland

Scientific reports of DST measurements in Dinophysis species in UK are scarce (Figure 10 A). However, more data are available from field monitoring databases (Dhanji-Rapkova et al., 2018; Salas and Clarke, 2019).

In Ireland, D. acuta was reported to synthesize high quantity of diarrheic toxins: OA (from 24 to 85 pg cell-1) and DTX2 (20 to 78 pg cell-1), and to a lesser extent PTX2 (14 to 27 pg cell-1) (Fernández Puente et al., 2004; Fux et al., 2010; James et al., 1997).

Diarrheic OA, DTX1 and DTX2 were found mainly in mussels and oysters, co-occurring with PTX2 in a few samples, but also in clams, razor clams and scallops (Figure 10 B-C).

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Figure 10: Great Britain and Republic of Ireland. Distribution of DST measurements A. in Dinophysis species and B. in marine animals, and C. in marine animals in Southern Scotland

I-10-2-4) Spain and Portugal

Several records of Dinophysis species are available for the Galician Rias (Northwestern Spain), site of an intensive (2-3 x 105 t per year) mussels cultivation (Reguera et al., 2014). The main species found in these areas were D. acuta, D. acuminata, the tropical to warm water species D. caudata, and to a lesser extent D. sacculus, D. fortii, D. tripos and D. rotundata (P. rotundatum) (Figure 11 A). The main toxins found were OA (up to 45 pg cell-1 in D. acuta (García-Portela et al., 2018), DTX2 and PTX2 (up to 205 pg cell-1 in D. fortii (Rodríguez et al., 2012), and PTX11 and OA D8 was recorded once in D. acuta (Pizarro et al., 2009). Only traces of OA and DTX2 (< 1 pg cell-1) were found in D. rotundata (P. rotundatum) (Fernández et al., 2001). (Figure 11 A). The temperate species D. sacculus was reported on both the Mediterranean and Atlantic coasts of Spain and synthesized only OA (from 3 to 17 pg cell-1; Delgado 1996, Canete et al. 2008) or OA and PTX2 in laboratory cultures (8 and 13 pg cell-1, respectively; Riobó et al. 2013) (Figure 11 B). According to García-Altares et al. (2016) study, intracellular DSP toxin concentrations in D. sacculus plankton concentrates were overestimated (i.e. maximum of 668 pg PTX2 cell-1).

In the same area, the main contaminated shellfish were mussels followed by clams, oysters, razor clams, cockles and limpets in which OA, DTX2, PTX2sa and DTX3 were reported (Figure 11 C).

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Figure 11: Spain and Portugal. Distribution of DST measurements A. and B. in Dinophysis species in Western and Southern Iberia and Western Spain, respectively and C. in marine animals

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I-10-2-5) Morocco, Tunisia, Italia, Croatia, Greece and Russia (Black Sea)

Toxic species of Dinophysis were reported in Italy (Figure 12 A). Indeed, D. caudata and D. fortii were found in the Northern Adriatic Sea (Italy, Croatia) and D. sacculus was found in Sicilia (Figure 12 A). The main toxin found was PTX2 (from 90 to 253 pg cell-1 in D. fortii Draisci et al., 1996, Sasaki et al., 1999) and to a lesser extent OA. D. sacculus synthesized low amounts (< 1 pg cell-1) of OA and DTX1, which is surprising for this species (Table 15 available online at: https://doi.org/10.17882/76433), and could be due to the analyses.

Mussels from the Atlantic coasts of Morocco to the Mediterranean Sea Tunisia, Sardinia, Greece (in a major shellfish production area; Reizopoulou et al. (2008)) and the Black Sea were mainly contaminated by OA (Figure 12 B).

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I-10-2-6) Nigeria, Angole and South Africa

Only few reports of DSTs are available for Africa and they are mostly from South Africa (Figure 13 B-C). The main organism reported to synthesize OA in this country is D. acuminata (from net haul samples rich in D. acuminata) and it contained DTX1, PTX2, PTX2sa, and to a lesser extent PTX11 (< 1 pg cell-1) (Pitcher and Probyn, 2011) (Figure 13 B).

However, several records in mussels from South Africa showed the accumulation of OA (Figure 13 C). This toxin was also found in clams in Angola (Rangel et al., 2008) and in SPATT associated with PTX2 in Nigeria (Zendong et al., 2016b). These records suggest that DSTs are present along the West African coasts and could be synthesized by species belonging to the D. acuminata- complex.

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I-10-2-7) North America

In USA, DST measurements and DSP events are localized in the Gulf of Mexico (Campbell et al., 2010) (Figure 14 D), and in North West (Taylor et al., 2013) (Figure 14 B) and North East USA (Wolny et al., 2020) (Figure 14 C). However, additional measurements are available in databases (e.g. CBP 2020, McKenzie et al., 2020).

D. acuminata, mainly found in the North East part of the USA and South East of Canada, was the dominant species while D. ovum was mainly present in the Gulf of Mexico (Swanson et al., 2010) (Figure 14 D). In South East Canada, the presence of D. norvegica was noticed (Cembella, 1989; Subba-Rao et al., 1993) (Figure 14 C ) and recently, D. norvegica from the Gulf of Maine has been successfully cultivated (Deeds et al., 2020). Okadaic acid was mainly found, especially in D. ovum (from 11 to 50 pg cell-1), and PTX2 was found to be produced mainly by D. acuminata, in association with DTX1 and OA. The presence of DTX3 was also reported in Dinophysis from Washington state (Trainer et al., 2013) (Figure 14 B).

So far, only few published records of mollusk contaminations are available in these areas, but mostly mussels and oysters were contaminated by OA group toxins (Trainer et al., 2013) (Figure 14 A), may be due to the fact that PTXs are not regulated in the US (Wolny et al., 2020) .

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I-10-2-8) Central and South America

DST measurements are not available for Central American countries and for Colombia, Venezuela and Northern Brazil (Reguera et al., 2014) (Figure 15 A-H).

On the Pacific coasts (Peru and Chile) D. acuminata (and D. acuminata-complex) and to a lesser extent D. caudata were reported (Figure 15 E, G). They mainly produced PTX2 (from 1 to 90 pg cell-1 in D. acuminata (Blanco et al., 2007, Alcántara-Rubira et al., 2018) and PTX11 (Fux et al., 2011; Krock et al., 2009), and a low amount of OA (< 1 pg cell-1) (Alcántara-Rubira et al., 2018).

In Peru, scallops were contaminated by PTX2 and OA (Figure 15 A). In Northern Chile, shellfish (including clams, scallops and piures) were found to accumulate non-diarrheic toxins (PTX2 and PTX2sa) (Figure 15 C), while in the Southern part of the country, which is the main shellfish farming region (Alves-de-Souza et al., 2019), diarrheic toxins (i.e. OA, DTX1 and DTX4) and PTX2 were repeatedly found in mussels, clams and also sea snails (Figure 15 C).

The Eastern coasts, i.e. in Southern Brazil, Uruguay and Argentina were also exposed to D. acuminata-complex, D. acuminata, D. tripos and D. ovum. Okadaic acid was the predominant toxin found, followed by PTX2 and PTX11 (Figure 15 F, H).

Marine animals contaminated by OA and DTX1 were found in Brazil, including mussels, clams and oysters but also octopuses and fish (Figure 15 B, D) (Mafra et al., 2015, 2014).

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I-10-2-9) Japan and Republic of Korea

D. acuminata and D. fortii occur along the Japanese Sea of Japan and Pacific Ocean coasts. In Southern Japan (Figure 16 E) the above-mentioned species co-occur with D. caudata whereas additional toxic species, e.g. D. tripos, D. rotundata (P. rotundatum) and D. norvegica (Figure 16 A-D) are found in the north. Suzuki et al., (2009) reported the presence of a toxic D. infundibulus (Figure 16 A) and Uchida et al., (2018) of D. mitra (P. mitra) (Figure 16 A). In Korea, D. acuminata was reported once in 2010 (Kim et al., 2010) (Figure 16 E). The main toxins found were PTX2, DTX1 and OA (Figure 16 A-E).

Although these toxic species have been found repeatedly in the Japanese and South Korean waters, only a few records of contaminated mussels, oysters and scallops by DSTs (OA, DTX1, DTX2, DTX3 and PTX2 and PTX6) have been reported (Figure 17 A-B).

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Figure 17: Japan and Republic of Korea. Distribution of DST measurements in marine animals in A. Southern Republic of Korea and B. North East of Japan

I-10-2-10) East Asia

Records of Dinophysis in Chinese waters are very rare. Few studies have reported the occurrences of D. acuminata (from cultures; (Gao et al., 2017; Jia et al., 2019) and D. caudata and their associated toxins (mainly PTX2) in these waters (Figure 18 A).

However, accumulation of DSTs (OA and DTX1) in mussels, oysters and scallops from Hong- Kong to the Northern China have been reported, suggesting the presence of more OA-producers (Figure 18 B).

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Figure 18: East Asia. Distribution of DST measurements A. in Dinophysis species and B. in marine animals in China and Hong Kong

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I-10-2-11) India and the Arabian Sea

In India, a DSP intoxication was listed due to contaminated oysters, but no toxin analysis was performed (Karunasagar et al., 1989) (Figure 19). In the Arabian sea, OA has been detected in mussels and associated with the presence of several Dinophysis species (Al Muftah et al., 2016) (Figure 19).

Figure 19: West Asia. Distribution of DST measurements in Dinophysis species and marine animals in India and Qatar

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I-10-2-12) South Asia and North Oceania

D. caudata and D. miles, from Republic of Singapore and Indonesia coastal waters were found to contain OA and DTX1 (Holmes et al., 1999; Holmes and Lewis, 2002; Marasigan et al., 2001) (Figure 20).

Figure 20: South Asia and North Oceania. Distribution of DST measurements in Dinophysis species in Republic of Singapore and Indonesia

I-10-2-13) Oceania

In Australia, only one published report showed that D. caudata synthesized PTX2 (and PTXsa) (Fernandez et al., 2013) (Figure 21 A). However, more data exist in databases for Australia (Farrell et al., 2018; Brett et al., 2020).

Contamination of oysters, mussels, clams, scallops and pipis by PTX2 (and PTX2-sa and 7-epi- PTX2-sa), OA and DTX1 (also in Tasmania) have been reported (Burgess et al., 2003; Madigan et al., 2006) (Figure 21 C).

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In New Zealand, D. acuta and to a lesser extent D. acuminata have been found to synthesize high amounts of PTX2 (up to 108 pg cell-1) (MacKenzie et al., 2005) and PTX11 (up to 65 pg cell-1) (MacKenzie et al., 2005) and traces of PTX12sa and PTX13 were found in D. acuta, co-occurring with traces of OA and DTX1 (MacKenzie et al., 2005) (Figure 21 B).

One report showed the contamination of mussels in New Zealand by OA, DTX1 and PTX2 (Figure 21 D) (MacKenzie et al., 2019).

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Figure 21: Oceania. Distribution of DST measurements in Dinophysis species in A. Australia and B. New Zealand, and in marine animals respectively for the same locations C. and D

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I-10-3) Conclusion of the DSTs distribution

Globally, since the review of Reguera et al. (2014) no trend of increasing frequency of DSP in monitored areas has been observed. However this representation underestimates the number of DSTs reports due to the fact that (i) only report of DST measurements in scientific publications are shown and (ii) reports mainly concern areas where aquaculture is developed and is an important economic sector (e.g. Western Europe and Chile) (Reguera et al., 2012), which suggests that DSTs may be present in areas where no monitoring programs exist (e.g. Africa, India) and (iii) misdiagnosis of DSP syndrome is made, due to the fact that the symptoms (especially diarrhea) are similar to bacterial contamination (Reguera et al., 2014).

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Chapter 2: Ecophysiological experiments Part 1: Factorial design on Teleaulax and predator feeding

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II-1) Combined effects of temperature, irradiance and pH on Teleaulax amphioxeia (Cryptophyceae) physiology and feeding ratio for its predator Mesodinium rubrum (Ciliophoran)

This Chapter 2-Part 1, has been published in Journal of Phycology in January 2020:

Gaillard, S., Charrier, A., Malo, F., Carpentier, L., Bougaran, G., Hégaret, H., Réveillon, D., Hess, P., Séchet, V., 2020. Combined effects of temperature, irradiance, and pH on Teleaulax amphioxeia (Cryptophyceae) physiology and feeding ratio for its predator Mesodinium rubrum (Ciliophora). Journal of Phycology. 56, 775-783. https://doi.org/10.1111/jpy.12977. Open access: https://hal.archives-ouvertes.fr/hal-02880016

II-1-1) Graphical abstract

Importance prey concentration

Factorial design 23* – T. amphioxeia Nutrition – . u um

M 400 17.6 Acclimation O  HL condition D pH 7.6 E Pigments LL 100  L Fv/Fm condition 17.6 pH Semi- S 7.6 continuous

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II-1-2) Résumé

Le cryptophyte Teleaulax amphioxeia est une source de plastes pour le cilié Mesodinium rubrum. Ces deux organismes sont membres de la chaîne trophique de plusieurs espèces de Dinophysis. Il est important de mieux comprendre l'écologie des organismes aux premiers niveaux trophiques avant d'évaluer l'impact des principaux facteurs du changement global sur Dinophysis spp. Par conséquent, les effets combinés de la température, de l'irradiance et du pH sur le taux de croissance, l'activité photosynthétique et la teneur en pigments d’une souche tempérée de T. amphioxeia ont été étudiés en utilisant un plan factoriel complet (plan composite centré 23*) dans 17 bioréacteurs contrôlés individuellement. Le modèle dérivé prédit un taux de croissance optimal pour T. amphioxeia à une intensité lumineuse de 400 µmol photons · m-2 · s-1, un pH plus acide (7,6) que la moyenne actuelle et une température de 17,6 ° C. Une interaction entre la température et l'irradiance sur la croissance a également été trouvée, tandis que le pH n'a pas eu d'effet significatif. Par la suite, pour étudier les impacts potentiels de la qualité et de la quantité des proies sur la physiologie du prédateur, M. rubrum a été nourri avec deux rapports proies : prédateurs différents et avec des cultures de T. amphioxeia préalablement acclimatées à deux intensités lumineuses différentes (100 et 400 µmol photons · m-2 · s-1). La croissance de M. rubrum semble dépendre de manière significative de la quantité de proies, tandis qu’aucun effet de la qualité des proies n'a pu être observé. Cette étude multiparamétrique suggère une probable augmentation de la présence de T. amphioxeia dans les conditions climatiques futures et interroge sur un potentiel effet sur l’augmentation des occurrences de Mesodinium spp. et Dinophysis spp., qui devrait faire l’objet d’études plus approfondies.

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II-1-3) Abstract

The cryptophyte Teleaulax amphioxeia is a source of plastids for the ciliate Mesodinium rubrum and both organisms are members of the trophic chain of several species of Dinophysis. It is important to better understand the ecology of organisms at the first trophic levels before assessing the impact of principal factors of global change on Dinophysis spp. Therefore, combined effects of temperature, irradiance and pH on growth rate, photosynthetic activity and pigment content of a temperate strain of T. amphioxeia were studied using a full factorial design (central composite design 23*) in 17 individually controlled bioreactors. The derived model predicted an optimal growth rate of T. amphioxeia at a light intensity of 400 µmol photons · m-2 · s-1, more acidic pH (7.6) than the current average and a temperature of 17.6 °C. An interaction between temperature and irradiance on growth was also found, while pH did not have any significant effect. Subsequently, to investigate potential impacts of prey quality and quantity on the physiology of the predator, M. rubrum was fed two separate prey: predator ratios with cultures of T. amphioxeia previously acclimated at two different light intensities (100 and 400 µmol photons · m-2 · s-1). M. rubrum growth appeared to be significantly dependant on prey quantity while effect of prey quality was not observed. This multi-parametric study indicated a high potential for a significant increase of T. amphioxeia in future climate conditions but to what extent this would lead to increased occurrences of Mesodinium spp. and Dinophysis spp. should be further investigated.

II-1-4) Key words

Dinophysis; ecophysiology; full factorial design; global change; Mesodinium rubrum; Teleaulax amphioxeia

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II-1-5) Introduction

The cryptophyte Teleaulax amphioxeia (Conrad) Hill (Hill, 1992) is observed worldwide and has been reported to forms red tides in coastal waters (Yoo et al. 2017). This organism is a prey and a source of plastids for the mixotrophic ciliate Mesodinium rubrum (Lohmann 1908, = Myrionecta rubra Jankowski 1976), which is also known to form red-colored blooms in coastal ecosystems (Lindholm, 1985). The ingested plastids and nuclei of T. amphioxeia are incorporated in M. rubrum (Johnson et al., 2007; Johnson and Stoecker, 2005; Yih et al., 2004) and remain photosynthetically and transcriptionally active to sustain growth of the ciliate (Johnson et al., 2007; Kim et al., 2017). Cryptophytes, as T. amphioxeia, play an important role in ecosystem dynamics as they are a ‘common food organism’ (Yih et al., 2004) of several protists (Peterson et al., 2013; Smith and Hansen, 2007). Interestingly, the mixotrophic and harmful species of the dinoflagellate genus Dinophysis (Ehrenberg 1841) exhibit chloroplasts of cryptophyte origin, obtained by ingestion of M. rubrum (Park et al., 2006; Wisecaver and Hackett, 2010). A both relationship between T. amphioxeia and M. rubrum and between M. rubrum and occurrence of Dinophysis spp. has been suggested in natural environments (Hamilton et al., 2017; Herfort et al., 2011; Peterson et al., 2013). The influence of M. rubrum concentration on growth (Hattenrath-Lehmann and Gobler, 2015; Kim et al., 2008; Nagai et al., 2011; Park et al., 2006; Smith et al., 2018; Tong et al., 2011) and toxin production (Gao et al., 2017) of Dinophysis spp. was even observed in lab experiments. M. rubrum growth depends on cryptophytes including T. amphioxeia (Johnson, 2011; Yih et al., 2004) but also on abiotic factors, such as light (Moeller et al., 2011), pH (Smith and Hansen, 2007) or temperature (Basti et al., 2018). However few studies have focused on the physiology of T. amphioxeia and its effects on growth and pigment content of M. rubrum. Such studies are thus required to improve knowledge on the bottom of the food chain of Dinophysis spp., and consequently on the understanding of environmental dynamics of both M. rubrum and Dinophysis spp. growth and blooms. It is widely recognized that climate change modifies harmful algal bloom duration and frequency (Glibert et al., 2014; Gobler et al., 2017) and in this context, according to Wells et al. (2015), temperature, light and pH appear to be key variables.

Therefore, we investigated the effects of these three parameters on the ecophysiology of T. amphioxeia. First, a full factorial design (central composite design 23*) was applied to assess the direct combined effects of temperature, irradiance and pH as well as their interactions on the

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Chapter 2-Part 1: Factorial design on Teleaulax and predator feeding maximum growth rate, pigment content and maximum quantum yield of the photosystem II 3* (Fv/Fm). The central composite design 2 required 15 experimental conditions with triplicate cultures for the central condition (Lundstedt et al., 1998). A culture device composed of 17 photo- bioreactors, previously developed by Marchetti et al. (2012), was used to perform the factorial design. This approach minimizes the number of experiments that need to be carried out to assess the effects of parameters on a specific response. Also, this design easily allows for the development of statistical models of the maximum growth rate, pigment quantity and Fv/Fm. Finally, M. rubrum was fed two photoacclimated cultures (100 and 400 µmol photons · m-2 · s-1) of T. amphioxeia displaying different pigment contents and at two different prey: predator ratios to study the effect of prey physiology and quantity on the ciliate.

II-1-6) Materials and methods

II-1-6-1) Full factorial design experiment on T. amphioxeia

II-1-6-1-1) Culture of T. amphioxeia

The cryptophyte Teleaulax amphioxeia (AND-A0710) was cultivated in L1 medium without silicate (L1-Si) at salinity 35 (R. Guillard and Hargraves, 1993). Cultures were maintained at 17.8 ± 0.6 °C, a light intensity of ~ 100 µmol photons · m-2 · s-1 provided by cool-white and pink fluorescent tubes (fluora and cool-white fluorescent light, Osram, Munich, Germany) and a 12: 12 light: dark (L: D) cycle (supplementary materials Table 21). T. amphioxeia was maintained in a semi-continuous culture (i.e. bi-weekly dilutions), allowing for constant physiological conditions. Cultures were not axenic.

II-1-6-1-2) Factorial design

The direct effects, interactions and optima of temperature, irradiance and pH on the maximum growth rate (µmax), the maximum quantum yield of the photosystem II (Fv/Fm) and the pigment content of T. amphioxeia were studied using a central composite design (23*, supplementary materials Equation 1). Five levels were used for each factor to estimate the second order quadratic component of the relationship between a factor and the three parameters. After the determination of a central value, limits and axial points (i.e. star points) for each factor (Table 18), the 17 required

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Chapter 2-Part 1: Factorial design on Teleaulax and predator feeding measurements (i.e. 15 experimental conditions with a triplicate for the central one) were performed thanks to a culture device consisting of 17 photo-bioreactors placed in a software-controlled incubator. Each photo-bioreactors was thermo-regulated by a heater connected to a temperature sensor, light was supplied by a xenon lamp and pH was measured using a pH electrode (Mettler- ® Toledo ) and controlled by CO2 injections (Marchetti et al. 2012). As pH was only controlled by injection of CO2, it was only possible to limit the increase in pH during the light period; overall variations in pH did not exceed the regulated pH by 0.5 unit.

The day of the experiment, the photo-bioreactors were sterilized with a solution of 0.5% of DEPTIL PA 5 (Hypred SAS, Dinard, France) and thoroughly rinsed with culture medium. The photo- bioreactors were thereafter filled with 150 mL of inoculum at a concentration of 3.5 × 105 cells  mL-1 and randomly placed in the culture device with a 12: 12 (L: D) cycle. A bi-daily sampling of 1 mL of each culture was used for cell counting. During the exponential growth phase, 10 mL of each photo-bioreactor were sampled for pigment analysis and Fv/Fm measurements.

Table 18: Factor levels in the factorial design experiment, where α is the axial distance between star points and the center of the experimental domain

Factors - α - 1 0 + 1 + α

Temperature (°C) 13.0 15.4 19.0 22.6 25.0

pH 6.5 6.9 7.6 8.3 8.6

Irradiance (µmol photons· m-2 · s-1) 40 194 420 645 800

II-1-6-2) Effect of the prey on M. rubrum

II-1-6-2-1) Photoacclimation of T. amphioxeia

Semi-continuous cultures in flasks were established in order to acclimate T. amphioxeia to two light conditions (Wood et al., 2005). The low light (LL; 100 µmol photons · m-2 · s-1) and high light (HL; 400 µmol photons · m-2 · s-1) conditions were chosen based on the results of the factorial design experiment (supplementary materials Equation 1 and Table 22) to induce contrasting µmax,

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Fv/Fm and pigment contents. Temperature was set according to the optimal growth rate conditions (i.e. 17.6 °C) and pH was uncontrolled as previous experiments indicated that pH did not significantly influence µmax, Fv/Fm and pigment contents. Growth was monitored every day and cultures were diluted every two days by adding fresh L1-Si medium. The maximum growth rate, pigment content and Fv/Fm were measured to monitor the acclimation of the cultures (Wood et al., 2005).

II-1-6-2-2) Feeding experiment

The ciliate Mesodinium rubrum (AND-A0711) was routinely maintained in sterilized sea water in the same conditions as T. amphioxeia (supplementary materials Table 21) and fed three times a week at a ratio of 1: 1 (prey: predator). The ciliates were starved one week before the experiment to reduce the number of plastids. The day of the experiment, 80 mL M. rubrum cultures at a concentration of 5 × 103 cells  mL-1 were fed T. amphioxeia acclimated at LL or HL conditions and at a prey: predator ratio of 1: 1 (low fed LL or HL) or 10: 1 (high fed LL or HL). In addition, three controls were used, one unfed culture of M. rubrum and two cultures of T. amphioxeia previously acclimated to LL and HL conditions but maintained in sterilized sea water (i.e. without L1-Si medium enrichment) as for M. rubrum. All the cultures were placed in a culture chamber at a temperature of 17.6 °C and an irradiance of 100 µmol photons · m-2 · s-1 (i.e. corresponding to the LL condition). The monitoring of cell growth, pigment content and Fv/Fm was performed during the exponential growth phase of M. rubrum and for the control cultures.

II-1-6-3) Experimental set-up

II-1-6-3-1) Counting and growth rate

Counting of T. amphioxeia during the factorial design experiment was directly performed on fresh samples by flow cytometry on a Accuri C6 flow cytometer (Becton Dickinson Accuri™, Oxford, UK) equipped with blue and red lasers (488 and 640 nm), detectors of forward (FSC) and side (SSC) light scatter, and fluorescence detectors: 585 ± 20 nm (FL2 and 675 ± 12.5 nm (FL4). FL2 vs FL4 channel density plots, corresponding to phycoerythrin and chlorophyll a, were used to count T. amphioxeia, using Accuri™ C6 software. Counting during the semi-continuous experiment and

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Chapter 2-Part 1: Factorial design on Teleaulax and predator feeding the feeding experiment were performed on a particle counter equipped with a 100 µm aperture tube (Multisizer 3, Coulter Counter, Beckman, Paris, France).

The maximum growth rates were calculated from the slope of the linear regression for the natural logarithm-transformed values of population size during the time interval of exponential growth phase (i.e. ranging from 2 to 4 days for both species) (Guillard 1973).

II-1-6-3-2) The maximum quantum yield of the photosystem II (Fv/Fm)

Fv/Fm is considered to be a proxy of algal health (Kromkamp and Forster, 2003; Moeller et al., 2011; Woźniak et al., 2002) and was assessed with the Pulse Amplitude Modulation (PAM) method (Schreiber et al., 1986) in a Phyto-PAM (Walz GmbH, Effeltrich, Germany).

II-1-6-3-3) Pigment analysis

Pigment concentrations were measured by filtering 3 mL of cultures onto 25 mm Whatman Gf/F filters (Whatman, Sigma-Aldrich, Maidstone, UK). Filters were immediately frozen in liquid nitrogen and stored in the dark at -80 °C (Zapata et al., 2000). The analysis of pigments was performed by using HPLC with UV or fluorescence detection as previously described by Ras et al. (2008). Total chlorophyll a (TChl a) (sum of chlorophyll a and chlorophyllid a), chlorophyll c (Chl c) and total carotenoids (TCarotenoids) (sum of alloxanthin, crocoxanthin and α-carotene) were expressed on a per cell basis (pg · cell-1) of T.amphioxeia or M. rubrum. The hydrosoluble phycoerythrin, which is a typical pigment of Cryptophyceae (Jeffrey et al., 2011), was not measured in this work.

II-1-6-3-4) Statistical analyses

Statgraphics v 18.1.02 was used to analyze the full factorial design experiment and statistical analyses were computed on RStudio v 1.1.463. After checking the assumptions of independence (Durbin-Watson test), homoscedasticity (Bartlett test) and normality (Shapiro-Wilk test) of the residuals, direct effects of temperature, irradiance and pH and their interactions were investigated using two-way ANOVA for the factorial design experiment. For the other experiments, t-test or one-way ANOVA followed by a Tukey post hoc test were performed. Otherwise Mann-Withney U or Kruskal-Wallis tests were used, followed by a Conover test. Differences were considered

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Chapter 2-Part 1: Factorial design on Teleaulax and predator feeding statistically significant when P < 0.05, for a significance level of α = 0.05. Values are expressed as mean ± SD. Experiments were performed in triplicate.

II-1-7) Results

II-1-7-1) Direct effects, interactions and optimum of temperature, irradiance

and pH on the physiology of T. amphioxeia

II-1-7-1-1) Effect on the maximum growth rate

3* According to the 2 experimental design, the model of µmax (µmax th) explained 90% of the observed variability (regression coefficients and equation of the model for µmax th are shown in supplementary materials Equation 1 and Table 22). Both significant linear and quadratic effects of temperature (one-way ANOVA, F2,14 = 9.51, P = 0.02) and irradiance (one-way ANOVA, F2,14 = 7.31, P = 0.03) were observed on µmax (Figure 22 A) while pH was not significant across the experimental domain (Figure 22 A-B). In addition, a significant interaction (two-way ANOVA,

F2,14 = 10.93, P = 0.01) between temperature and irradiance (Figure 22 C) was noted, with a positive effect of temperature on µmax at low irradiance and the opposite effect under high irradiance. The predicted value of µmax for T. amphioxeia was 0.88 d-1, obtained for a temperature of 17.6 °C, a pH of 7.6 and an irradiance of 400 µmol photons · m-2 · s-1 under a circadian cycle 12: 12 (L: D)

(Figure 22 D). After the experiment, µmax th was checked under the predicted optimal conditions using three photo-bioreactor replicates in the same culture device and the µmax obtained was in very good agreement with the predicted growth rate (0.873 ± 0.003 d-1).

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A B 0.88 I-I T-T * *

) 0.78 1

T-I - T I 0.68 pH-I

T-pH (d rate Growth 0.58 pH-pH pH 0.48 0 1 2 3 4 5 6 15.4 22.6 6.9 8.3 194 646 Standardized effect T pH I C 1 D

0.80 * 1

)

) 1

1 0.80

- - 0.60 0.60 0.40 0.40 0.20

0 Growth rate (d rate Growth 0.20 (d rate Growth -0.20 800 -0.40 600 0 400 12 15 200 I 15.4 22.6 15.4 22.6 6.9 8.3 18 21 24 27 0 T T-pH T-I pH-I

II-1-7-1-2) Effect on the Fv/Fm and the pigment content

Models of maximum quantum yield of the photosystem II (Fv/Fm th), total chlorophyll a (TChl a th), chlorophyll c (Chl c th) and total carotenoids (TCarotenoids th) explained 98%, 69%, 69% and 67% of the observed variability, respectively. Briefly, across the experimental domain Fv/Fm th was influenced by the same factors as µmax (i.e. optimal Fv/Fm th at intermediate temperature and irradiance) whereas optimum of TChl a th and Chl c th were obtained at low irradiance (supplementary materials Figure 23). We further checked independently and in triplicate the accuracy of predicted modeled values using the conditions corresponding to µmax th. The measured values were in good agreement with predicted ones for Fv/Fm th, TChl a th and Chl c th but not for

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TCarotenoids th. Regression coefficients and equation of the models for Fv/Fm th, TChl a th and

Chl c th were shown in supplementary materials Equation 1 and Table 22.

II-1-7-2) Light acclimation of T. amphioxeia

After 27-30 generations, µmax of LL and HL-acclimated T. amphioxeia cultures were stable but -1 significantly higher in HL condition (0.85 ± 0.09 vs. 0.77 ± 0.10 d , t-test, T1,14 = 2.30, P = 0.03;

Table 2). However, TChl a and Chl c contents were significantly (t-test, T1,1 = 9.75 and 9.27 respectively, P = 0.001) ca. twice higher in T. amphioxeia grown in LL while similar TCarotenoids contents were observed between the two light conditions (Table 19). Fv/Fm were high for both photoacclimated T. amphioxeia (> 0.6) but significantly higher for the LL-acclimated condition (t- test, T1,1 = 4.53, P = 0.01; Table 19). The maximum growth rate, TChl a, Chl c and Fv/Fm of the HL-acclimated culture of T. amphioxeia were close to the modeled values, whereas for the LL- acclimated culture, µmax was 1.75-fold higher and around 2-fold lower TChl a and Chl c than modeled values (Table 19).

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Acclimation conditions LL HL Generations 27 30 µmax 0.77 ± 0.10 0.85 ± 0.09

µmax th 0.44 0.88

Fv/Fm 0.68 ± 0.01 0.61 ± 0.02

Fv/Fm th 0.71 0.65 TChl a 0.41 ± 0.03 0.24 ± 0.01

TChl a th 0.78 0.24 Chl c 0.05 ± 0.01 0.03 ± 0.001

Chl c th 0.12 0.03 TCarotenoids 0.13 ± 0.01 0.10 ± 0.004

II-1-7-3) Feeding experiment

The maximum growth rates, maximum cellular concentrations and pigment contents of M. rubrum were not significantly different when using LL or HL-acclimated T. amphioxeia cultures (Table 20). However, these responses depended significantly on the nutrition ratio applied. Indeed, high- fed condition resulted in 1.5-fold higher µmax (t-test, T1,9 = 4.99, P = 0.001), 2.5 times higher maximum cellular concentrations (t-test, T1,9 = 15.09, P < 0.001) and twice as high TChl a, Chl c and TCarotenoids (t-test, T1,4 = 8.89, 9.15 and 7.08, respectively, P < 0.001; Table 20). The Fv/Fm ranged from 0.68 ± 0.01 to 0.72 ± 0.02 and were not significantly different among the nutrition conditions. The unfed control of M. rubrum did not show a positive growth and had a significantly lower maximum cellular concentration Fv/Fm, TChl a, Chl c and TCarotenoids (one-way ANOVA,

F2,9 = 19.54, 55.18, 65.67, 70.48 and 37.22, respectively, P < 0.001, Table 20). The two T. amphioxeia controls maintained in sterilized sea water and previously acclimated to LL and HL conditions had similar µmax, Fv/Fm, TChl a, Chl c and TCarotenoids contents after three days of

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Chapter 2-Part 1: Factorial design on Teleaulax and predator feeding growth in LL condition. When compared to the semi-continuous cultures, the control LL culture of T. amphioxeia possessed 1.5 times less TChl a (t-test, T1,4 = 9.40, P = 0.001) and the control HL culture had a 1.2-fold lower µmax (t-test, T1,16 = 3.59, P = 0.002) whereas its Fv/Fm increased (t- test, T1,4 = 5.71, P = 0.005) (Tables 2 and 3).

Control M. High fed Control T. amphioxeia Nutrition Low fed rubrum not

conditions fed LL HL LL HL LL HL 0.31 ± 0.29 ± 0.20 ± 0.20 ± µmax - 0.76 ± 0.02 0.70 ± 0.01 0.04 0.03 0.03 0.04 Maximum concentration (× 19 ± 1.5 18 ± 1.8 7.6 ± 0.78 7.4 ± 0.42 5.8 ± 0.15 452 ± 10.1 407 ± 17.8 103) 0.69 ± 0.72 ± 0.68 ± 0.68 ± Fv/Fm 0.58 ± 0.02 0.68 ± 0.02 0.69 ± 0.01 0.01 0.02 0.01 0.01 0.26 ± 0.26 ± TChl a 3.1 ± 0.45 3 ± 0.5 1.4 ± 0.05 1.4 ± 0.08 1.1 ± 1.2 0.004 0.003 0.38 ± 0.37 ± 0.16 ± 0.16 ± 0.04 ± 0.04 ± Chl c 0.12 ± 0.02 0.05 0.06 0.01 0.02 0.001 0.001 0.57 ± 0.58 ± 0.10 ± 0.10 ± TCarotenoids 1.1 ± 0.17 1.1 ± 0.18 0.51 ± 0.10 0.02 0.04 0.002 0.002

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II-1-8) Discussion

II-1-8-1) Full factorial design experiment on T. amphioxeia and

photoacclimation in semi-continuous culture

The present study first investigated the effect of temperature, irradiance, pH and their interactions on the growth of Teleaulax amphioxeia thanks to a factorial design experiment. Beforehand, we tested a wide range of values for each factor (temperature 13-30 °C, irradiance 20-800 µmol photons · m-2 · s-1, pH 6-10) and tried to determine the factor levels (Table 18) as conditions allowing for growth in order to increase the robustness of obtained models.

Results of the 23* full factorial design underlined the importance of temperature and irradiance on growth, while pH was not significant on the strain AND-A0710 of T. amphioxeia (direct effects). Interestingly, a significant interaction between the two factors temperature and irradiance was also observed for the maximum growth rate. Generally, the interaction between these two factors results in µmax increasing with temperature under light saturation (Ojala 1993, Edwards et al., 2016, Wirth et al., 2019). However, our results showed that µmax was strongly affected at temperatures higher than the optimal one when irradiance was high. Hence the increase of temperature did not allow the strain to better cope with photoinhibition. The decrease in µmax beyond 400 µmol photons · m-2 · s-1 may suggest photoinhibition of the photosystem II. It has been shown for microalgae that an increase of temperature can reduce the carboxylase activity of the Rubisco (i.e. the catalytic enzyme of photosynthesis) while promoting the production of oxygen radicals that lead to oxidation of lipids and reaction centers of photosystem II (Kale et al., 2017; Ras et al., 2013) and ultimately to photoinhibition. We hypothesized that with our conditions of culture and especially because of the wide range of temperatures we applied, the increase of temperature here enhanced the effect of photoinhibition but further work is needed to understand how this interaction impacts T. amphioxeia.

The optimal maximum growth rate for this temperate strain was obtained for intermediate tested irradiance (400 µmol photons · m-2 · s-1) and temperature (17.6 °C), whereas a deviation towards high temperature and irradiance (>22.6 °C and 646 µmol photons · m-2 · s-1) led to lower µmax.

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The maximum growth rate was ca. 0.88 d-1 according to the factorial design and values measured from the independent triplicate verification.

As far as we know, the same strain of T. amphioxeia was used in two other studies with similar culture conditions. In the first one, Rial et al. (Rial et al., 2013) found a maximal µmax of 0.98 and 1.6 d-1 at 70 and 200 µmol photons · m-2 · s-1, respectively, also suggesting a high impact of light on growth under stress conditions, i.e. without photoacclimation. In the other study, García-Portela et al. (2018b) photoacclimated the cultures and did not observe a significant difference of µmax between 200 and 650 µmol photons · m-2 · s-1. This is similar to what we observed for our photoacclimated cultures, as µmax at 100 and 400 µmol photons · m-2 · s-1 were only 10% different (but still significantly different). The differences in µmax between the current study and the previous ones (García-Portela et al., 2018b; Rial et al., 2013) are likely explained by a different experimental setup. Nevertheless, our results and those from García-Portela et al. (2018b) confirmed the high photoacclimation ability of T. amphioxeia.

T. amphioxeia can also tolerate or acclimate to other abiotic factors. Indeed, Lee et al. (Lee et al., 2019) reported that several strains of T. amphioxeia isolated from cold and temperate waters (i.e. 5.4 to 28.9 °C) can all be acclimated to the same temperature in the lab (i.e. 20 °C). Our results also showed that T. amphioxeia was able to grow under stress conditions across all the experimental domain except under high temperature (> 22.6 °C), high irradiance (> 646 µmol photons · m-2 · s- 1) or the combination of both (i.e. n°13, 4, 11 and 16 in supplementary materials Table 23).

In addition, a previous lab study on a Danish strain of T. amphioxeia reported a positive growth at elevated pH 9.4 (Smith and Hansen 2007). Our results suggested a high tolerance to pH of the T. amphioxeia strain used, including for values already occasionally found in some coastal waters (e.g. < 8) (Feely et al., 2008). According to some predictions, in 2100, average pH in the world ocean would be around 7.7 (Brewer, 1997; Gattuso and Hansson, 2011; Haugan and Drange, 1996; Orr et al., 2005) and would thus not significantly impact the growth of T. amphioxeia. Altogether, these results suggest an important plasticity of the species which may explain why T. amphioxeia is found in diverse ecological niches.

It should be noted that we observed some discrepancies between values predicted by the surface response and values really observed during the factorial design experiment and for the photoacclimated cultures. For the HL-acclimated condition, the experimental µmax, Fv/Fm and

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Chapter 2-Part 1: Factorial design on Teleaulax and predator feeding pigment content fitted well with the predicted modeled values. However, for the LL-acclimated condition, observed and predicted values of µmax, TChl a and Chl c were not in agreement (Table 19). These differences may arise from the fact that this type of factorial design is performed to determine optimal conditions across an experimental domain and not extreme values (Lundstedt et al., 1998). Indeed, while there was almost no difference for the central points between predicted and measured values in the factorial design, we noted for the irradiance star point (supplementary materials Table 23 n°6) a difference of 57%, 5%, 30% and 29% for µmax, Fv/Fm, TChl a and Chl c, respectively. These levels of difference were reflected in the LL-acclimated culture between predicted and observed values. In addition, experiments were performed in stress condition while the semi-continuous cultures led to acclimation as reflected by the stable µmax, Fv/Fm and pigment contents after more than 27 generations. The presence of bacteria in the cultures of T. amphioxeia cannot be excluded. T. amphioxeia can feed on bacteria (Yoo et al., 2017), especially in light- limited conditions (Marshall and Laybourn-Parry, 2002), thus its mixotrophic ability might also explain the discrepancies observed in LL on growth rate and pigment content.

The factorial design experiment helped to better understand the physiological responses of the temperate strain of T. amphioxeia which belongs to the Teleaulax/Plagioselmis/Geminifera clade (Hansen et al., 2012). T. amphioxeia is an important donor of plastids to M. rubrum (García-Portela et al., 2018b; Hernández-Urcera et al., 2018; Johnson et al., 2018; Peterson et al., 2013) and thus indirectly to Dinophysis spp. (Park et al., 2006).Low light conditions (< 100 µmol photons · m-2 · s-1) coupled with intermediate tested temperature resulted in limited growth of T. amphioxeia

(while maintaining high pigment content and Fv/Fm) and helped prevent the organism “outgrowing” M. rubrum and Dinophysis spp. in routine laboratory cultures. Furthermore, controlling the pH of T. amphioxeia cultures may be not necessary as this species appears to tolerate pH from 6.0 (this study, data not shown) up to 9.4 (Smith and Hansen, 2007). Thanks to the factorial design experiment, the present study determined the optimal conditions to obtain a large biomass that may be useful to exploit the beneficial role of T. amphioxeia as a food enrichment (Peltomaa et al. 2018 and Lee et al. 2019). This approach also provided information on factors driving the bloom initiation of T. amphioxeia. In natural environments, T. amphioxeia has been found in higher concentration several meters below the surface (Peterson et al., 2013), indicating that the species easily moves in the water column. This field observation coincides with our experimental observation that high light intensity diminishes µmax. We thus hypothesized that T. amphioxeia

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Chapter 2-Part 1: Factorial design on Teleaulax and predator feeding can perform photosynthesis at a water depth of several meters and can thus escape conditions of higher light intensity at surface. Nonetheless, the intraspecific variability of the species should be explored, including strains from polar and temperate regions.

II-1-8-2) Feeding experiment of M. rubrum

M. rubrum is a mixotrophic organism which acquires and maintains photosynthesis by ingestion of cryptophyte plastids (Gustafson et al., 2000; Johnson et al., 2006; Yih et al., 2004) and cryptophyte nuclei (Johnson et al., 2007; Kim et al., 2017). As the contribution of carbon fixation through photosynthesis appears higher than the contribution of prey (Moeller et al., 2011), the aim of the nutrition experiment was to assess the effect of the physiology of the prey on its predator M. rubrum. We first photoacclimated T. amphioxeia in semi-continuous cultures and obtained a LL-acclimated culture with higher pigment content and Fv/Fm compared to the HL-acclimated culture. The pigment profile of photoacclimated T. amphioxeia was similar to the ones observed by Rial et al. (2013) and García-Portela et al. (2018) with the major lipophilic pigments being Chl a and alloxanthin. Alloxanthin to TChl a ratio was higher in HL-acclimated culture, as already shown for Rhodomonas salina (Schlüter et al., 2000), but in our study it was due to a decrease in TChl a, thus not supporting the reported photoprotection role of alloxanthin in T. amphioxeia (Laviale and

Neveux, 2011; Roy et al., 2011). Values of Fv/Fm suggested a good cellular health (Moeller et al., 2011) in both LL and HL acclimations, with a significantly higher value in the LL-acclimated compared to the HL-acclimated culture (0.68 ± 0.01 and 0.61 ± 0.02), as already observed by García-Portela et al. (2018b) with the same strain of T. amphioxeia. Overall, differences obtained in terms of pigment contents and Fv/Fm suggested that the LL-acclimated culture of T. amphioxeia may be a better quality food source in term of photosynthetic capacity for M. rubrum

However, feeding with prey of different physiology (i.e. LL or HL-acclimated T. amphioxeia) yielded no significant effect on µmax, Fv/Fm and pigment content of M. rubrum. Unfortunately, the control of T. amphioxeia acclimated to the HL condition had already converted its pigment content to a content equivalent to the LL condition after 3 days. Therefore the difference of physiological status of the preys was not maintained during all the feeding experiment and to what extent this influenced the results should be further elucidated. Interestingly, this observation highlights

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Chapter 2-Part 1: Factorial design on Teleaulax and predator feeding another evidence of the plasticity of T. amphioxeia, which can easily acclimate to different light conditions.

This study clearly confirmed the positive effect of prey quantity on M. rubrum, i.e. a high feeding ratio 10: 1 (prey: predator) yielded significantly higher µmax, maximum cellular concentrations,

Fv/Fm and pigment contents compared to the low feeding ratio 1: 1. Indeed, prey quantity had been shown to be beneficial for M. rubrum growth (Peltomaa and Johnson 2017), with a 75% increase in µmax for a feeding ratio of 44: 1 compared to 1: 1. These experimental studies corroborate the occurrence of Mesodinium spp. in situ being quantitatively related to the presence of T. amphioxeia (Hamilton et al., 2017; Herfort et al., 2011; Peterson et al., 2013). However, in the environment, different factors may also contribute to bloom development of Mesodinium spp. For instance, in the Columbia River estuary, development and structure of M. rubrum blooms may also be explained by changes in abiotic factors (e.g. increases of light intensity and dissolved organic compounds, a decrease of turbulence), or biotic factors (e.g. prey preference and availability), or a combination of those factors (Herfort et al., 2011; Peterson et al., 2013). Nonetheless, the impact of nutrient limitations and ratios on growth and photosynthetic activity of T. amphioxeia should be further investigated, as they also appeared to drive M. rubrum bloom initiation (Hamilton et al., 2017; Peterson et al., 2013) and directly impact growth of M. rubrum (Hattenrath-Lehmann and Gobler, 2015).

The preponderant effect of feeding ratio was also evident on pigment content, which was twice as concentrated in the high fed M. rubrum. Comparing maximal pigment contents between both M. rubrum and T. amphioxeia, the ciliate had around 8 times more pigments per cell. This observation is consistent with the 6 to 36 plastids of T. amphioxeia harbored by M. rubrum (Hansen and Fenchel 2006). However, there is probably a high intra-specific variability between strains of M. rubrum in terms of behavior, size and prey preference, possibly related to different haplotypes (Herfort et al. 2011), thus extrapolation of results should be done with care.

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II-1-9) Conclusion

This study shows the impact of two key variables of global change (temperature and irradiance) on the physiology of the cryptophyte T. amphioxeia, which is one of the first level organism of the trophic chain of Dinophysis spp. Also, importantly, pH appeared to not impact on growth of at least the strain in this study. While a slight increase of irradiance and temperature would lead to an increased concentration of T. amphioxeia, a negative interaction was observed for high temperature combined with high irradiance. It is not evident whether such a condition is of high environmental relevance for an organism which has been observed to occur at several meter water depth. This study suggests that future climate conditions appear not detrimental to T. amphioxeia. An increase of T. amphioxeia abundance would favour M. rubrum growth and pigment content, which in turn might lead to increased occurrence of Dinophysis spp.

II-1-10) Acknowledgements

This work was funded by the project CoCliME which is part of ERA4CS, an ERA-NET initiated by JPI Climate, and funded by EPA (IE), ANR (FR), BMBF (DE), UEFISCDI (RO), RCN (NO), and FORMAS (SE), with co-funding by the European Union (Grant 690462). We thank David Jaén (LCCRRPP, Huelva, Spain) for T. amphioxeia and M. rubrum strains, Céline Dimier and Joséphine Ras from the SAPIGH analytical platform of the Laboratoire d’Océanographie de Villefranche (CNRS-France).

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II-1-11) Supplementary materials

Equation 1: The general quadratic model fitted to the data for theoretical maximum growth rate

( max th).

2 2 2 Model th = ß0+ß1 X1+ ß2 X2+ ß3X3+ ß12 X1X2+ ß13 X1X3+ ß23 X2X3+ ß11X1+ ß22X2+ ß33X3+ ε

Here, ß0 is the model error, ßi , ßii and ßij are the model coefficients (Table 22), Xi is the main effect 2 of the factor I, Xij is the interaction between the factors i and j, Xi is the quadratic effect of the factor i and ε is the residual error. X1, X2 and X3 were chosen for temperature, irradiance and pH, respectively. Noted that Equation 1 could be used for the calculation of µmax th, Fv/Fm th, TChl a th and Chl c th.

A B I I T I-I T-I T T-T T-T I-I T-I pH-pH pH T-pH T-pH pH-I pH-pH pH pH-I 0 2 4 6 8 10 12 0 0.5 1 1.5 2 2.5 3 Standardized effect Standardized effect C I I-I T-T T T-I pH-pH pH T-pH pH-I 0 0.5 1 1.5 2 2.5 3 Standardized effect

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Table 21: Culture conditions of strains used in this study.

Temperature Irradiancea (µmol Species Origin Strain ID Medium (°C) photons · m-2 · s-1)

Teleaulax Huelva AND-A0710 L1-Si 17.8 ± 0.6 ~100 amphioxeia (Spain)

Mesodinium Huelva Sterilized AND-A0711 17.8 ± 0.6 ~100 rubrum (Spain) seawater

aCultures were subjected to light in the PAR domain during a circadian cycle 12 h: 12 h (light: dark).

Table 22: Regression coefficients for the models of maximum growth rate (µmax th), maximum quantum yield of the photosystem II (Fv/Fm th), total chlorophyll a (TChl a th) and chlorophyll c

(Chl c th), where β0 is the model error, 1 is for temperature, 2 for pH and 3 for irradiance

Notations terms µmax th Fv/Fm th TChl ath Chl cth

ß0 -8.39136 1.43632 -3.72339 -7.07201E-01

ß1 6.41200E-01 2.28393E-01 2.81699E-01 3.84370E-02

ß2 6.21468E-01 -8.52622E-01 5.88921E-01 1.42017E-01

ß3 6.23805E-03 3.94179E-03 -2.20618E-03 -4.36766E-04

ß12 1.49100E-02 4.93690E-03 -9.27244E-03 -6.37145E-04

ß13 2.36085E-04 -1.75225E-04 -4.18822E-05 -5.52009E-06

ß23 2.36466E-04 -5.26063 -3.22218E-05 2.26091E-06

ß11 -1.22927E-02 -6.25934E-03 -5.63233E-03 8.96128E-04

ß22 -2.96434E-02 5.15398E-02 -2.85943E-02 8.96177E-03 ß 4.87390E-06 1.31250 2.74671E-06 4.52000E-07 33

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Irradiance TChl a Chl c Temperature (µmol µmax TCarotenoids n° pH Fv/Fm (pg · (pg · (°C) photons (d-1) (pg · cell-1) cell-1) cell-1) · m-2 · s-1) 1 19 7.6 420 0.84 0.59 0.22 0.03 0.12 2 15.4 8.2 194 0.59 0.69 0.36 0.05 0.14 3 15.4 6.9 646 0.56 0.6 0.31 0.04 0.15 4 19 7.6 800 - - - - - 5 22.6 6.9 194 0.86 0.65 0.34 0.04 0.16 6 19 7.6 40 0.11 0.69 1.37 0.22 0.71 7 19 6.4 420 0.71 0.61 0.26 0.03 0.12 8 15.4 8.2 646 0.62 0.58 0.18 0.03 0.14 9 19 7.6 420 0.86 0.59 0.20 0.03 0.12 10 15.4 6.9 194 0.58 0.71 0.29 0.05 0.16

11 22.6 8.2 646 - - - - -

12 13 7.6 420 0.62 0.61 0.16 0.02 0.08 13 25 7.6 420 - - - - - 14 22.6 8.2 194 0.67 0.71 0.09 0.02 0.04 15 19 8.6 420 0.75 0.58 0.26 0.03 0.10 16 22.6 6.9 646 - - - - - 17 19 7.6 420 0.86 0.59 0.16 0.02 0.07

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II-2) Effect of a 24 h salinity stress on the growth, biovolume, toxins, osmolytes and metabolite profiles on three strains of the Dinophysis acuminata-complex (“D. sacculus”)

This Chapter 2-Part 2, has been submitted in Harmful Algae in September 2020:

Gaillard, S., Réveillon, D., Danthu, C., Hervé, F., Sibat, M., Carpentier, L., Hégaret, H., Séchet, V., Hess, P., 2020. Effect of a 24 h salinity stress on the growth, biovolume, toxins, osmolytes and metabolite profiles on three strains of the Dinophysis acuminata-complex (“D. sacculus”). Submitted in Harmful Algae (2020-09-01)

II-2-1) Graphical abstract

Hypo- hyperosmotic stress  Osmolytes (42) . Osmoregulation 25 35 (control) 42 = µ, volume, . toxins and Metabolomic metabolites . Tolerance Isolation 3 strains “Dinophysis sacculus” Intraspecific variability

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II-2-2) Résumé

La microalgue Dinophysis est le principal dinoflagellé responsable de l'intoxication diarrhéique par les mollusques (DSP) chez l’Homme, due à la consommation de bivalves filtreurs contaminés par des toxines diarrhéiques lipophiles. La distribution de ce genre est mondiale et elle est déterminée par exemple par la température, l'irradiance, la salinité et les nutriments. Ces facteurs sont sensibles au changement climatique. Le complexe D. acuminata peut contenir plusieurs espèces, dont D. sacculus. Ce dernier a pu être observé dans les estuaires, les zones semi-fermées et les plans d'eau soumis à de rapides variations de salinité. De plus, sa répartition naturelle suggère une certaine tolérance à la salinité. Cependant, les réponses des souches du complexe D. acuminata (« D. sacculus ») soumises à un stress de salinité et les mécanismes sous-jacents n'ont jamais été étudiés en laboratoire. Dans cette étude, des stress hypoosmotique (25) et hyperosmotique (42) de 24 h ont été réalisés in vitro sur trois souches cultivées de « D. sacculus », isolées des côtes atlantique et méditerranéenne françaises. Le taux de croissance, le biovolume, les osmolytes (proline, glycine bétaïne et diméthylsulfoniopropionate (DMSP)) et les toxines ont été mesurés et une approche métabolomique a été réalisée. Les teneurs en osmolytes étaient plus élevées à la salinité la plus élevée, mais seule une augmentation significative de la glycine bétaïne a été observée entre le contrôle (35) et l'état hyperosmotique. L’approche métabolomique a révélé des profils significativement différents pour différentes salinités et dépendants de la souche étudiée. Ces résultats, ainsi que l'absence d'effets sur le taux de croissance, le biovolume, les teneurs en acide okadaïque (OA) et en pecténotoxines (PTXs), suggèrent que les souches de « D. sacculus » étudiées sont très tolérantes aux variations de salinité, ce qui pourrait être un avantage dans leurs habitats et dans le contexte du changement climatique.

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II-2-3) Abstract

Dinophysis is the main dinoflagellate responsible for diarrheic shellfish poisoning (DSP) in human consumers of contaminated filter feeding bivalves by lipophilic diarrheic toxins. This genus has a worldwide distribution driven by e.g. temperature, irradiance, salinity and nutrients, which are sensitive to climate change. The D. acuminata-complex may contain several species, including D. sacculus. This latter has been found in estuaries and semi-enclosed areas, water bodies subjected to quick salinity variations and its natural repartition suggests some tolerance to salinity. However, the responses of strains of D. acuminata-complex (“D. sacculus”) subjected to salinity stress and underlying mechanisms have never been studied in the laboratory. Here, a 24 h hypoosmotic (25) and hyperosmotic (42) stress was performed in vitro on three cultivated strains of “D. sacculus” isolated from French Atlantic and Mediterranean coasts. Growth rate, biovolume, osmolytes (proline, glycine betaine and dimethylsulfoniopropionate (DMSP)), and toxins were measured and a metabolomic approach was performed. Osmolyte contents were higher at the highest salinity, but only a significant increase in glycine betaine was observed between the control (35) and hyperosmotic condition. Metabolomics revealed significant and strain-dependent differences in metabolite profiles for different salinities. These results, as well as the absence of effects on growth rate, biovolume, okadaic acid (OA) and pectenotoxin (PTXs) contents, suggested that the “D. sacculus” strains studied are highly tolerant to salinity variations, which could be an advantage in their habitats and in the context of climate change.

II-2-4) Key words

Dinoflagellates; DMSP; glycine betaine; okadaic acid; pectenotoxins; proline

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II-2-5) Introduction

Diarrhetic shellfish poisoning (DSP) - a human intoxication due to the consumption of contaminated mollusks - is characterized by nausea, abdominal pain, vomiting and diarrhea (Yasumoto et al., 1985). This intoxication is mainly caused by lipophilic toxins synthesized by toxic species of the genus Dinophysis (Ehrenberg, 1841), namely okadaic acid (OA) and dinophysistoxins (DTXs). In addition, some species can also produce pectenotoxins (PTXs), another group of non-diarrheic toxins ((Ito et al., 2008; Reguera et al., 2012). Since 1976 (Yasumoto et al., 1978, 1980), worldwide observations of DSP outbreaks and indirect impacts on human health have been reviewed (Reguera et al., 2014). Consequently, national surveillance programs have been implemented in many countries, as the REPHY/REPHYTOX (hereafter named REPHY) in France that monitors toxic phytoplankton species in seawater since 1984 and their toxins in seafood (Belin et al., 2020). Shellfish farming and recreational harvesting are forbidden when the concentration of regulated toxins exceeds sanitary thresholds (e.g. a limit of 160 µg OA eq. per kg of fresh whole bivalves meat in Europe, America and Japan (DeGrasse and Martinez-Diaz, 2012; EU.Commission, 2011; Hess, 2012; National Shellfish Sanitation Program, 2017; Suzuki and Watanabe, 2012).

Moreover, recent studies highlighted direct effects of Dinophysis spp. on bivalve shellfish species of a commercial interest, such as hypersecretion of mucus and paralysis of the Japanese and noble scallops (Basti et al., 2015b), reduction of clearance rate of the blue mussel (Nielsen et al., 2020) or an increase of oocyte mortality and a decrease of fertilization success of the Pacific oyster (Gaillard et al., 2020b). Altogether, the effects of Dinophysis blooms could lead to environmental, sanitary, societal and economic issues (Reguera et al., 2012; Van Dolah, 2000).

Numerous field studies investigated the effect of salinity variation on the abundance and distribution of toxic species of the genus Dinophysis. Still, there is no consensus within the scientific community as both positive and negative significant correlations have been reported (Ajani et al., 2016; Alves et al., 2018; Caroppo et al., 2001; Hattenrath-Lehmann et al., 2015; Ninčević-Gladan et al., 2008). Moreover, most likely related to their mixotrophic capacity, Dinophysis spp. are able to thrive at significant depth, i.e. well below the euphotic zone (Fux et al., 2010), and can be found in salinity stratified-areas (Alves-de-Souza et al., 2019; Diaz et al., 2011; Ninčević-Gladan et al., 2008; Peperzak et al., 1996), over a wide range of salinities (Alves and

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Mafra, 2018; Hoshiai et al., 2003), e.g. D. acuminata was found at salinities ranging from 5 to 31 in Chile (Diaz et al., 2011).

Since the successful mixotrophic culture of D. acuminata by Park et al. in 2006, several laboratory studies have been carried out on Dinophysis spp. about the effects of (1) abiotic factors, such as nutrient availability, concentration and uptake (e.g. Nagai et al. 2008b, Hattenrath-Lehmann and Gobler 2015, Hattenrath-Lehmann et al., 2015, Tong et al., 2015a, Gao et al., 2018, García-Portela et al., 2020), temperature (Basti et al., 2015c, 2018; Gao et al., 2017; Kamiyama et al., 2010), salinity (Fiorendino et al., 2020), light intensity and quality (e.g. García-Portela et al., 2018b; Hansen et al., 2016; Kim et al., 2008; Nielsen et al., 2012) or (2) biotic factors, such as prey type (Gao et al., 2017; García-Portela et al., 2018a; Smith et al., 2018), prey concentration (Hattenrath- Lehmann and Gobler, 2015; Kim et al., 2008; Smith et al., 2018) and prey exudates (Gao et al., 2019; Nagai et al., 2011), nutritional status (García-Portela et al., 2020) or (3) geographical origin (Fux et al., 2011; Tong et al., 2015b) on the physiology and/or toxicity, whereas laboratory studies exploring the effect of salinity stress on physiology and metabolites production (including toxins) are still lacking.

D. acuminata and D. sacculus were described by morphology only, but several studies pointed out the considerable morphological plasticity (Lassus and Bardouil, 1991; Zingone et al., 1998). Genetically, the sequences are closely related and do not support the distinction into several species. For instance, D. ovum, another member of the complex is now considered as possibly conspecific with D. acuminata (Park et al., 2019). For D. sacculus, although it seems to have a different ecological niche compared to D. acuminata, its taxonomical identity is not clear and genetic data could not separate it clearly from other species of the complex (Séchet et al., 2020).

Indeed, D. sacculus found on European Atlantic coasts and in the Mediterranean Sea, is likely to perform well in closed or semi-enclosed areas, e.g. lagoons, rias or bays (Escalera et al., 2010; Ninčević-Gladan et al., 2008; Pizarro et al., 2008; Zingone et al., 1998), where shellfish farming activities are located. In coastal waters, salinity can change over short time spans due to runoff waters, precipitation, evaporation and extreme events related to global changes (Kirst 1989, IPCC 2013, Wells et al. 2019). Furthermore, salinity is likely to increase in the future in North Atlantic temperate and Mediterranean waters (up to 0.4 in the Mediterranean), notably as the concomitant result of an increase of evaporation and a decrease of precipitation (Skliris et al., 2014).

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Salinity stress can affect microalgae, impacting their osmotic potential, ion ratios (uptake and loss) due to the difference in osmolarity between the internal cellular and external environment (Kirst, 1989). In order to maintain an equilibrium, cells rely on different physiological responses to a salt stress, as the modification of growth rate, inhibition of photosynthesis or respiration, reactive oxygen species (ROS) production and osmoregulation (Borowitzka, 2018; Mayfield and Gates, 2007). This last process has been described as a three-step mechanism: microalgae can (1) adjust their turgor pressure by fluxes of water, (2) adjust their ion concentrations and (3) synthesize osmolytes (i.e. compatible solutes) (Hagemann, 2016, 2011; Kirst, 1989), which are typically small, water-soluble compounds containing nitrogen and sulfur (Keller et al., 2004; Rhodes et al., 2002). The most commonly found osmolytes are amino acids (e.g. proline) (Hagemann, 2016; Stefels, 2000), quaternary ammonium derivatives (glycine betaine, GBT) (Dickson and Kirst, 1986), as well as the tertiary sulfonium compound dimethyl sulfonio-propionate (DMSP) (Keller et al. 1999), which has been found in massive blooms of coccolithophores (Malin et al., 1993) but is also commonly produced by dinoflagellates (Caruana and Malin, 2014). However, no laboratory studies addressed the production of proline, GBT and DMSP by Dinophysis spp. and especially in a context of sudden salinity variations.

In addition to the quantitation of known metabolites, untargeted LC-HRMS approaches such as metabolomics (i.e. metabolic fingerprinting) can be used to study the response organisms, including microalgae to salinity changes. Indeed, metabolomics allows to get a fingerprint of a large set of small metabolites affected by the environmental changes in any organism (Bundy et al., 2009; Fiehn, 2002). Other molecules acting like osmolytes have been described (Rhodes et al., 2002) and characterized after salinity changes by liquid chromatography coupled to high resolution mass spectrometry (e.g. sucrose) (Georges des Aulnois et al., 2020).

In this study, three well-fed “D. sacculus” strains isolated from French coasts (two semi-enclosed and one estuarine species) and in exponential growth were subjected to 24 h hypoosmotic and hyperosmotic stresses. In addition to growth and biovolume measurements, toxins and osmolytes (proline, GBT and DMSP) were quantified using ultra high performance liquid chromatography coupled to low resolution tandem mass spectrometry (LC-LRMS/MS) to determine “D. sacculus” putative mechanisms of osmoregulation under salinity stress conditions. Finally, a non-targeted

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II-2-6) Materials and methods

II-2-6-1) Culture maintenance

Three strains of D. acuminata-complex isolated from the French Atlantic coast at Loscolo (Vilaine Bay, strain IFR-DSA-01Lo = Dsa-Lo) and Meyran (Arcachon Bay, strain IFR-DSA-01Me = Dsa- Me) and one strain from Thau Lagoon (Crique de l’Angle, strain IFR-DSA-02Th=Dsa-Th) in the Mediterranean Sea were used. Characterization of these strains has been realized by Séchet et al. (2020) who investigated morphology and genetics and showed that they all clustered in the subclade B in the cox1 phylogenetic analysis, corresponding presumably to “D. sacculus”.

This mixotrophic species was cultivated according to a classic three-step culture method (Park et al., 2006) with the ciliate Mesodinium rubrum (Lohmann, 1908) (strain MBL-DK2009) and the cryptophyte Teleaulax amphioxeia (Conrad) Hill (Hill, 1992) (strain AND-0710) as described in Gaillard et al. (2020). Dinophysis cultures were maintained in sterilized seawater at salinity 35, 17.8 ± 0.6 °C, a light intensity of ~ 100 µmol photons m-2 s-1 with a circadian cycle of 12: 12 (L: D; supplementary materials Table 25). All the cultures were monoclonal and xenic.

II-2-6-2) Experimental design

Cultures of “D. sacculus” in exponential growth phase were filtered on a nylon sieve (mesh 11 µm) and gently rinsed with sterilized seawater to remove any cryptophyte and ciliate cells. Each strain was then resuspended in nine replicates of 10 mL in sterilized seawater (salinity 35), at a cell density of 2200 ± 350 cells mL-1 and fed M. rubrum at a ratio of 6: 1 (prey: predator). Growth was monitored for seven days until the total consumption of M. rubrum and the exponential growth phase was reached.

Then, for each strain, the same volume of either milliQ water or salted-milliQ water was added in three replicates to decrease the salinity down to 25, increase the salinity up to 42 and maintain the

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Chapter 2-Part 2: Salinity stress on D. sacculus salinity at 35 (control condition). NaCl was chosen because it is the main dissolved salt in natural seawater (Hagemann, 2016).

Salinities 25 and 42 correspond approximately to the minimum (at Loscolo) and maximum (at Thau Lagoon) salinities observed in the field sampling locations (Figure 24) (REPHY 2019). Moreover, a preliminary experiment on the same three strains revealed no positive growth rate at salinity 45.

One day (24 h) after the salinity stress, cultures were filtered on a nylon sieve (mesh 11 µm) and the remaining supernatants were filtered on 0.2 µm (cellulose) syringe filters to remove any potential cryptophyte and ciliate or bacteria. Filters, with “D. sacculus” cells were stored at -80 °C for further toxin and metabolomic analyses. Supernatants, stored at -80 °C, were analyzed for their toxin contents.

42.5

40.0

37.5

35.0 Salinity 32.5

30.0

27.5

25.0

Loscolo Meyran-Arcachon Thau Lagoon Locations

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Figure 24: Box plot of the sea water salinity at the three locations where the studied strains of “D. sacculus” were isolated. The 1st and 3rd quartiles are represented by the lower and upper limits of the box, respectively, with the median in bold horizontal line. Error bars represent both lower and higher values within 1.5 times interquartile range below the 1st and above the 3rd quartiles, with outside values (plain dot) below or above these limits. Mean salinities are represented by red circles. Data were extracted from the REPHY database from 2015 to 2018 and correspond to weekly (September to May) or biweekly (June-July-August) salinity measurements of sea water samples by a conductivity meter.

II-2-6-3) Growth rate calculation and biovolume measurement

Two daily samples of 90 L of “D. sacculus” cultures were fixed in acidic Lugol solution (1% final concentration) and a minimum of 100 cells of “D. sacculus” and M. rubrum (before full consumption) were counted using a Nageotte cell counting chamber with a light microscope. The maximum growth rates were calculated from the slope of the linear regression for the natural logarithm-transformed values of population size during the time interval of exponential growth phase (Guillard, 1973).

To assess the biovolume, at least 30 individuals of each “D. sacculus” strain were pictured under a light microscope (Leica DMR, Germany). Cells of “D. sacculus” correspond to a flattened ellipsoid (Olenina et al., 2006), thus the biovolume was calculated after measuring height and large diameter using ImageJ software. The small diameter was estimated according to Olenina et al. (2006).

II-2-6-4) LC-LRMS/MS and LC-HRMS

Intracellular metabolites from cells were extracted with methanol (0.5 mL for 25x103 cells, i.e. between 0.5 to 1.4 mL methanol were added) and sonicated at 25 kHz for 15 min. Extracellular metabolites were recovered from the supernatant after liquid-liquid extraction with dichloromethane, evaporated under a flow of nitrogen (Gaillard et al., 2020b) and resuspended in 0.5 to 1.4 mL of methanol. All the samples were filtered (0.2 µm, Nanosep, MF, Pall) before analysis.

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II-2-6-4-1) Toxin analysis

Toxins were quantified as in Gaillard et al. (2020). Analysis was performed by liquid chromatography coupled to low resolution tandem mass spectrometry (LC-LRMS/MS) with a UHPLC system (UFLC XR Nexera, Shimadzu, Tokyo, Japan) coupled to a triple quadrupole/ion- trap mass spectrometer (API 4000 QTrap, Sciex, Redwood City, CA, USA), equipped with a turboV® electrospray ionization (ESI) source. Certified calibration solutions of pectenotoxin 2 (PTX2), okadaic acid (OA), dinophysistoxin 1 and 2 (DTX1 and DTX2) were obtained from the National Research Council Canada (NRCC, Halifax, NS, Canada). Intracellular pectenotoxin 2 equivalent (PTX2eq) was the sum of pectenotoxin 2, pectenotoxin 2b, pectenotoxin 2 seco-acid and 7-epi-pectenotoxin-2 seco-acid, all quantified with the PTX2 standard by assuming similar molar responses. Limit of detection/quantification were 1/3 ng mL-1 for OA, DTX1 and DTX2, and 0.2/0.6 for PTXs.

II-2-6-4-2) Proline, GBT and DMSP analysis

Quantification of intracellular proline, glycine betaine and DMSP was adapted from Georges des Aulnois et al. (2019) and performed on the analytical system used for toxin analysis. The chromatographic column was a Hypersil GOLD HILIC (hydrophilic interaction liquid chromatography) (150 x 2.1 mm; 3 µm; ThermoScientific, Waltham, MA, USA) and the flow rate was 0.25 mL min-1. Compounds were quantified using external 5-point calibration curves of standards (Sigma-Aldrich, Saint-Quentin Fallavier, France) solubilized in methanol, with concentrations from 50 nM to 5000 nM. As biovolumes were similar among conditions (see section “II-2-7-1) Growth rate and biovolume”), concentrations were expressed on a per-cell basis.

II-2-6-4-3) Metabolomics

Metabolomic profiles were acquired as in Estevez et al. (2020) by liquid chromatography coupled to high resolution mass spectrometry (LC-HRMS). The system was a UHPLC (1290 Infinity II, Agilent technologies, Santa Clara, CA, USA) coupled to a quadrupole-time of flight mass spectrometer (QTOF 6550, Agilent technologies, Santa Clara, CA, USA) equipped with a Dual Jet Stream ESI interface. Both positive and negative full scan modes of were used over a mass-to- charge ratio (m/z) ranging from 100 to 1700. The only difference was the injection volume of 10

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µL. Pool samples (QC) were prepared and injected ten times at the beginning of the batch sequence and then every five samples (including blanks). Blanks were prepared as “D. sacculus” samples.

LC-HRMS raw data (.d) were converted to .mzXML format using MS-Convert (ProteoWizard 3.0) (Chambers et al., 2012) and pre-processed with theWorkflow4Metabolomics (W4M; http://workflow4metabolomics.org) e-infrastructure (Guitton et al., 2017). Peak picking, grouping, retention time correction, and peak filling were performed with the “CentWave”, “PeakDensity”, “PeakGroups”, and “FillchromPeaks” algorithms. Annotation (isotopes, adducts) was conducted with the “CAMERA” algorithm (Kuhl et al., 2012). Intra-batch signal intensity drift was corrected by fitting a locally quadratic (loess) regression model to the QC values (Dunn et al., 2011; Van Der Kloet et al., 2009).

Three successive filtering steps using in-house scripts on R were applied, as in Georges des Aulnois et al. (2020). Pre-processing of +MS and -MS data matrices led to 8458 and 2651 variables respectively, and 540 and 203 variables remained after the filtrations. The two matrices were concatenated, log transformed and Pareto scaled before statistical analyses.

In parallel, HRMS data were acquired by autoMS/MS in an attempt to annotate the significant features revealed by the metabolomic approach, as in Georges des Aulnois et al. (2019).

II-2-6-5) Statistical analyses

Statistical analyses were performed on RStudio v 1.1.463. When the assumptions of independence (Durbin-Watson test), homoscedasticity (Bartlett test) and normality (Shapiro-Wilk test) of the residuals were validated, a two-way ANOVA was computed on the two factors location and salinity. When a significant difference was highlighted for one factor, a one-way ANOVA followed by a Tukey post hoc test was performed. When the previous assumptions were not met, data were log-transformed. Differences were considered statistically significant when P < 0.05, for a significance level of α = 0.05. Values were expressed as mean ± SD.

For metabolomics, the statistical analyses were carried out with MetaboAnalyst 4.0 (Chong et al., 2018). As the PLS-DA models were not validated (determined by a permutation test), ANOVA followed by a Tukey post hoc test (for the strains Dsa-Lo and Dsa-Me) and t-tests (for the strain

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Dsa-Th) were used to assess the features significantly affected between the three salinity conditions.

II-2-7) Results

II-2-7-1) Growth rate and biovolume

The three strains of “D. sacculus” (supplementary materials Table 25) survived to the 24 h hypoosmotic (25) and hyperosmotic (42) stress conditions and maintained a positive growth (Table 24). The growth rates ranged from 0.16 ± 0.04 and 0.33 ± 0.06 d-1 and the biovolumes from 12.4 ± 1.50 x 103 to 14.5 ± 1.65 x 103 µm3 (Table 24). However, neither growth rates nor biovolumes were significantly different between salinities and locations.

Table 24: Growth rate (µ, d-1) and biovolume (x103 µm3) of the 3 strains of “D. sacculus” after 24 h of salinity stresses at 25 and 42 and for the control. Values are expressed as mean ± SD (n = 3 for and n ≥ 30 for biovolume).

µ Biovolume Strain Salinity (d-1) (x103 µm3) 25 0.17 ± 0.06 13.2 ± 1.67 Dsa-Lo 35 0.23 ± 0.13 14.1 ± 1.70 42 0.33 ± 0.06 12.4 ± 1.50 25 0.25 ± 0.04 14.0 ± 1.75 Dsa-Me 35 0.16 ± 0.04 13.4 ± 1.77

42 0.20 ± 0.06 13.8 ± 1.75 25 0.28 ± 0.10 14.1 ± 1.77 Dsa-Th 35 0.18 ± 0.05 14.2 ± 2.29 42 0.20 ± 0.11 14.5 ± 1.65

II-2-7-2) Toxins

The three strains of “D. sacculus” exhibited the same qualitative toxin profile constituted by 80- 91% of PTX2eq and 9-20% of OA (Figure 25), with some quantitative differences. The strain Dsa-

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Lo synthesized significantly less OA (2.0 ± 0.4 pg cell-1; P < 0.001; mean of the three salinities) than Dsa-Me and Dsa-Th (8.4 ± 1.3 and 10 ± 0.8 pg cell-1, respectively; Figure 25) and the highest PTX2eq concentration was found in the strain Dsa-Me (83 ± 9.0 pg cell-1; P < 0.001; Figure 25).

However, no significant effect of hypoosmotic or hyperosmotic stresses was observed on toxin contents after this 24 h experiment. Furthermore, the proportion of intracellular toxins was high for both OA (> 85%) and PTX2eq (> 88%) but similar among strains and salinities (supplementary materials Table 26).

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II-2-7-3) Proline, GBT and DMSP

The three strains of “D. sacculus” synthesized proline, GBT and DMSP and the origin of the strains had no effect on the concentrations (i.e. no intraspecific variability) (Figure 3).

The concentration of proline, GBT and DMSP appeared to be greater at the highest salinity tested (42; Figure 26). The mean concentration of proline in control (35) for the three strains was 4.0 ± 0.6 fmol cell-1 and the salinity stresses led to 1.1-fold less and 1.6-fold more proline at salinity 25 and 42, respectively (Figure 26 A). More specifically, for the strains Dsa-Lo and Dsa-Me, a 2-fold significantly higher concentration of proline (P < 0.05) was observed between salinity 42 and 25 (6.2 ± 1.4 vs. 2.7 ± 0.8 fmol cell-1 and 4.7 ± 0.7 vs. 2.0 ± 1.2 fmol cell-1 respectively; Figure 26 A).

For GBT, the mean concentration in the controls was 3.2 ± 1.4 fmol cell-1 and was similar at the lowest salinity (25) (Figure 26 B). However, for the strain Dsa-Lo a significant 6.3-fold higher GBT concentration was observed at salinity 42 (10 ± 5.0 fmol cell-1) compared to control (1.7 ± 0.3 fmol cell-1) and around 2-fold for the strain Dsa-Th and Dsa-Me (Figure 26 B). For this last strain, a 4-fold higher accumulation of GBT was observed at 42 compared to 25 (Figure 26 B).

In control conditions (salinity 35), “D. sacculus” synthesized 1.0 ± 0.2 pmol cell-1 of DMSP (Figure 26 C) with a non-significant trend of accumulation at 42 compared to 35 (1.1-fold) and at 35 compared to 25 (1.2-fold). The only significant difference was for the strain Dsa-Lo, with a 1.4- fold higher DMSP content at salinity 42 compared to 25 (P < 0.05; Figure 26 C).

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o b

r M n a

7.5 i 7.5 B 0.75

y l

B G 5.0 ab 5.0 AB 0.50 A AB a A a 2.5 2.5 a 0.25

0.0 0.0 0.00

DSA-LoDsa-Lo DsaDSA-Me-Me DsaDSA-Th-Th DsaDSA-Lo-Lo DsaDSA-Me-Me DsaDSA-Th-Th DsaDSA-Lo-Lo DsaDSA-Me-Me DsaDSA-Th-Th Strain Strain Strain Strains of “D. sacculus” Strains of “D. sacculus” Strains of “D. sacculus”

Figure 26: Intracellular concentration of A. proline (fmol cell-1), B. glycine betaine (fmol cell-1) and C. DMSP (pmol cell-1) in the three strains of “D. sacculus” after 24 h of salinity stresses at 25 and 42 and for the control. Values are expressed as mean ± SD (n = 3). Conditions with different superscript letters were significantly different

II-2-7-4) Metabolomics

Metabolomics and the resulting PCA illustrated the intraspecific variability of the three strains of “D. sacculus” (Figure 27). We could not however discriminate the control from the hypoosmotic and hyperosmotic conditions, thus no clear salinity effect was observed despite the presence of six hundred features in the data matrix (Figure 27). Nevertheless, 19, 2 and 32 features were significantly affected by salinity in the strain Dsa-Me, Dsa-Lo and Dsa-Th respectively (supplementary materials Table 27 A-C).

Among them, adenosine and eicosapentaenoic acid (EPA) (features M268T51 and M301T600 in supplementary materials Table 27 C), were identified in Dsa-Th (supplementary materials Table 27, Figure 28 A-B) based on (Georges des Aulnois et al., 2020) and by comparison with a standard (same retention time, mass and similar MS/MS spectra), respectively. These two compounds were significantly less synthesized at the highest salinity (i.e. 42; Figure 28; P < 0.05).

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30 Dsa-Me S25 Dsa-Me S35 Dsa-Me S42 20 Dsa-Lo S25 Dsa-Lo S35 Dsa-Lo S42 10 Dsa-Th S25 Dsa-Th S35 Dsa-Th S42

0 PC(11.4%) 2 -10

-20

-30 -25 -20 -15 -10 -5 0 5 10 15 20 25 30 PC 1 (14.7%)

Figure 27: Principal component analysis (PCA) score plot obtained from LC-HRMS profiles of the three “D. sacculus” strains exposed to the three salinities (S)

II-2-8) Discussion

This study highlighted the good salinity tolerance of “D. sacculus” after 24 h salinity stresses. Indeed, growth rates and biovolumes were not significantly affected and there was no release of toxins (i.e. 85-99% were intracellular) that could be synonym of cell lysis and death (Smith et al., 2012). Moreover, the metabolomic approach could not discriminate the control from the hypoosmotic and hyperosmotic stressed cells, also showed by the toxin concentrations, which suggests that only minor metabolic modifications occurred. However, some osmoregulation has occurred since cells maintained a positive growth.

Similar to our observations, Errera and Campbell (2011) and Sunda et al. (2013) did not report any significant difference in the growth rates of several strains of Karenia brevis during a hypoosmotic stress (from 36 to 27). However and unlike our results, both studies noted a quick (i.e. in 3 minutes)

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Chapter 2-Part 2: Salinity stress on D. sacculus increase in biovolume, significant for Sunda et al. (2013) but not for Errera and Campbell (2011). It should be noted that dinoflagellates of the genus Karenia are naked cells, while Dinophysis is constituted of cellulosic thecal plates (Jensen and Daugbjerg, 2009). Thus, these solid plates and the rapidity of water flux exchanges (Kirst, 1989) may explain the absence of biovolume modification under hypoosmotic or hyperosmotic conditions after 24 h in this study.

One of our hypotheses was that DSTs acted like osmolytes. In toxic phytoplankton, de novo synthesis has been suggested to play a role in osmoregulation during salinity stress. Indeed Errera and Campbell (2011) suggested that toxins of K. brevis (brevetoxins) may “facilitate osmoregulation” because of their interaction with ion channels that can affect ion concentrations. Indeed, these authors pointed out an important (up to 14-fold compared to control) and rapid (< 3 h) accumulation of brevetoxins in several strains of K. brevis subjected to a hypoosmotic stress. Nonetheless, this was not confirmed by Sunda et al. (2013) with a similar experiment and strains, who refuted the interaction of brevetoxins and sodium channels. Accumulation of another phycotoxin, domoic acid, has been shown for the toxic diatom Pseudo-nitzschia australis after 24 and 48h hypoosmotic stress (35 to 30), which suggests that this toxin did not act like an osmolyte (Ayache et al., 2019).

Our study indicated that the two “D. sacculus” toxins, i.e. OA and PTX2, were not involved in osmoregulation since no significant change in their concentrations was observed after hypo- or hyperosmotic stress, as would be expected for osmolytes (Kirst, 1989).

A crucial step in the osmoregulation of photosynthetic microorganisms is the synthesis of osmolytes (Kirst, 1989). This work shows that three strains of “D. sacculus” were able to accumulate nitrogen and sulfur containing osmolytes, respectively proline, GBT and DMSP, at the highest salinity tested, when compared to the lowest, and even during hyperosmotic stress condition with GBT (strain Dsa-Lo) (Hagemann, 2011; Keller et al., 2004, 1999; Kirst, 1989).

To our knowledge, there is a lack of short-term (i.e. sampling times ≤ 24 h) studies on proline, GBT and DMSP contents in dinoflagellates subjected to rapid salinity stress conditions. However, time is a key parameter in osmoregulation (Gwinn et al., 2019; Hagemann, 2011). After acclimation of a strain of Prorocentrum minimum (salinities from 16 to 36), Gebser and Pohnert (2013) showed a constant level of DMSP, while a 20-fold increase of GBT was observed after hyperosmotic acclimation. Caruana et al. (2020) observed an increase of DMSP for only one strain of

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Alexandrium minutum (only on a per-cell basis) after hyperosmotic acclimation (38), whereas GBT content decreased after hypoosmotic acclimation (33) for one strain of A. pacificum. Overall, these studies showed the important role of GBT, and to a lesser extent of DMSP, in the osmoregulation process even after acclimation toxic phytoplankton to different salinity conditions of toxic phytoplankton.

This observation may be explained by the fact that DMSP is less active (in stabilization or catalysis of macromolecules) than GBT and proline (Kirst, 1996; Rhodes et al., 2002), the latter being one of the most efficient osmolytes (Kirst, 1989). Caruana and Malin (2014) classified DMSP as a “major osmolyte” for D. acuminata but according to our study, DMSP was likely to be a “medium to minor osmolyte” in “D. sacculus”, with a maximum increase of 16 mM between salinity 35 and 42 for Dsa-Lo. Data from Caruana and Malin (2014) were obtained from a field study on a natural population of D. acuminata by Jean et al. (2005), who reported 477 mM of DMSP. The difference could be attributed to the fact that in the environment, sulfur-derived compounds (e.g. DMSP) may represent a more important part of synthesized osmolytes than nitrogen-derived compounds (e.g. proline or GBT ), because nitrogen is usually more limiting than sulfur (Gwinn et al., 2019; Keller et al., 1999; Raven and Giordano, 2016; Rhodes et al., 2002).

Whether the adjustment of proline, GBT and DMSP concentrations by “D. sacculus” is the only mechanism of osmoregulation for this species/genus and whether this synthesis is effective enough to maintain intracellular homeostasis during salinity stress remains to be elucidated.

Despite the absence of discrimination between control and hypo- hyperosmotic stress conditions by the LC-HRMS analysis, several features were significantly affected by the different salinities (supplementary materials Table 27 and Figure 28), including EPA and adenosine. The polyunsaturated fatty acid (PUFA) EPA is mainly observed in marine eukaryotic organisms (especially diatoms and dinoflagellates) (Peltomaa et al., 2019) and is an essential fatty acid for heterotrophic consumers. It has for instance a beneficial effect on growth and fecundity of marine organisms, and it prevents or treats several human diseases (Bajpai and Bajpai, 1993; Galloway and Winder, 2015; Okuyama et al., 2008). EPA produced by toxic dinoflagellates and raphidophytes displayed toxic effects when associated with ROS and was suggested to be involved in prey capture mechanism of M. rubrum by Dinophysis spp. (Mafra et al., 2016 and references therein). A meta-analysis on phytoplankton fatty acid profiles according to environmental

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Chapter 2-Part 2: Salinity stress on D. sacculus conditions by Galloway et al. (Galloway and Winder, 2015), showed a negative correlation between dinoflagellates long-chain essential fatty acids (e.g. EPA) and salinity, thus corroborated our observation, and may be explained by modification of fatty acid metabolism by salinity variation (Kirst, 1990).

The other unidentified features may also play a role in the osmoregulation process. In the literature, some other potential osmolytes such as carbohydrates (Rhodes et al., 2002) or polyols (Borowitzka, 2018; Gebser and Pohnert, 2013) have been described but their mechanism of action are still unknown. Still, the absence of effects of salinity on metabolic profiles consistent between strains may also be attributed to the fact that: (1) cell densities of D. sacculus cultures were low, possibly giving rise to a lack of sensitivity with our LC-HRMS analysis and/or (2) only the methanol-soluble part of the metabolome was analyzed which may not be optimal for the extraction of highly polar molecules. Indeed, one of the main difficulties of this experiment was to obtain dense cultures before the experiment, thus also limiting number of extractions.

The dinoflagellate D. sacculus is known for its higher preference for semi-enclosed areas (Zingone et al., 1998) which are, as many coastal areas, frequently subjected to potential salinity variations (Fu et al., 2012; IPCC, 2013; Skliris et al., 2014). Two out of the three strains of “D. sacculus” used in this study were isolated from semi-enclosed environments, from the Atlantic (Dsa-Me, Arcachon Bay) and Mediterranean (Dsa-Th, Thau Lagoon) coasts respectively, while the third strain (Dsa-Lo, Loscolo) originated from an estuarine region. These three sites show some salinity fluctuations (Figure 24). Indeed, while the mean salinity appears around 35 and is quite stable in Meyran-Arcachon (between 30.1-35.7), it can decrease down to 25.3 at Loscolo, sometimes subjected to fresh water releases from a dam. On the opposite, the salinity at Thau Lagoon (Crique de l’Angle) can reach up to 41.3. Salinity in this Mediterranean lagoon is influenced by the balance between evaporation, rainfall and exchange with the sea among other factors (Collos et al., 2009). Importantly, Dinophysis spp., including “D. sacculus”, were observed in these locations at all salinities reported (Gaillard et al., 2020b; REPHY, 2019). Other Dinophysis species, such as D. acuminata, a species genetically close to “D. sacculus” (Séchet et al., 2020), have been shown to naturally grow in a wide range of salinities. Indeed, in Chile and Brazil, D. acuminata has been found at salinities between 5 to 31 and 20 to 34, respectively (Alves-de-Souza et al., 2019; Diaz et al., 2011) and at salinities 8 to 27 in North East USA (Hattenrath-Lehmann and Gobler, 2015),

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Chapter 2-Part 2: Salinity stress on D. sacculus reflecting an important tolerance to salinity fluctuations. Moreover, in a recent study, the salinity tolerance in vitro of two strains of D. acuminata-complex, D. ovum (DoSS3195) and D. acuminata (DAVA01) was highlighted by a positive growth rate over a range from 22 to 34, although the optimal growth were between 22 and 26.

Some toxic species of this genus, such as D. acuta (Farrell et al., 2012; Fux et al., 2010) and D. acuminata (Alves-de-Souza et al., 2014; González-Gil et al., 2010; Velo-Suárez et al., 2010; Xie et al., 2007) have been found to perform well in thin layers. These thin layers (often less than 0.5 m thickness) are nutrient-rich layers, where phytoplankton are concentrated, and blooms formed by e.g. temperature and/or salinity stratification (GEOHAB, 2008). Thus, this study, and other (Fiorendino et al., 2020), showed that D. sacculus and other toxic species of the genus Dinophysis are tolerant to variations of salinity, and could consequently display mechanisms of osmoregulation. Whether this trait is solely due to variations of the known measured osmolytes (i.e. proline, GBT and DMSP) remains to be shown, as these variations were also sometimes not significantly different from the control condition (except for GBT content in the strain Dsa-Lo), but simply showed a coherent trend from the lowest to the highest salinity.

In future studies, it would be interesting to measure the ion concentrations, such as sodium, potassium and ion fluxes, as well as the activity of ion channels, known to mediate a rapid response to salinity stress (Hagemann, 2011), with epifluorescence, NMR or radiotracer methods during osmotic stress (Hagemann, 2016; Louzao et al., 2006). It also appears fundamental to investigate kinetics of the synthesis of osmolytes with longer-term (> days) responses to salinity stress measurement and/or acclimation of several species of the genus Dinophysis including different chemotypes and non-toxic species. Finally, identification of all molecular features significantly affected by salinity in this study is challenging but necessary in order to better understand and characterize the osmoregulation of Dinophysis and other toxic phytoplankton.

II-2-9) Acknowledgements

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University of Copenhagen, Helsingør, Denmark) for M. rubrum cultures. We also thank Nicolas Chomérat for the corrections on the manuscript.

II-2-10) Supplementary materials

Table 25: Origin and culture conditions of the strains used in this study

Irradiancea Temperature Species Location Strain ID Code Medium (µmol photons (°C) m-2 s-1) Teleaulax Huelva AND- - L1-Si 17.8 ± 0.60 ~ 100 amphioxeia (Spain) A0710 Mesodinium Helsingør Harbor MBL- - L1/20-Si 17.8 ± 0.60 ~ 100 rubrum (Denmark) DK2009 Loscolo IFR-DSA- Sterilized Dsa-Lo 17.8 ± 0.60 ~ 100 (French Atlantic) 01Lo sea water Meyran-Arcachon IFR-DSA- Sterilized “Dinophysis Dsa-Me 17.8 ± 0.60 ~ 100 (French Atlantic) 01Me sea water sacculus” Thau Lagoon (Crique IFR-DSA- Sterilized de l’Angle) Dsa-Th 17.8 ± 0.60 ~ 100 02Th sea water (French Mediterranean) a Cultures were subjected to light in the PAR domain with a circadian cycle 12 h: 12 h (light: dark).

“Dinophysis sacculus” strains Salinity Intracellular OA (%) Intracellular PTX2eq (%)

25 94 ± 1.0 99 ± 0.3 Dsa-Lo 35 94 ± 0.9 99 ± 0.2 42 94 ± 1.3 99 ± 0.2

25 92 ± 0.8 97 ± 0.5 Dsa-Me 35 85 ± 15 88 ± 17 42 86 ± 4.6 95 ± 1.9 25 93 ± 1.4 98 ± 0.3 Dsa-Th 35 88 ± 4.2 97 ± 1.3 42 90 ± 2.5 97 ± 1.4

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A feature m/z rt (s) mode Salinity (significant) M155T299 155.071 299 NEG S35-42 Significantly < at 25 M169T320 169.086 320 NEG S35-42 M191T48 191.019 48 NEG S35-42 M214T464 214.253 464 POS S35-42 Dsa-Me

M258T457 258.279 457 POS S35-42 M284T651 284.257 651 POS S35-42

M288T424 288.290 424 POS S35-42 M288T434 288.291 434 POS S35-42

M300T496 300.325 496 POS S35-42

M316T503 316.321 503 POS S35-42 M318T484 318.327 484 POS S35-42

M332T447 332.316 447 POS S35-42 M411T475 411.299 475 POS S35-42

M424T529 424.318 529 POS S35-42

M628T451 628.452 451 POS S35-42 M795T757 795.139 757 S35-42

Significantly < at 25 and M316T466_2 316.320 466 POS S35 42 M288T444 288.290 444 POS S25-35 Significantly < at 42 Significantly > 25 (vs. M414T599 414.393 599 POS S25*-35 35 and 42) and 35 > 42

B feature m/z rt (s) mode Salinity (significant) M434T730 433.810 730 POS S25-35 Significantly < at 42 Dsa-Lo M315T94 315.154 94 POS S25 Significantly < at 25

C feature m/z rt (s) mode Salinity (significant) Dsa-Th M124T47 124.039 47 POS S25-35 Significantly < at 42 M166T48 166.087 48 POS S25-35

M167T40 166.833 40 NEG S25-35

M219T513_2 219.175 513 NEG S25-35 M268T51 268.104 51 POS S25-35 Adenosine

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M301T600 301.216 600 NEG S25-35 EPA

M327T622 327.232 622 NEG S25-35 M360T39 360.149 39 POS S25-35 M624T721 623.502 721 POS S25-35

M658T622 657.525 622 NEG S25-35 M787T484 787.463 484 POS S25-35

M791T733 790.548 733 POS S25-35

M803T700 803.449 700 NEG S25-35 M803T483 803.457 483 NEG S25-35

M805T483 805.462 483 NEG S25-35 M812T794 811.591 794 NEG S25-35

M816T796 815.529 796 NEG S25-35

M835T807 834.6106 607 POS S25-35 M903T513 903.474 513 NEG S25-35

M905T481 905.432 481 POS S25-35 M905T589 905.434 589 POS S25-35

M921T587 921.425 587 NEG S25-35

M921T479 921.425 479 NEG S25-35 M924T568 924.474 568 POS S25-35 M924T661 924.475 661 POS S25-35 M940T521 940.471 521 POS S25-35 M941T481 941.472 481 POS S25-35 M989T520 989.412 520 NEG S25-35 M1126T828 1125.675 828 NEG S25-35 M1194T828 1193.662 828 NEG S25-35 M553T765 553.197 765 POS S42 Significantly > at 42 M387T320 387.213 320 POS S42

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120 Chapter 2-Part 2: Salinity stress on D. sacculus

100 A B

1 80

l l

e Salinity c

g 25 p (area) 35

2 60

X 42

P EPA EPA (area)

Adenosine 40 * *

20 Dsa-Th Dsa-Th

0 Figure 28: A. Adenosine (feature M268T51) and B. eicosapentaenoic acid (EPA) (feature DSA-Lo DSA-Me DSA-Th M301T600) areas at the three salinities for the “D. sacculus” strainStrain from Thau lagoon (Dsa-Th).

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Chapter 3: Effects on marine organisms Part 1: Dinophysis and its toxins towards oyster gametes

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III-1) Cultures of Dinophysis sacculus, D. acuminata, and pectenotoxin 2 affect gametes and fertilization success of the Pacific oyster, Crassostrea gigas

This Chapter 3-Part 1, has been published in Environmental Pollution in May 2020:

III-1-1) Graphical abstract

Exposition to Oocytes and/or Spermatozoa

Mortality and ROS production

4 + 3 2

0 -C 0.5 c.ml-1 5 c.ml-1 50 c.ml-1 500+ c.ml-1 Fertilization rate

100 Dinophysis sacculus Dinophysis acuminata Fertilization 75

0 -C 0.5 c.ml-1 5 c.ml-1 50 c.ml-1 500+ c.ml-1 Pectenotoxin 2 Okadaic acid Dinophysis or toxin concentrations

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III-1-2) Résumé

Les efflorescences de microalgues toxiques et nuisibles (ou « harmful algal blooms », i.e. HAB en anglais) sont une menace pour la santé humaine. C’est le cas par exemples d’espèces de dinoflagellés toxiques du genre Dinophysis, qui sont principalement responsables de l'intoxication diarrhéique par les mollusques (DSP) chez les consommateurs de coquillages contaminés. Une telle contamination entraîne des fermetures de la pêche et collecte des animaux dans les zones contaminées, et donc de fermes conchylicoles, causant ainsi des problèmes socio-économiques majeurs. Les effets directs de nombreuses espèces d’HAB ont été démontrés sur les bivalves adultes, alors que les effets sur les premiers stades de vie des bivalves, étapes les plus critiques, restent relativement inexplorés. Cette étude visait à évaluer les effets in vitro des souches cultivées de D. sacculus et D. acuminata isolées en France ou de leurs toxines associées (à savoir l'acide okadaïque (OA) et la pecténotoxine 2 (PTX2)) sur la qualité des gamètes de l’espèce de coquillages la plus commercialisée en France, l'huître creuse du Pacifique, Crassostrea gigas. Des mesures de la production de ROS et la viabilité des gamètes par cytométrie en flux, ainsi que l’évaluation du succès de la fécondation à l'aide de comptages microscopiques ont été réalisées. Les ovocytes ont été plus affectés que les spermatozoïdes et leur mortalité ainsi que leur production de ROS a augmenté en présence de D. sacculus et de PTX2, respectivement. Une diminution du succès de la fécondation a aussi été observée à de faibles concentrations : 0,5 cellules par ml-1 de Dinophysis spp. et 5 nM de PTX2, alors qu'aucun effet de l'OA n'a pu être observé. L'effet sur le succès de la fécondation s’est accentué lorsque les deux types de gamètes ont été préalablement exposés et de manière concomitante par rapport à des expositions séparées, ce qui suggère un effet synergique. Nos résultats suggèrent également que les effets pourraient être dus au contact direct entre cellules algales et gamètes. Ces résultats mettent en évidence un effet potentiel de Dinophysis spp. et de la PTX2 sur la reproduction et le recrutement de l'huître du Pacifique.

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III-1-3) Abstract

Harmful algal blooms (HABs) of toxic species of the dinoflagellate genus Dinophysis are a threat to human health as they are mainly responsible for diarrheic shellfish poisoning (DSP) in the consumers of contaminated shellfish. Such contamination leads to shellfish farm closures causing major economic and social issues. The direct effects of numerous HAB species have been demonstrated on adult bivalves, whereas the effects on critical early life stages remain relatively unexplored. The present study aimed to determine the in vitro effects of either cultivated strains of D. sacculus and D. acuminata isolated from France or their associated toxins (i.e. okadaic acid (OA) and pectenotoxin 2 (PTX2)) on the quality of the gametes of the Pacific oyster Crassostrea gigas. This was performed by assessing the ROS production and viability of the gametes using flow cytometry, and fertilization success using microscopic counts. Oocytes were more affected than spermatozoa and their mortality and ROS production increased in the presence of D. sacculus and PTX2, respectively. A decrease in fertilization success was observed at concentrations as low as 0.5 cell mL-1 of Dinophysis spp. and 5 nM of PTX2, whereas no effect of OA could be observed. The effect on fertilization success was higher when both gamete types were concomitantly exposed compared to separate exposures, suggesting a synergistic effect. Our results also suggest that the effects could be due to cell-to-cell contact. These results highlight a potential effect of Dinophysis spp. and PTX2 on reproduction and recruitment of the Pacific oyster.

III-1-4) Key words

Dinophysis spp.; okadaic acid; pectenotoxins; oyster gametes; fertilization success

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III-1-5) Introduction

Harmful algal blooms (HABs) of toxic microalgae are increasing in terms of frequency, intensity and duration due, in part, to climate change and eutrophication (Gobler et al., 2017; Hallegraeff, 1993; Wells et al., 2019). Toxins associated with HABs can accumulate in marine bivalves (Landsberg, 2002; Shumway, 1990; Simões et al., 2015), causing a threat to human health through direct contact with toxins or consumption of contaminated organisms (Hallegraeff, 2010, 1993; Van Dolah, 2000). Consequently, national surveillance programs monitoring phytoplankton and phycotoxin concentrations in water and bivalves have been implemented (e.g. REPHY in France). The European Council has set a maximum limit of 160 µg OA eq. per kg of fresh whole bivalve meat (EU Commission, 2011), above which shellfish harvesting (farming and recreational) is forbidden in order to protect human consumers (Nielsen et al., 2012).

Shellfish farming is an important economic sector worldwide. In France, the Pacific oyster, Crassostrea gigas (= Magallana gigas) (Thunberg, 1793) represents the majority of annual shellfish sales (ca. 118,000 tons; France Agrimer, 2018). Shellfish farmers in France annually suffer economic losses due to the presence of several toxic species of the genus Dinophysis (Ehrenberg, 1841; Marcaillou et al., 2005; Trainer et al., 2020). Indeed, D. acuminata and D. sacculus are the main responsible for shellfish farm closures, that can last for several weeks per year (Belin and Soudant, 2018; Marchand et al., 2009). Along French coasts, Dinophysis spp. are regularly observed at a concentration of 102 cells L-1 (supplementary materials Figure 34, REPHY, 2019), which is similar to concentrations typically reported in the literature (Reguera et al., 2012). However, blooms of Dinophysis spp. can occasionally reach cell densities up to 103 - 107 cells L-1 (reviewed in Reguera et al. (2012b)), including one instance of 8 x 105 cells L-1 reported in France (REPHY, 2019).

These dinoflagellates can produce two types of lipophilic toxins, okadaic acid (OA) and its analogs dinophysistoxins (DTXs), and pectenotoxins (PTXs; Marcaillou et al. 2005, Reguera et al. 2014). Okadaic acid and DTXs are responsible for diarrheic shellfish poisoning (DSP) in humans following shellfish consumption (Lawrence et al., 2000; Reguera and Pizarro, 2008), with symptoms that include diarrhea, nausea, vomiting and abdominal pain (Yasumoto et al., 1978). In contrast, PTXs are not considered diarrheic shellfish toxin (DST) as they do not cause diarrhea in

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Chapter 3-Part 1: Dinophysis and its toxins towards oyster gametes humans (Matsushima et al., 2015). However, PTXs are lethal to mice by intraperitoneal injection (Miles et al., 2004).

While HABs have mostly been studied in relation to public health, another fundamental issue is the direct effect they have on filter-feeding bivalves (Landsberg, 2002; Matsuyama et al., 2001; Shumway and Cucci, 1987; Shumway, 1990). National monitoring programs along the French Atlantic coast indicate that spawning, development and recruitment of larvae may co-occur with Dinophysis spp. (supplementary materials Figure 34) (Pouvreau et al., 2016; REPHY, 2019). While adult and juvenile bivalves can mechanically escape toxic microalgae by cessation of filtration and closing their shells (Hégaret et al., 2007), the planktonic early life stages such as gametes and embryos are directly exposed to HABs and their toxins in the water column and appear more sensitive than adults (Castrec et al., 2019; Glibert et al., 2007; Stoecker et al., 2008; Wang et al., 2006; Yan et al., 2001).

Many studies have focused on the effects of toxic dinoflagellate species on oyster gametes, embryos and larvae, e.g. for the genera Alexandrium (Banno et al., 2018; Basti et al., 2015a; Castrec et al., 2020, 2019; Matsuyama et al., 2001; Mu and Li, 2013), Karenia (Leverone et al. 2006, Rolton et al. 2014, 2015, 2016, Basti et al. 2015a), Heterocapsa (Basti et al., 2013, 2011), Gymnodinium (Matsuyama et al., 2001), Karlodinium and Prorocentrum (Glibert et al., 2007; Stoecker et al., 2008). Nevertheless, due to the mixotrophy of toxic species of the genus Dinophysis and the resulting difficulty in their cultivation until recently (Park et al., 2006), few studies have investigated the effects of Dinophysis spp., their toxins or combinations of both on bivalves, such as oysters. The few available studies indicate that Dinophysis spp. producing PTXs induce hypersecretion of mucus and pseudofeces, paralysis, alteration of the tissues within the digestive gland and reduced escape response in adult scallops (Basti et al., 2015b). Mccarthy et al., (2014) demonstrated that exposure of OA to adult Pacific oysters and blue mussels increased DNA fragmentation. Further studies also highlighted modified hemocyte functions in both Mediterranean mussels (Malagoli et al., 2008; Prado-Alvarez et al., 2012) and carpet shell clams (Prado-Alvarez et al., 2013) exposed to Dinophysis spp. and their toxins.

The present study investigated the in vitro effects of whole culture, resuspended cells and culture filtrate of Dinophysis sacculus (Stein, 1883), whole culture of D. acuminata (Claparède and Lachmann, 1859) and certified standards of OA and PTX2 on (i) gamete cellular characteristics

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(i.e., ROS production, mortality, and morphology), and (ii) fertilization success of oocytes and spermatozoa of the Pacific oyster.

III-1-6) Materials and methods

III-1-6-1) Microalgal cultures

Monoclonal cultures of D. sacculus (Stein, 1883) (strain IFR-DSA-01Lt) and D. acuminata (Claparède and Lachmann, 1859) (strain IFR-DAU-02Ar) were isolated in Arcachon, France, in 2015 and 2018, respectively. These mixotrophic species were cultivated in 0.2 µm filter-sterilized natural seawater (FSSW) for D. sacculus and L1/20-Si + K/2-Si (Hernández-Urcera et al., 2018) for D. acuminata at salinity 35 and fed every two days with ciliate prey Mesodinium rubrum (Lohmann, 1908) (strain MBL-DK2009) at a ratio of 1: 1 (predator: prey) according to Park et al. (2006). The ciliate M. rubrum was fed three times a week with the cryptophyte Teleaulax amphioxeia (Conrad) (Hill, 1992) (strain AND-0710). Both M. rubrum and T. amphioxeia were cultivated in flasks respectively in L1/20-Si and L1-Si (Guillard and Hargraves, 1993) and diluted every two days for the ciliates and every week for the cryptophyte. All cultures were maintained at 17.8 ± 0.6 °C, at a light intensity of ~ 100 µmol photons m-2 s-1 provided by cool-white and pink fluorescent tubes (fluora and cool-white fluorescent light, Osram, Munich, Germany) and a 12: 12 (L: D) cycle (supplementary materials Table 30). To increase the biomass of Dinophysis spp., cultures were fed at a ratio of 1: 10 (predator: prey) for 4 months before the experiment. One week before the experiment, cultures of Dinophysis spp. were filtered on a nylon sieve (mesh 11 µm) and gently rinsed with 75 mL of FSSW to remove any cryptophyte and ciliate. Cultures were then resuspended in 20 mL of FSSW and starved for one week before the experiment to obtain cells from the mid exponential growth phase on the day of the experiment.

III-1-6-2) Experimental design

All experiments are summarized in supplementary materials Figure 33 and performed with 3 to 11 replicates. A replicate is either a pool of several females or males, or one male or female, as detailed below.

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III-1-6-2-1) Experiment 1 (Exp. 1) - Effect of whole culture of D. sacculus upon gametes

The aim of Exp. 1 was to determine the effect of whole cultures of D. sacculus on gamete cellular characteristics and fertilization success. In total, four fertilization experiments were performed by crossing exposed or non-exposed oocytes and/or spermatozoa to D. sacculus.

- (i) Oocytes exposed to D. sacculus (n = 8 pools of gametes, each from 3 different females) were crossed with non-exposed spermatozoa (a pool of spermatozoa from 5 males)

-or (ii) Spermatozoa exposed to D. sacculus (n = 8 pools of gametes, each from 3 different males) were crossed with non-exposed oocytes (a pool of oocytes from 5 females).

- (iii) In addition, fertilization success was determined after exposure of both oocytes and spermatozoa to D. sacculus (n = 4 pools of gametes, each from 3 different organisms).

- (iv) The effects of whole cultures of D. sacculus on fertilization success was also investigated by exposing gametes only during fertilization. Oocytes, spermatozoa and D. sacculus (at final concentrations of 0 (control), 0.5, 5, 50 and 500 cells mL-1) were put in contact simultaneously in FSSW (n = 11 pools of gametes, each from 3 different organisms).

Briefly, for (i), (ii) and (iii); oocytes or spermatozoa were exposed for 2 h in glass vials at 20 ± 1 °C to the whole culture of D. sacculus at a final concentration of 0 (control), 0.5, 5, 50 and 500 cells mL-1 in FSSW. These D. sacculus cell concentrations were selected to mimic the cell densities in natural blooms occurring in France, from typical (1 x 102 cells L-1) to exceptionally dense blooms (8 x 105 cells L-1, supplementary materials Figure 34) (REPHY, 2019).

III-1-6-2-2) Experiment 2 (Exp. 2) - Effect of resuspended cells and culture filtrate of D. sacculus upon gametes

Experiment 2 was designed to determine the respective effect of resuspended cells and extracellular medium (culture filtrate) of D. sacculus on gametes. (i) Oocytes or (ii) spermatozoa (n = 5 individual females or males) were exposed for 2 h in glass vials either to the whole D. sacculus culture at 500 cells mL-1 (similar to Exp. 1), or to D. sacculus cells only, obtained by filtration (11 µm-mesh nylon sieve) of a culture at 500 cells mL-1 and resuspended in FSSW (to remove the extracellular metabolites) or to culture filtrate obtained by filtration (0.2 µm-mesh nylon sieve) of the whole culture (500 cells mL-1) to measure the effect of only the extracellular metabolites of

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III-1-6-2-3) Experiment 3 (Exp. 3) - Effect the whole culture of D. acuminata upon gametes

The aim of Exp. 3 was to determine the effect of a 2 h exposure of whole cultures of D. acuminata (at a final concentration of 0 (control), 0.5, 5, 50 and 500 cells mL-1) on gamete cellular characteristics and fertilization success of both exposed oocytes and spermatozoa (n = 4 pools of gametes, each from 3 different organisms), as described in Exp. 1 (iii).

III-1-6-2-4) Experiment 4 (Exp. 4) - Effect of OA PTX2 standards on gametes

Experiment 4 investigated the effect of 0 (control), 5, 10, 20 and 50 nM solutions of okadaic acid (OA) and pectenotoxin 2 (PTX2) in FSSW and to the corresponding methanol (MeOH) control (2.3 % final) on cellular characteristics and fertilization success of (i) oocytes or (ii) spermatozoa, (iii) both oocytes and spermatozoa exposed to toxins and (iv) gametes exposed to toxins only during fertilization (n = 3 to 4 pools of gametes from 3 different organisms). These concentrations of toxins approximately corresponded to the minimum concentration of OA (i.e. 5 nM) found in the studied strains of D. sacculus and D. acuminata (sum of intra and extracellular toxins of 500 cells) to ca. the maximum concentration of PTX2 (i.e. 50 nM; Table 28). All measurements were performed as detailed in Exp. 1.

III-1-6-3) Toxin analyses of Dinophysis spp. strains

Toxin analysis was adapted from Sibat et al. (2018) and García-Portela et al. (2018a) and performed on 1 mL (n = 3) sub samples of Dinophysis spp. cultures collected in exponential growth phase. After centrifugation (3500 g, 4 °C, 15 min), both cells (pellets) and culture filtrates (supernatants) were extracted. The pellet (intracellular toxins) was extracted with 0.5 mL methanol and sonicated at 25 kHz for 15 min. Extracellular toxins were recovered from the supernatant after liquid-liquid extraction with dichloromethane, which was evaporated under nitrogen and resuspended in 0.5 mL of methanol. Subsequently, samples were filtered (0.2 µm, Nanosep, MF, Pall, Northborough, MA,

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USA). Analyses were performed using ultra high performance liquid chromatography coupled to low resolution tandem mass spectrometry (UHPLC-LRMS/MS) with a UHPLC system (UFLC XR Nexera, Shimadzu, Tokyo, Japan) coupled to a triple quadrupole/ion-trap mass spectrometer (API 4000 QTrap, ABSciex, Redwood City, CA, USA), equipped with a turboV® ESI source (see details in García-Portela et al. 2018). Certified calibration solutions of PTX2, OA, dinophysistoxin 1 and 2 (DTX1 and DTX2) were obtained from the National Research Council Canada (NRCC, Halifax, NS, Canada). Intracellular (intra) and total (sum of intracellular and extracellular) toxin contents were expressed on a per cell basis (pg cell-1) while extracellular (extra) as equivalent (eq) pg cell- 1. Pectenotoxin 2 eq (PTX2eq) was the sum of pectenotoxin 2, pectenotoxin 2b, pectenotoxin 2 seco-acid and 7-epi-pectenotoxin 2 seco-acid, all quantified with PTX2 standard by assuming similar molar responses.

III-1-6-4) Sampling and maintenance of oysters

Sexually mature C. gigas were collected at La Pointe du Chateau-Baie de Daoulas, France (48°20’01.8”N 4°19’02.6”W) in summer 2018 and 2019 or obtained from IFREMER experimental facilities as described in Castrec et al. (2019). Individuals from the field were cleaned with filtered sea water (FSW) to remove sessile organisms. All oysters were maintained in an aerated tank at 16 ± 1 °C with a continuous flow of FSW for one to three days before the experiments.

III-1-6-5) Collection of gametes

Gonads were dissected and placed in individual Petri dishes to collect gametes according to Song et al. (2009) for oocytes and Boulais et al. (2015) for spermatozoa. Briefly, for each oyster, gametes were collected in 10 mL FSSW and sieved through 100 µm mesh to isolate gametes from gonad debris. Only motile spermatozoa and rounded oocytes were selected for the experiments (Rolton et al., 2015). Spermatozoa and oocyte concentrations were determined by flow cytometry (FCM) according to Le Goïc et al. (2013, 2014) and diluted in FSSW at 107 cells and 105 mL-1, respectively.

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III-1-6-6) Flow-cytometry analysis - morphology, viability and ROS

production

Analyses of morphology, viability, and reactive oxygen species (ROS) production (i.e. cellular characteristics) of oyster gametes by FCM were adapted from Le Goïc et al. (2013, 2014) and carried out with an EasyCyte Plus cytometer (Guava Technologies, Millipore, Luminex Billerica, USA) equipped with a 488-nm argon laser and three fluorescence detectors: green (525 ± 30 nm), yellow (583 ± 26 nm) and red (680 ± 30 nm).

For cell morphology measurements, values of the forward scatter (FSC) and side scatter (SSC), respectively proxies of cell size and complexity, were used to estimate cell morphology of spermatozoa and oocytes. Spermatozoa viability was assessed using double staining spermatozoa solution with 2 µL of propidium iodide (PI) and 2 µL of SYBR-14 (Live/Dead® Sperm Viability Kit, Molecular Probes, Eugene, OR, USA) at final concentrations of 2 µg mL-1 and 1 µM during 10 min in the dark (Le Goïc et al., 2013). For oocyte viability, PI and SYBR-Green-1 (1/10,000 of the commercial solution; Molecular Probes, Eugene, OR, USA) were used (Le Goïc et al., 2014).

ROS production of gametes was measured by staining 200 µL of oocytes or spermatozoa solution with 2 L (final concentration of 10 M) of dye 2’7,7’-dichlorofluorescein diacetate DCFH-DA (Sigma, St Quentin Fallavier, France) for 1 h in the dark (Le Goïc et al., 2014; Vignier et al., 2017).

ROS production was expressed as percentage of control (a.u.). For both viability (expressed as mortality) and ROS production measurements, oocytes and spermatozoa concentrations were 5 x 104 and 5 x 106 cells mL-1, respectively.

III-1-6-7) Microscopy analysis - fertilization success assessment

To assess fertilization success, oocytes and spermatozoa were inoculated in FSSW (20 ± 1 °C) at a ratio of 1: 100 (5 x 103 oocytes: 5 x 105 spermatozoa) in 12-well plates in a final volume of 4 mL FSSW (Boulais et al., 2017). When fertilized oocytes in the control (i.e. non-exposed oocytes and spermatozoa) reached 80 % or, after 2 h of incubation, samples were fixed with 1 % formaldehyde (final concentration). Fertilization success (%) was assessed under an inverted light microscope (Axio observer.Z1, Zeiss, Oberkochen, Germany) by counting fertilized and unfertilized oocytes

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(polar body extrusion from 2 to 8-cell stages vs. no polar body). For the fertilization success measurements in all experiments except conditions (iv), the concentration of Dinophysis spp. or toxins were minimum 30-fold lower due to dilution in FSSW and their potential contribution to the observed effect were considered not significant.

III-1-6-8) Statistial analysis

Statistical analyses were performed on RStudio v 1.1.463. After checking the assumptions of independence (Durbin-Watson test), homoscedasticity (Bartlett test) and normality (Shapiro-Wilk test) of the residuals, t-test or one-way ANOVA followed by a Tukey post hoc test were computed. Otherwise, Mann-Withney U or Kruskal-Wallis tests were used, followed by a Conover test. Differences were considered statistically significant when P < 0.05, for a significance level of α = 0.05. Values were expressed as mean ± SD.

III-1-7) Results

III-1-7-1) Toxin contents of Dinophysis spp. cultures

The D. sacculus strain synthesized OA and three PTX2 derivatives (pectenotoxin 2, pectenotoxin 2b, pectenotoxin 2 seco-acid, 7-epi-pectenotoxin 2 seco-acid) whereas the D. acuminata strain synthesized only OA. Neither of the two algal species produced DTX1 or DTX2 (Table 28).

For Exp. 1 and Exp. 2, D. sacculus produced similar amounts of total OA (4.5 ± 1.4 and 6.0 ± 0.46 pg cell-1) and total PTX2eq, i.e. sum of concentrations (95 ± 36 and 76 ± 19 pg cell-1; Table 28). The majority of OA was in the extracellular compartment in contrast to PTX2 which was mainly intracellular. For Exp. 3 with D. acuminata, the total OA content per cell was 16-fold higher (87 ± 11 pg cell-1) than D. sacculus and >90 % was intracellular (P < 0.001; Table 28).

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OA PTX2eq Total Total Extra Intra Extra Total (nM in Intra Total (nM in Exp. (eq pg (pg cell- (eq pg (pg cell- 500 (pg cell-1) (pg cell-1) 500 cells cell-1) 1) cell-1) 1) cells mL-1) mL-1) 1 2.2 ± 0.52a 2.3 ± 1.1a 4.5 ± 1.4a 2.8 ± 0.9a 73 ± 26 22 ± 9.0 95 ± 34 55 ± 20 D. 3.9 ± 6.0 ± sacculus 2 2.1 ± 0.17a 3.7 ± 0.3a 64 ± 16 12 ± 2.9 76 ± 19 44 ± 11 0.29a 0.46a D. 3 80 ± 9.0b 7.5 ± 1.9b 87 ± 11b 54 ± 6.8b < LD < LD < LD < LD acuminata

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III-1-7-2) Exp. 1 - Effect of the whole culture of D. sacculus on gametes

None of the tested concentrations affected spermatozoa cellular characteristics. However, mortality of oocytes was 2.9-fold-higher when exposed to D. sacculus at a concentration of 500 cells mL-1 compared to the control (P < 0.05), whereas no effect was observed on ROS production. Moreover, a significant increase in FSC was observed for the same exposure condition compared to control (P < 0.05; Table 29).

Fertilization success decreased significantly when (i) oocytes or (ii) spermatozoa were exposed to 50 and 500 cells mL-1 of D. sacculus compared to their respective controls (89 vs. 56 and 2 % and 83 vs. 58 and 33 %, respectively; P < 0.001; Figure 29 A-B). Interestingly, a 17-fold difference was noted between spermatozoa and oocytes when exposed to 500 cells mL-1 (P < 0.001; Figure 29 A-B) with oocytes being more sensitive. When (iii) both gametes were exposed to D. sacculus, fertilization success compared to control was significantly reduced by 25, 44 and 93 % at 5, 50 and 500 cells mL-1, respectively (P < 0.001; Figure 29 C). The fertilization success in presence of D. sacculus during the fertilization (iv) was significantly reduced by 18, 39 and 57 % when exposed to 0.5, 5 and 50 cells mL-1, respectively (P < 0.001). Fertilization was however, totally impeded at 500 cells mL-1 (Figure 29 D).

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Oocytes Spermatozoa Mortality ROS Mortality FSC SSC ROS (%) FSC SSC (%) (%) (%) 79.7 57 ± Control 359 ± 27a 1752 ± 65 100 1.7 ± 1.5a 100 2.8 ± 1.3 ± 2.1 1.7 Exp. 79.9 55 ± 123 ± 0.5 360 ± 28a 1737 ± 70 91 ± 23 2.1 ± 1.6ab 4.8 ± 1.9 1 ± 1.8 3.1 21 D. 79.8 55 ± 134 ± 5 360 ± 28a 1738 ± 66 96 ± 37 2.6 ± 1.9ab 3.5 ± 1.2 sacculus ± 1.6 2.9 21 (cells 80.1 56 ± 131 ± 50 371 ± 26a 1746 ± 66 92 ± 8.8 2.3 ± 1.5ab 2.8 ± 1.1 mL-1) ± 1.7 2.0 25 79.3 56 ± 118 ± 500 473 ± 30b 1733 ± 66 106 ± 43 4.9 ± 1.6b 3.5 ± 2.0 ± 1.5 1.6 25 97.6 74 ± Control 300 ± 11a 1780 ± 38 100 3.8 ± 0.8a 100 16 ± 4.4 ± 1.6 1.7 Exp. 3 97.8 75 ± 92 ± 0.5 303 ± 7.9a 1781 ± 42 110 ± 42 3.7 ± 0.6a 14 ± 2.2 D. ± 2.5 2.9 27 acuminat 98.7 75 ± 91 ± 5 304 ± 17ab 1783 ± 36 113 ± 90 3.7 ± 0.4a 17 ± 5.8 a ± 1.6 1.7 14 (cells 96.9 76 ± 108 ± 50 323 ± 7.6ab 1761 ± 40 136 ± 45 n.a 19 ± 4.5 mL-1) ± 2.0 1.1 32 96.9 75 ± 69 ± 500 348 ± 38b 1756 ± 69 113 ± 58 10.4 ± 2.5b 18 ± 6.5 ± 0.9 1.0 12 98.1 77 ± Control 326 ± 11 1676 ± 76 100a 1.4 ± 1.0 100 4.9 ± 2.4 ± 1.5 2.4 120 ± 97.9 77 ± 163 ± MeOH 329 ± 10 1677 ± 76 2.3 ± 0.9 6.2 ± 2.3 32ab ± 1.7 1.2 124 97.5 78 ± 206 ± OA 5 329 ± 8.8 1688 ± 71 100 ± 46a 1.6 ± 0.9 4.3 ± 2.3 ± 1.5 1.4 56 97.7 78 ± 193 ± OA 10 330 ± 7.8 1681 ± 69 75 ± 25a 1.5 ± 0.4 4.5 ± 2.8 ± 1.8 2.1 51 97.9 78 ± 168 ± Exp. 4 OA 20 328 ± 9.4 1680 ± 69 91 ± 30a 1.4 ± 0.5 5.2 ± 3.8 ± 2.1 1.3 82 Toxins 97.7 77 ± 155 ± (nM) OA 50 328 ± 8.4 1672 ± 58 74 ± 35a 2.2 ± 0.5 5.7 ± 5.6 ± 1.7 1.7 61 97,6 78 ± 199 ± PTX2 5 329 ± 9.6 1679 ± 67 94 ± 36a 2.4 ± 1.1 4.0 ± 2.7 ± 1.5 1.1 68 PTX2 97,6 77 ± 179 ± 332 ± 10 1674 ± 71 90 ± 28a 2.3 ± 1.0 4.4 ± 2.1 10 ± 1.5 1.8 52 PTX2 97.6 77 ± 173 ± 333 ± 9.7 1678 ± 68 105 ± 52a 2.8 ± 0.9 5.1 ± 3.3 20 ± 1.8 2.1 95 PTX2 203 ± 97.2 76 ± 193 ± 353 ± 14 1678 ± 65 3.1 ± 0.8 4.8 ± 3.5 50 108b ± 2.8 2.1 77

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a 100 A 100 a a B a a a b 75 b 75 c

with with exposed 50 50 with exposed

25 25

Fertilization Fertilization

oocytes oocytes (%) Fertilization Fertilization 0 spermatozoa (%) 0 Control 0C 0.5 0.5c.ml-1 5 c.ml-1 5 50 c.ml-150 500 500 c.ml-1 ControlC 0.5 0.5c.ml-1 5 c.ml-1 5 50 c.ml-150 500 500 c.ml-1 (control) C 100 a D 100 ab bc 75 75 c a b

50 50

in in presence of with with both (%) c

gametes gametes (%) d 25 25

sacculus

. ertilization

ertilization e

D F exposed 0 0 ControlC 0.5 0.5c.ml-1 5 c.ml-1 5 50 c.ml-150 500 500 c.ml-1 ControlC 0.5 0.5c.ml-1 5 c.ml-1 5 50 c.ml-150 500 500 c.ml-1

D. sacculus concentration (cells mL -1) D. sacculus concentration (cells mL -1)

III-1-7-3) Exp. 2 - Effect of resuspended cells and culture filtrate of D. sacculus

on gametes

The negative effect of gametes exposed to whole cultures (500 cells mL-1) was confirmed (P < 0.001), with fertilization success 10 times lower following exposure of oocytes vs. spermatozoa (P < 0.001; Figure 30 A-B). While similar results were obtained using resuspended D. sacculus cells (P < 0.05), no significant difference was noted between the controls and gametes exposed to culture filtrates (Figure 30 A-B).

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a A 100 a B 100 a ab

75 75

b b with with exposed with with exposed 50 50

25 25

b b

Fertilization Fertilization

spermatozoa (%) Fertilization Fertilization oocytes oocytes (%) 0 0 ControlC 500 500 c.ml-1 Resuspended SupernatantFiltrate (eq ControlC 500 500 c.ml-1 Resuspended SupernatantFiltrate (eq cells (500)(500 c.ml- 500 (eq c.ml-1)500) cells (500)(500 c.ml- 500 (eq c.ml-1)500) 1) 1) Conditions of exposure to D. sacculus (cells mL -1) Conditions of exposure to D. sacculus (cells mL -1)

III-1-7-4) Exp. 3 - Effect the whole culture of D. acuminata on gametes

Only oocytes exposed to 500 cells mL-1 of D. acuminata were significantly affected, with a 2.7- fold higher mortality (P < 0.001) and a 16 % increase in FSC (P < 0.001; Table 29). Again, spermatozoa were not affected. A significant decrease in fertilization success after exposure of both gametes was observed from as few as 5 cells mL-1 (1.8-fold; P < 0.05), while fertilization was almost completely inhibited at 500 cells mL-1 of D. acuminata (P < 0.001; Figure 31).

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100 a a 75 b

50 with with both

c gametes gametes (%) 25

ertilization d exposed F 0 ControlC 0.5 0.5c.ml-1 5 c.ml-1 5 50 c.ml-150 500 500 c.ml-1

D. acuminata concentration (cells mL-1)

III-1-7-5) Exp. 4 - Effect of OA and PTX2 standards on gametes

Production of ROS was around twice higher in oocytes exposed for 2 h to 50 nM of PTX2 (P < 0.05) compared to the control while spermatozoa were not affected by exposure to OA and PTX2 standards (Table 29).

A decrease in fertilization success was observed with (i) oocytes exposed for 2 h to 20 and 50 nM of PTX2 compared to control (16 and 0 % vs. 77 %, respectively; P < 0.001; Figure 32 A). Similarly, (ii) spermatozoa exposed to 20 and 50 nM of PTX2 decreased fertilization success compared to control (35 and 25 % vs. 81 %; P < 0.001; Figure 32 B). However, oocytes exposed to 50 nM, exhibited a more pronounced effect than exposed spermatozoa on fertilization (P < 0.05; Figure 32 A-B). Exposure of (iii) both oocytes and spermatozoa to 5, 10, 20 and 50 nM of PTX2 reduced the fertilization success by 25, 37, 78 and 97 % compared to control (P < 0.01; Figure 32 C). The fertilization in presence of toxins (iv) was significantly diminished only at a concentration of 50 nM of PTX2 (32 %) compared to control (76 %; P < 0.001; Figure 32 D). Neither OA (at any of the concentrations tested) nor MeOH in the control affected the fertilization success.

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A 100 B 100 a a a a a a a a a a a a a

75 a 75 a a exposed

50 50 b b b 25 25

Fertilization with Fertilization with

exposed oocytes (%) Fertilization with Fertilization with 0 spermatozoa (%) 0 ControlMeOH MeOH OA OA OA OA PTX2PTX PTXPTX2 PTXPTX2 PTXPTX2 Control MeOHMeOH OA OA OA OA PTX2PTX PTXPTX2 PTXPTX2 PTXPTX2 5 nm5 10 10 nM 20 20 nM 5050 5 5nm 10 10 nM 20 5050 5 nm5 10 10 nM 20 20 nM 5050 5 5nm 10 10 nM 20 5050 nM nM nM nM nM nM

C 100 a a ab ab D 100 a a a a a a abc abc a bc c 75 75 a ab 50 50

b toxins toxins (%) d 25 25

Fertilization with Fertilization with both

exposed gametes (%) Fertilization Fertilization in 0 presence of 0 ControlMeOH MeOH OA OA OA OA PTX2PTX PTXPTX2 PTXPTX2 PTXPTX2 Control MeOHMeOH OA OA OAOA OA PTX2PTX PTX2PTX PTXPTX2 PTXPTX2 5 nm5 10 10 nM 20 20 nM 5050 5 5nm 10 10 nM 20 5050 5 5nm 10 10 nM 20 20 nM 50 50 5 5nm 10 10 nM 2020 50 nM nM nM nM nM nM Toxins concentration (nM) Toxins concentration (nM)

III-1-8) Discussion

The present work demonstrated that Dinophysis sacculus and D. acuminata as well as one of their toxins PTX2 impaired cellular characteristics and fertilization success in gametes of the Pacific oyster, Crassostrea gigas, a species of commercial interest.

The concentrations of Dinophysis spp. used in this exposure study were selected because of their environmental relevance, since concentrations of > 1000 cells L-1 were frequently observed in the four regions studied on the French Atlantic coast, whereas concentrations of > 10,000 cells L-1 were occasionally observed in two of the regions, including major oyster production sites, i.e. the Bay of Arcachon and the Bay of Brest (supplementary materials Figure 34). These concentrations,

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Chapter 3-Part 1: Dinophysis and its toxins towards oyster gametes observed in France over a ten-year period, are moderate compared to other areas affected by Dinophysis spp. blooms, e.g. India or Norway, where concentrations of up 1.5 x 106 and 2.3 x 107 cells L-1 have been observed (Reguera et al., 2012).

The toxin exposure concentrations (i.e. from 5 to 50 nM) corresponded to the maximum amounts of OA and PTX2, respectively, produced by 500 cells of D. sacculus in our experiment. Noteworthy, PTX2 caused an increase in ROS production of oocytes exposed to 50 nM, which could reflect a stimulation of oocyte metabolism or cellular stress. Indeed, the production of ROS is a key mechanism involved in stress responses (Kadomura et al., 2006), but an excess of ROS could lead to cellular toxic effects such as destruction of membrane integrity by lipid peroxidation, DNA damage and associated alteration of cell functioning and ultimately cell death (Cavallo et al., 2003; Landsberg, 2002; Lesser, 2006). This may explain the observed reduced fertilization success of oocytes exposed to PTX2. Similarly, Le Goïc et al. (2014) observed an increased ROS production of oocytes in the presence of the toxic dinoflagellate Alexandrium minutum, and suggested this increase production may have reduced oocyte quality.

Pectenotoxin 2 at concentrations as low as 20 nM reduced fertilization success when either oocytes or spermatozoa were pre-exposed. This is true also at concentration as low as 5 nM of PTX2 when oocytes and spermatozoa were both pre-exposed. This suggests that both oocytes and spermatozoa were negatively affected by PTX2 and that the effect of PTX2 on oyster gametes is cumulative. Secondly, this toxicity is likely mediated by a different mechanism than ROS production since no increase in ROS production was observed below 50 nM concentration of PTX2.

Using mammalian and finfish cell lines (e.g. human, rat, rabbit, salmon), PTXs have been shown to interfere with actin assembly/disassembly, thereby affecting cell cytoskeletal functions and leading to cell death, at concentrations ranging from nM to µM (Ares et al., 2005; Dominguez et al., 2010; Spector et al., 1999). It has been shown that PTX2 causes actin depolymerization (Dominguez et al., 2010), sequestration of monomeric actin (at a concentration of 20 nM) (Spector et al., 1999), disrupted F-actin (Hori et al., 1999), and inhibited actin polymerization by a capping process at the barbed-end of F- and G-actin (Allingham et al., 2007). The reduced fertilization success observed in this study could be associated with impairment of the oocyte and spermatozoan cytoskeleton by PTX2, as well as fertilization itself since actin polymerization is a crucial mechanism in oysters, involved in spermatozoan motility and the penetration of the oocyte (Ledu

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Chapter 3-Part 1: Dinophysis and its toxins towards oyster gametes and McCombie, 2003). In the literature, the indirect evidence of the involvement of Dinophysis spp. in mortalities observed in the natural environment (reviewed in Landsberg 2002, Basti et al., 2015b) were almost exclusively related to D. caudata, a producer of high cellular contents of PTX2 (Basti et al., 2015b, 2015c; Fernández et al., 2006; Marasigan et al., 2001). The action of PTX2 on actin in oyster gametes would be worthy of investigation in further studies.

Okadaic acid, in contrast to PTX2, affected neither the gametes nor fertilization success, at concentrations up to 50 nM. Okadaic acid is reported to be an inhibitor of serine/threonine protein phosphatase 1 and 2 activities (Bialojan and Takai, 1988; Mccarthy et al., 2014) and is also believed to be a tumor promoter in humans (Lago et al., 2005). This toxin induced chromosome loss, apoptosis and DNA damages in mammalian cell lines (see references in Prado-Alvarez et al., 2013). Okadaic acid has been shown to induce an increase in DNA fragmentation in adult Pacific oyster and blue mussel (Mccarthy et al., 2014) and modified hemocyte functions in several bivalve species (Malagoli et al., 2008; Prado-Alvarez et al., 2013, 2012) at concentrations between 1.2 to 50 and 10 to 500 nM, respectively. The absence of effects of OA on oyster gametes in this study could be due to the relatively short exposure time and low OA concentrations, reaching respectively 2 h and a maximum of 50 nM concentration of certified standard and 4.5 nM concentration of extracellular OA produced by 500 cells of D. acuminata in our experiment.

The main structural difference between OA and PTX2 is that PTX2 is a macrocyclic lactone (i.e. cyclic ester). Pectenotoxin 2 biological activity on cytoskeletal dynamics is clearly associated with the macrocyclic ester as the activity disappeared when the esters were hydrolyzed and the macrocycle was opened (Allingham et al., 2007; Ares et al., 2007; Miles et al., 2006b). The resulting analogue, pectenotoxin 2 seco-acid, is structurally very similar to OA, and is not active on the cytoskeleton (neither is OA). Interestingly, PTX2 has a high structural similarity with goniodomin A, another algal macrocyclic lactone, which has also been reported to affect the cytoskeleton via F-actin (Espiña et al., 2016). In addition, it should be noted that among the three species of Alexandrium that produce goniodomins (Harris et al., 2020), A. monilatum has been clearly associated with fish kills since the middle of the last century (Howell, 1953) and more recently also with shellfish mortalities (Harding et al., 2009; May et al., 2010).

Our study also revealed that PTX2 is likely not to be the only bioactive compound responsible for the toxicity of Dinophysis spp. on oyster gametes and fertilization success. Firstly, fertilization

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Additionally, the present study indicated that the decreased fertilization success, specifically for D. sacculus, was derived from cells and not from the extracellular compartment, as filtrate had no activity, unlike resuspended cells. This observation could be explained by cell-to-cell contact and the effect of (a) mechanical damages and/or (b) surface-bound toxins and/or (c) quick release of intracellular bioactive compounds (Landsberg, 2002).

Contact with Dinophysis spp. cells, or by the mean of feeding peduncle (Ojamäe et al., 2016), may have resulted in (a) mechanical damage to the membranes of oyster gametes, as suggested by the increase in FSC morphological parameter of oocytes. This proxy of cell size could indicate a swelling of the cells, when exposed to 500 cells mL-1 of D. sacculus associated to an increase in mortality.

Furthermore, it has been hypothesized that (b) the presence of toxins on the cell surface of another HAB species, H. circularisquama can affect pearl oyster larvae after contact (Basti et al., 2011). In addition, Mu and Li (2013) suggested that the release by A. catenella of surface-located toxins may affect Pacific oyster egg hatching success. Surface-bound toxins from Dinophysis spp., Alexandrium spp. and Heterocapsa spp. have not been reported yet but could potentially explain the observed effect on gametes and fertilization success.

Another explanation could be (c) a rapid release of intracellular compounds, different from the already known and characterized toxins, with activity towards oyster gametes. While most attention has been paid to toxins affecting humans (i.e. DST in a sanitary context), other bioactive compounds from Dinophysis spp. have been overlooked despite some interesting observations. Basti et al. (2015b) observed mortality in adult mollusks fed D. caudata independently of PTX2 content, thus they hypothesized the presence of unknown toxins and/or other bioactive compounds. Similarly, Mafra et al. (2016) hypothesized that the mechanism of prey capture of several toxic species of Dinophysis spp. involved uncharacterized allelochemical compounds, other than the known DSTs, which debilitates M. rubrum. The existence of such allelochemicals for Karenia

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Chapter 3-Part 1: Dinophysis and its toxins towards oyster gametes brevis affecting C. virginica larvae (Rolton et al., 2014) as well as for A. minutum affecting early life stages and adults of C. gigas has also been observed (Castrec et al., 2020, 2018). Bioguided- fractionation approaches combining suitable bioassays (Long et al., 2018a) as well as chromatography coupled to e.g. high resolution mass spectrometry (Nothias et al., 2018) may be useful to identify these molecules. However, the difficulties inherent to the highly challenging culture of Dinophysis spp. requiring prey organisms may be a limitation in these kinds of studies.

Oyster spermatozoa are motile and small (2 µm) cells, which may thus have limited contact with Dinophysis spp. cells or their toxins, as opposed to the immotile and large (75 µm) oocytes, which are more likely to make physical contact with the dinoflagellate or its toxins. Moreover, spermatozoa and oocytes are different in term of composition (e.g. biochemical content), metabolism (e.g. mobility and embryonic development, respectively) and plasma membrane (Boulais et al., 2017, 2015). These observations may explain the different sensitivity between gametes exposed to Dinophysis spp. and PTX2 and the absence of effects observed by flow cytometry for spermatozoa.

In addition, the decrease in fertilization success when spermatozoa were exposed to D. sacculus and D. acuminata and the synergistic effect observed when both gametes were exposed, suggested that spermatozoa were indeed impacted by Dinophysis spp. Further measurements on spermatozoa should be focused on motility and velocity, as well as energetic metabolism and mitochondrial membrane potential, which can affect flagellar movements and ultimately fertilization capacity (Boulais et al., 2017, 2015; Le Goïc et al., 2013).

The inhibition of fertilization success was higher with gametes exposed to D. acuminata than to D. sacculus for the same concentration, however whether this species is more toxic or has a different mechanism of action is still unknown.

Some preliminary results also indicate that when gametes were exposed to D. sacculus, abnormal development of D-shaped larvae could be observed (personal communications), leading to the question of the effects on early life stages of C. gigas and ultimately, recruitment. Similarly, when C. gigas larvae were exposed to A. minutum, anomalies in swimming behavior, feeding and growth were observed which led to a decrease in survival and settlement of older larvae stages (Castrec et al., 2020).

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Analysis of the data collected in the REPHY and VELYGER monitoring programs clearly demonstrated that the concentrations of Dinophysis spp. used in this study are environmentally relevant and can occur during spawning of C. gigas (supplementary materials Figure 34). If shifts in climate lead to increased co-occurrence of Dinophysis spp. and oyster spawning periods, effects on reproduction could potentially increase. Economic impact assessment of Dinophysis spp. blooms in French coasts is underway as part of the CoCliME project. Additionally, any significant increase of Dinophysis spp. concentrations, e.g. through increased eutrophication is likely to also amplify such effects. Indeed, Dinophysis spp. blooms have been related to nutrient pollution in France (Souchu et al., 2013) and globally (Hattenrath-Lehmann and Gobler, 2015; Hattenrath- Lehmann et al., 2015).

III-1-9) Conclusion

This study highlighted for the first time that low cellular concentrations (i.e. 5 x 102 to 5 x 103 cells L-1) of toxic species of the genus Dinophysis, i.e. D. sacculus and D. acuminata, and low PTX2 concentration (5 nM) can interfere with fertilization success of C. gigas and can potentially affect reproduction of this species.

The adverse effects observed on oyster fertilization success and gamete cellular characteristics were similar for both Dinophysis species. Whether this activity is a general trait of the genus Dinophysis and results from similar mechanisms requires further investigation. Future studies should also include other Dinophysis spp., as this is a very diverse genus with species showing different toxin profiles, including some that do not produce toxins. It is important to explore the intraspecific and interspecific diversity of this activity and its broader impacts on shellfish reproduction.

Therefore, studies focusing on the effects of Dinophysis spp., especially PTX-producers, and their associated allelopathic or bioactive compounds appear fundamental to better assess their effects on marine organisms (i.e. bivalve, fish, zooplankton and phytoplankton).

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III-1-10) Acknowledgements

This work was funded by the project CoCliME which is part of ERA4CS, an ERA-NET initiated by JPI Climate, and funded by EPA (IE), ANR (FR), BMBF (DE), UEFISCDI (RO), RCN (NO) and FORMAS (SE), with co-funding by the European Union (Grant 690462). We thank David Jaén (LCCRRPP, Huelva, Spain) for T. amphioxeia and Per Juel Hansen for M. rubrum cultures. We acknowledge Justine Castrec for her valuable help during the experiment, as well as Nadine Neaud- Masson and Marc Sourisseau for extraction of data from Quadrige2 (the database of the REPHY network).

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III-1-11) Supplementary materials

Table 30: Culture conditions of strains used in this study

Temperature Irradiancea (µmol Species Origin Strain ID Medium (°C) photons m-2 s-1)

Teleaulax Huelva (Spain) AND-A0710 L1-Si 17.8 ± 0.6 ~ 100 amphioxeia

Mesodinium Helsingør Harbor MBL- L1/20-Si 17.8 ± 0.6 ~ 100 rubrum (Denmark) DK2009

Dinophysis Arcachon IFR-DSA- Sterilized 17.8 ± 0.6 ~ 100 sacculus (France) 01Lt sea water

Dinophysis Arcachon IFR-DAU- L1/20-Si + 17.8 ± 0.6 ~ 100 acuminata (France) 02Ar K/2-Si

a Cultures were subjected to light in the PAR domain during a circadian cycle 12 h: 12 h (light: dark)

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Gamete FCM Fertilization Experiments expositions Cellular caracteristics success

(i) + X X +

(ii) + X X + Exp. 1 X (iii) + X Cross together

(iv) + +

resuspended or X (i) + filtrate + Exp. 2 resuspended or X (ii) + filtrate +

Exp. 3 + X X Cross together

(i) + OA or PTX2 X X +

(ii) + OA or PTX2 X X + Exp. 4 X (iii) + OA or PTX2 X Cross together

(iv) OA or PTX2 + +

Figure 33: Experimental design Chapter 3-Part 1. Oysters gametes were exposed, or not, to a gradient of concentration from 0.5 to 500 cells mL-1 of (Exp. 1) Dinophysis sacculus and (Exp. 3) D. acuminata, to (Exp. 2) D. sacculus (500 cells mL-1), resuspended cells (500 cells mL-1) or to culture filtrate (eq 500 cells mL-1) and to (Exp. 4) a gradient of concentration from 5 to 50 nM of okadaic acid (OA) and pectenotoxin 2 (PTX2). Cellular characteristics (morphology, mortality and ROS production) were then analyzed, or not, by flow cytometry (FCM). Finally, depending on each case, exposed gametes were fertilized with non-exposed gametes, with exposed gametes or fertilization were made in presence of D. sacculus or toxins.

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1.E +06 1,E +05 Chapter 3-Part 1: Dinophysis and its toxins towards oyster gametes

1.E +05 1.E +06 D-shaped larvae 1,E +05 Dinophysis spp. 1.E+06 1.E+05 A 1.E +04 1,E +04 1.E +05

1.E+05 Dinophysis Dinophysis

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1.E+07 1.E+05 C

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A   B N C D

Figure 34: Concentration of Dinophysis spp. (black dots; cells L-1; log10) and D-shaped larvae (grey dots; larvae 1.5 m-3; log10) of Crassostrea gigas from A. Bay of Brest, B. Bay of Bourgneuf, C. Marennes-Oléron, and D. Bay of Arcachon, respectively, from the Northern the Bay of Biscay to the South. Data are occurrences from 2008 to 2017 extracted from the REPHY and VELYGER databases. Notes that 100 cells L-1 of Dinophysis spp. is the limit of detection (placed on the x- axis) and occurrences < LOD are not included in the data set. VELYGER observatories are not available for the year 2008 at B. Bay of Bourgneuf

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III-2) Mortality and histopathological features of sheepshead minnow (Cyprinodon variegatus) larvae exposed to Dinophysis acuminata and pectenotoxin 2 during a 96 h bioassay

This Chapter 3-Part 2, is in preparation for submission in a journal

Gaillard, S., Mason P. L., Ayache, N., Sanderson, M., Smith, J. L., Giddings, S., McCarron, P.,

Séchet, V., Hégaret, H., Réveillon, D., Hess, P., Vogelbein, W. K.

III-2-1) Graphical abstract

100 PTX2  mortality

LC50 1231 nM 0 0 24 48 72 96 Dinophysis acuminata Suffocation or 96 h bioassay Hypertrophy gill Pectenotoxin 2 + lamaellae and Cyprinodon chloride cells variegatus Ripped tissues 3 weeks old larvae

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III-2-2) Résumé

Les écosystèmes marins côtiers et estuariens sont soumis à des efflorescences de microalgues toxiques et nuisibles (ou harmful algal blooms, i.e. HAB en anglais). Dans ces zones, les HAB cooccurrent avec les périodes de reproduction et de pontes de mollusques et de poissons d'importance écologique et économique. Les espèces toxiques du genre de dinoflagellé Dinophysis synthétisent des toxines diarrhéiques lipophiles, l’acide okadaïque (OA) et ses dérivés les dinophysistoxines (DTX), qui sont des polyéthers linéaires et qui peuvent s'accumuler dans les bivalves filtreurs et provoquer une intoxication diarrhéique par les mollusques (DSP pour « diarrheic shellfish poisoning » en anglais) chez les consommateurs humains. De plus, certains Dinophysis spp. produisent des polyéthers macrocycliques, les pecténotoxines (PTX). L'effet de Dinophysis et de ses toxines sur les animaux marins, et en particulier sur les premiers stades de vie, est encore mal étudié. Dans cette étude, des larves de fondule tête de mouton (Cyprinodon variegatus), qui est un poisson estuarien commun, ont été exposées à un gradient de concentrations de D. acuminata (une souche isolée de la baie de Chesapeake, productrice d'OA, de DTX1 et de PTX2), de cellules remises en suspension, de filtrat de culture et de lysat ou un standard de PTX2 pendant un essai biologique larvaire de 96 h. Aucune mortalité n'a été observée lorsque les larves étaient exposées à D. acuminata (de 5 à 5500 cellules mL-1) tandis que l’exposition à la PTX2, à des concentrations intermédiaires à fortes (de 250 à 4000 nM), induisaient 8 à 100% de mortalités après 96 h. Cependant, dans les deux expériences d'exposition, les caractéristiques histopathologiques évaluées par microscopie optique et par microscopie électronique à transmission (TEM pour « transmission electron microscopy » en anglais) ont révélé des altérations de l'épithélium des branchies, une hypertrophie, une redistribution et une modification de la morphologie des cellules à chlorure (= ionocytes) dans les lamelles secondaires. Les dommages histopathologiques pourraient être dus à l'interaction entre la PTX2 et les filaments d'actine des cellules piliers ou avec les cellules à chlorure, ainsi qu'au contact de cellule à cellule entre D. acuminata et les filaments branchiaux. Dans l'ensemble, les effets observés suggèrent une suffocation des larves de poissons. D'autres observations seront effectuées à des concentrations plus faibles ainsi que dans les organes digestifs afin de déterminer un potentiel effet toxique à des concentrations de D. acuminata et de PTX2 écologiquement pertinentes.

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III-2-3) Abstract

Marine coastal and estuarine ecosystems frequently experience harmful algal blooms (HABs) of toxic phytoplankton. These events often co-occur with reproduction and spawning periods of ecologically and economically important mollusks and finfish. Toxic species of the dinoflagellate genus Dinophysis synthesize lipophilic diarrheic toxins, that include the linear polyether, okadaic acid (OA) and its derivatives, the dinophysistoxins (DTXs). These toxins can accumulate in filter- feeding bivalves and cause diarrheic shellfish poisoning (DSP) in their human consumers. Additionally, some Dinophysis spp. also produce macrocyclic polyethers, the pectenotoxins (PTXs). The effects of Dinophysis and its toxins on marine animals, specifically the critical early life stages, are still poorly studied. Here, 3-week-old sheepshead minnows (Cyprinodon variegatus), a common finfish in eastern USA estuaries, were exposed to a gradient of concentrations of either live D. acuminata culture, culture filtrate, cell lysate, cells resuspended in clean media (Chesapeake Bay strain DAVA01, producing primarily PTX2 with minor production of OA, and DTX1) or a PTX2 standard using a 96 h toxicity bioassay. No mortality was observed when larvae were exposed to D. acuminata (from 5 to 5500 cells mL-1), whereas exposure to PTX2 at intermediate and high concentrations (from 250 to 4000 nM) induced 8 to 100% mortality by 96 h. However, in both exposure experiments, histopathology and transmission electron microscopy (TEM) revealed significant gill damage, including intercellular edema (fluid accumulation), necrosis and sloughing of the gill respiratory epithelia, and damage to the osmoregulatory epithelium, including hypertrophy, proliferation, redistribution and necrosis of chloride cells (= ionocytes). Significant tissue damage in gills is hypothesized to be caused by the interaction of PTX2 with the actin cytoskeleton of the affected gill epithelia. The minor pathology observed in challenge of larvae to live Dinophysis culture and fractions thereof, may be attributable to cell to cell contact between D. acuminata and fish gill. Overall, the significant gill pathology observed in the Ptx2 challenge suggested that death is due to loss of respiratory and osmoregulatory functions in the fish larvae. Further observations will be conducted at lower concentrations and in digestive tracts, in order to determine the potential harmful effect of natural field concentrations of D. acuminata and PTX2.

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III-2-4) Key words

Chesapeake Bay; Dinoflagellate; Gills; Harmful algal bloom; Microscopy

III-2-5) Introduction

Dinoflagellates of the genus Dinophysis synthesize toxic polyethers, including the pectenotoxins (PTXs), okadaic acid (OA) and its derivatives the dinophysistoxins (DTXs). These latter two groups of compounds are also referred to as diarrheic shellfish toxins (DSTs) (Ares et al., 2007; Dominguez et al., 2010), that can bioaccumulate in filter-feeding bivalves during Dinophysis blooms, and cause diarrheic shellfish poisoning (DSP) in human consumers. The symptoms of intoxication, named DSP after an outbreak in Japan in late 1970s (Murata et al., 1982; Yasumoto et al., 1980, 1978), include vomiting, nausea, abdominal pain and diarrhea (Van Dolah, 2000; Yasumoto et al., 1985).

Okadaic acid and several DTXs, including the analogue DTX1 and the acyl derivate DTX3 (Dominguez et al., 2010; Suzuki and Mitsuya, 2001), have been shown to cause in vivo cytotoxicity in the digestive organs (i.e. liver and intestine) of rodents (Berven et al., 2001; Ito and Terao, 1994; Terao et al., 1993; Tubaro et al., 2003) and death following oral and intraperitoneal injections, from 160 µg kg-1 for OA to 500 µg kg-1 for DTX3 (Dominguez et al., 2010). Both OA and DTXs are inhibitors of serine/threonine protein phosphatases (PP), especially PP1 and PP2A (Cruz et al., 2007; Fernandez et al., 2002) and can interfere with ion secretion cells in intestine, leading to the above-mentioned diarrheic symptoms (Cohen et al., 1990).

Currently, PTXs are no longer considered to be diarrheic toxins (Miles et al., 2004a), and are therefore not monitored by some sanitary programs (e.g. in the US; NSSP, 2017; Trainer et al., 2013). In some instances they were recently recommended for exclusion from monitoring programs (e.g. in New Zealand; Boundy et al., 2020). Most of the PTXs found in bivalves are metabolic products of PTX2, e.g. seco acid products (i.e. ring-opened), PTX1, PTX3 or PTX6 (Boundy et al., 2020; Miles et al., 2006b; Suzuki et al., 2005) or acyl esters of seco acids (Wilkins et al., 2006). While death and hepatotoxicity have been shown following intraperitoneal injections of nM concentration of PTX2 in rodents (Terao et al., 1986), studies on oral ingestion of PTX2 and seco acid products in mice did not induce death (> 5000 µg kg-1) (Ito et al., 2008; Miles et al., 2004a).

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However, exposure of mammalian cell lines to PTXs at nM to µM concentrations, has shown these compounds to interfere with actin by depolymerization, sequestration and disruption, and finally affecting the cytoskeletal functions (Allingham et al., 2007; Ares et al., 2005; Hori et al., 1999; Leira et al., 2002; Spector et al., 1999; Zhou et al., 1994).

Toxic species of the genus Dinophysis are cosmopolitan (i.e. D. acuminata) (Nagai et al., 2020) and they are a recurrent problem due to contamination of bivalves where shellfish farming activities are located (i.e. in Western Europe or in Japan) (Reguera et al., 2014). In the United States, Dinophysis blooms have recurred since the 1960s. However, they have been considered a real threat only since 2008 when a bloom of D. ovum in the Gulf of Mexico led to the first harvesting ban due to the presence of OA equivalent toxins exceeding the sanitary threshold in the oyster, Crassostrea virginica (Campbell et al., 2010; Deeds et al., 2010). Thereafter, OA and DTX1 were found in shellfish following blooms of D. acuminata in Long Island Sound (NY) (Hattenrath-Lehmann et al., 2013) and in Puget Sound (WA). These events led to shellfish harvesting closures (Lloyd et al., 2013; Trainer et al., 2013; Wolny et al., 2020).

In general, phycotoxins can directly affect marine animals through exposure to phytoplankton or dissolved toxins, following ingestion or uptake across the gills (e.g. filter-feeding bivalves, planktivorous fish). Indirect exposure has been reported to occur through the consumption of contaminated organisms (i.e. carnivorous fish, marine mammals and birds) (Landsberg, 2002). Filter-feeding bivalves, such as mussels or oysters, accumulate significant quantities of toxins due to their mode of nutrition but can metabolize them into less toxic forms (Blanco et al., 2018; Mafra et al., 2019). Similarly, filter-feeding fish such as clupeiforms are also vulnerable to acute and/or chronic toxicity due to ingestion of toxic phytoplankton (Landsberg, 2002). In this context, early life stages of fish are more sensitive to toxic phytoplankton and/or associated phycotoxins due to their immobility or low mobility compared to adult fish which could potentially escape contaminated waters (Wiegand et al., 1999).

Finfish mortalities were observed during a bloom of D. caudata (a known PTX2-producer; Basti et al., 2015; Luisa Fernández et al., 2006) in the Gulf of Thailand and in India (Okaichi, 1967; Santhanam and Srinivasan, 1996), but direct causality was not demonstrated. Unfortunately, since these events, only a few studies focused on the effects of Dinophysis spp. and their toxins on fish, which is probably related to the complexities of Dinophysis cultivation and experimental

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Some experiments have looked at the exposure of fish embryos to nM concentration of OA and observed pathology in digestive organs, genotoxicity and mortality (Escoffier et al., 2007; Le Du et al., 2017). Exposure of finfish larvae to µM concentrations of OA caused gill and liver pathology and developmental disorders (Figueroa et al., 2020; Souid et al., 2018).

However, to date only one study has focused on the effects of Dinophysis cells on fish. Rountos et al. (2019) exposed 1 to 7 days old larvae larvae of estuarine sheepshead minnow, Cyprinodon variegatus, and Inland silverside, Menidia beryllina, for 6 days to toxic strains of D. acuminata and Alexandrium catenella (1000 cells mL-1). Results indicated that a mix of the two toxic species increased the mortality in both fish, while D. acuminata alone decreased C. variegatus growth rate (Rountos et al., 2019), suggesting a harmful effect of associated toxins.

The eurythermic and benthic sheepshead minnow (C. variegatus) is important from an ecological and economical point of view in Central and North East America (Bennett and Beitinger, 1997; Bushong et al., 1988; Gervasi et al., 2019; Hall et al., 1994; Rosenfield et al., 2004). Moreover, spawning seasons of C. variegatus (i.e. spring-summer) (Rountos et al., 2019) are likely to overlap with bloom periods of several toxic species of the genus Dinophysis. Indeed, in the lower Chesapeake Bay, and other estuaries located in Atlantic coast of the United States, multiple toxic species of the genus Dinophysis, including D. acuminata, have been reported in monitoring programs since the 1960s (see references in Wolny et al., 2020), to occur from spring to fall, and more specifically in June (Marshall, 1982, 1980).

To date, few studies have specifically focused on potential adverse health effects of PTX2 in estuarine organisms. We recently observed deleterious effects of both PTX2 and the PTX2- producer D. sacculus on oyster oocytes, with an increase in reactive oxygen species (ROS) after 2h of exposure to 50 nM of PTX2 and higher mortality after 2 h of exposure to ecologically relevant concentrations (from 0.5 to 500 cells mL-1) of D. sacculus, while no negative effects of OA were observed (Gaillard et al., 2020b). These cellular effects led to a decrease in the fertilization success and could ultimately impact recruitment of oysters, both wild and cultured, highlighting the importance of additional studies on PTX-producers (Gaillard et al., 2020b).

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The chain forming and bioluminescent species Alexandrium monilatum produces goniodomin A, a compound which has structural similarity to PTX2 and a toxicological mechanism of action related to that of PTX2, as both compounds target actin (Harris et al., 2020; Murakami et al., 1988; Tainter et al., 2020). Blooms of this species have been observed in several American coastal areas, including the Chesapeake Bay (Harding et al., 2009), and have caused deleterious effects in finfish and shellfish (May et al., 2010 and references therein). The structural similarities of PTX2 and goniodomin A suggest a potentially detrimental effect of PTX2 on early life stages of oysters and thus their recruitment, prompting us to study and characterize the effect of PTX2 and a PTX2- producer on estuarine fish.

In the present study we applied a previously described 96 h larval fish bioassay using larval sheepshead minnows (Lovko et al., 2003; Vogelbein et al., 2002) to investigate the effects of (i) a cultivated isolate of D. acuminata (whole culture, cells resuspended in clean media, culture filtrate and lysate), and (ii) purified PTX2. We measured cumulative mortality, and investigated tissue pathology associated with exposures using routine paraffin histopathology and transmission electron microscopy.

III-2-6) Materials and methods

III-2-6-1) Microalgal cultures

The dinoflagellate Dinophysis acuminata (Claparède and Lachmann, 1859) (strain DAVA01) was isolated from the Chesapeake Bay, Nassawadox, Virginia, USA in May 2017. This mixotrophic species was cultivated in f/2 medium prepared with 0.22 µm filter-sterilized natural seawater (FSW) obtained from Wachapreague, Virginia, with a salinity of 35, and diluted with distilled water to reach a value of 25 (Table 33). The dinoflagellate was fed every two days with its ciliate prey Mesodinium rubrum (Lohmann, 1908) (strain JAMR) at a predator: prey ratio of 1: 5. The Mesodinium culture was fed twice a week with the cryptophyte Teleaulax amphioxeia (Conrad) Hill (Hill, 1992) (strain JATA) at a predator: prey ratio of 1: 10. Both T. amphioxeia and M. rubrum were isolated from Inokushi Bay, Oita prefecture, Japan in 2007 and cultivated in f/2 and f/12 media, respectively. All the species were grown in batch culture and maintained in an incubator at

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15 ± 1 °C, at a light intensity of ≈ 100 mol photons m-2 s-1 on a 12: 12 h (light: dark) cycle (Table 33).

One week before the experiment, a stock culture of D. acuminata was filtered on a 10-µm nylon mesh sieve to remove cryptophytes and ciliates, gently rinsed with FSW and resuspended in fresh medium. The culture was then starved to ensure that D. acuminata cells remained in mid exponential growth phase for one-week prior to the experiment.

III-2-6-2) Fish larvae

Three-week old sheepshead minnow larvae, Cyprinodon variegatus (Lacépède 1803), were purchased and delivered the day of each experiment from Aquatic Biosystems (Fort Collins, Colorado, USA). They were fasted for 48 hrs prior to over-night delivery, there were no mortalities due to shipping and all animals appeared to be in good health.

III-2-6-3) Experimental design

III-2-6-3-1) 96 h larval fish bioassay

The larval finfish bioassay was adapted from Vogelbein et al. (2002) and Lovko et al. (2003). Small (1 mL) glass beakers were filled with 0.5 mL (volume validated with pre-test experiment) of D. acuminata or PTX2 solutions (see details below). One fish larva was transferred per beaker using a disposable plastic pipette. Exposures were performed for 96 hours in covered 48-well plates at room temperature (22 to 25 °C). Every 24 h, larvae were monitored for cumulative mortality assessment under a binocular dissecting microscope or were sampled and processed for routine paraffin histological and transmission electron microscopical (TEM) analyses (see details below).

The calculation of an LC50 was based on the Hill coefficients (Hill, 1910) from dose-response curves (Prinz, 2010) using REGTOX macro on ExcelTM (Vindimian, 2010).

III-2-6-3-2) Exposure of D. acuminata to C. variegatus

The following four experimental treatments, plus the appropriate controls, including (i) whole cell D. acuminata culture, (ii) live filtered D. acuminata cells resuspended in clean f/2 media, (iii) culture filtrate, (iv) culture lysate, and (v) f/2 medium as control, were created and assayed for

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Chapter 3-Part 2: D. acuminata and PTX2 towards fish adverse effects on sheepshead minnow larvae in a 96 h bioassay. A culture of D. acuminata (in exponential growth phase) was diluted in f/2 at salinity 25 to reach cell concentrations of 5, 50, 250, 800, 2500, and 5500 cells mL-1. This range of concentrations was used because it corresponds to a field range from moderate and naturally occurring blooms of Dinophysis of 102-104 cells L-1 (Gaillard et al., 2020b; Nagai et al., 2011) to rare, but exceptionally dense blooms of 105-107 cells L-1 (Belin et al., 2020; Reguera et al., 2012). Treatment (ii) was produced by filtering live cells of an active culture with 5500 cells mL-1 and resuspending them in clean f/2 medium to a final concentration of 5500 cells mL-1 of D. acuminata. The filtrate obtained from the previous cell filtering process (eq. 5500 cells mL-1) was again filtered using a 0.2 µm (cellulose) syringe filter and then employed as treatment (iii), culture filtrate (e.g. aqueous fraction). Treatment (iv), cell lysate or the intracellular compartment of D. acuminata, was produced by sonication (15 min at 25 kHz) of a whole live culture with 5500 cells mL-1 eq., followed by centrifugation (3500 g, 15 min, 4 °C) to remove cell debris. Cumulative mortality was assessed with n = 14 per treatment and routine parafin histology microscopy and transmission electron microscopy (TEM) with n = 8 and n = 2 fish, respectively, for each treatment.

III-2-6-3-3) Exposure of PTX2 to C. variegatus

The effect of PTX2 (50, 250, 500, 750, 1000, 2000 to 4000 nM), including the two controls (FSW and FSW + ethanol (EtOH) 0.8 % final) were investigated through the 96 h larval fish bioassay. A purified (>98%) and accurately quantitated standard of PTX2 was obtained from the National Research Council Canada (NRCC, Halifax, NS, Canada). The PTX2 standard was gently evaporated to dryness under nitrogen and resuspended in ethanol in order to avoid the negative effect of methanol on fish larvae observed in a preliminary experiment. The PTX2 solutions were prepared by dilution of the stock solution in FSW at salinity 25. The lowest concentration of 50 nM of PTX2 corresponded approximately to the content (intracellular + extracellular) of 2500 cells mL-1 of D. acuminata DAVA01 (see section “Results-Toxin analyses”). Higher PTX2 concentrations were tested as an increasing gradient to observe the effects on C. variegatus mortality and histopathology. For each condition, the replicates were n = 38, n = 8 and n = 2 for the cumulative mortality assessment, light microscopy and TEM, respectively.

III-2-6-4) Histology: light microscopy and TEM techniques

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Larvae destined for light microscopic evaluation were processed by routine methods for paraffin histology (Prophet et al., 1994). Briefly, fish were euthanized by 15 min exposure (500 ppm in 200 ppt filtered seawater) to the finfish anaesthetic, tricaine methanesulfonate (MS-222) (Find Source: I’ll go look at the jar and add) for 15 min. Fish were then (n = 8 per condition) fixed and decalcified by immersion for at least 48 h in Bouin’s solution, rinsed overnight in running tap water, and transferred to 70% ethanol for storage. Tissues were dehydrated through a graded ethanol series, cleared in xylene substitute and infiltrated overnight with warm melted paraffin in an automatic tissue processor (Shandon Hypercenter, ThermoFisher, Waltham, MA). Tissues were then embedded in paraffin on a tissue embedding center (Tissue Tech, Sakura Finetek USA, Torrance, CA), and resulting tissue blocks were sectioned at 5 µm on a rotary microtome (Olympus Cut 4055, Center Valley, PA) and stained with hematoxylin and eosin (H&E) in an automatic slide stainer (Shandon Varistain 24-3, ThermoFisher, Waltham, MA). Slides were cover-slipped, oven-dried and evaluated histologically and photographed on an Olympus AX-70 photomicroscope.

For the TEM analyses, fish were processed as described in Vogelbein et al., (2002). Briefly, fish were euthanized with tricaine methanesulphonate (MS-222: see above for Histology), and fixed in 4% glutaraldehyde with 5% paraformaldehyde in 0.1M sodium cacodylate (NaCac) buffer, pH 7.2, at 4° C for 2 h. Samples were washed in three changes of 0.1M NaCac buffer, 15–30 min each, and stored overnight at 4-8° C in a third change of buffer. They were post-fixed with 1% OsO4 in 0.1M NaCac buffer, pH 7.2, at 4° C for 1 h and then washed in three changes of 0.1M NaCac buffer, pH 7.2, 15–30 min each. Tissues were then decalcified in 5% EDTA/0.1 M NaCac at 4° C for 24 hrs. They were then dehydrated in a graded ethanol series, cleared in propylene oxide, and infiltrated in epoxy resin (Spurr’s Medium) over 4-5 days, and polymerized in a warming oven for 48 hrs at 58° C. Tissue blocks were then sectioned on an ultra-microtome (Reichert Ultracut E) at 70-90 nm using a diamond knife, with sections mounted on single hole, Formvar-coated copper grids and post stained with lead citrate and uranyl acetate (REF). Sections were evaluated and photographed in a Zeiss EM 100 transmission electron microscope operated at 80 Kv.

III-2-6-5) Toxin analyses

The stock culture of D. acuminata was sampled (1 mL, n = 3) at the beginning of the experiment to measure intracellular and extracellular toxins. Samples were centrifuged (3500 g, 15 min at 4°C)

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Chapter 3-Part 2: D. acuminata and PTX2 towards fish and separated into cells (pellets) and culture filtrate (supernatant). The pellets (intracellular toxins) were extracted with 0.5 mL methanol, sonicated for 15 min (25 kHz), centrifuged (3500 g, 15 min, 4 °C), filtered (0.2 µm) and stored in glass vials at -20 °C for analysis. The supernatants (extracellular toxins) were extracted by solid phase extraction (SPE) using Oasis HLB 60 mg cartridges (Waters, Milford, Massachusetts, USA). The SPE column was conditioned with 3 mL methanol followed by 3 mL high purity water, and the sample was then passed though the SPE column. The cartridge was rinsed with 3 mL of pure MilliQ water (Millipore) then dried for 1 min. Toxins were eluted with 1 mL of methanol into a glass vial and stored at -20 °C for analysis. Toxin analysis was performed using ultra-performance liquid chromatography–tandem mass spectrometry (UPLC-MS/MS) equipped with a trapping dimension (trap) and at-column dilution (ACD) (UPLC-MS/MS with trap/ACD) (Xevo MS, Waters Corp., Milford, MA, USA). The chromatographic separation was carried out on a C18 column (Acquity BEH C18 130 Å 1.7 µm, 2.1 × 50 mm) equipped with a trapping column (XBridge BEH C18 130 Å 10 µm, 2.1 × 30 mm) using the analytical method developed by Onofrio et al. (2020). Certified standards (dinophysistoxin-1 (CRM-DTX1-b), dinophysistoxin-2 (CRM-DTX2-b), okadaic acid (CRM-OA- d) and pectenotoxin-2, (CRM-PTX2-b), CNRC, Halifax, Canada) were used for external calibration.

III-2-6-6) Statistical analyses

Statistical analyses were performed using RStudio. Pearson’s Chi-square test were computed on the cumulative mortality observations, with random, exclusive (dead or alive) and independent data with expected values > 10 (Mchugh, 2013). Then, a post hoc test was performed on adjusted residuals of the Pearson’s Chi-square test with Bonferroni correction (MacDonald and Gardner, 2000).

III-2-7) Results

III-2-7-1) Toxins of D. acuminata

The toxin profile of the D. acuminata strain DAVA01 was mainly constituted by intracellular (> 95%) PTX2 (22 ± 0.50 pg cell-1) and to a much lesser extent, by intracellular (100%) OA (0.17 ±

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0.07 pg cell-1) and DTX1 (1.2 ± 0.64 pg cell-1) (Table 31). The maximum cell concentration of D. acuminata used in the exposure experiment was 5500 cells mL-1 and corresponded to 140 ± 3.20 nM of PTX2 (Table 31).

OA DTX1 PTX2 Total Total Total Total Total Total (nM in 5500 (nM in 5500 (nM in 5500 (pg cell-1) (pg cell-1) (pg cell-1) cells mL-1) cells mL-1) cells mL-1) D. acuminata 0.17 ± 0.07 0.94 ± 0.45 1.2 ± 0.64 8.1 ± 4.3 22 ± 0.50 140 ± 3.20 DAVA01

III-2-7-2) Cumulative mortalities

No mortality of C. variegatus larvae was observed when exposed to (i) the whole culture (from 50 to 5500 cells mL-1), (ii) resuspended cells (5500 cells mL-1), (iii) culture filtrate (eq. 5500 cells mL- 1) or (iv) lysate (5500 cells mL-1) of D. acuminata.

However, exposure of C. variegatus to PTX2 showed dose- and time-dependent responses (Figure 35). Indeed, 4000 and 2000 nM of PTX2 induced 100% mortalities in 24 h (P < 0.001), whereas at 1000 nM, only 3 % of the fish larvae died. After 48 h, significant mortality (63 %; P < 0.05) of C. variegatus larvae exposed to 1000 nM was observed and reached 100 % after 72 h of exposure (P < 0.01; Figure 35). For fish larvae exposed to the intermediate concentrations of 750 nM, mortalities were first observed at 48 h (37 %), and were significant after 72 h (P < 0.01), at which point all larvae had died. An exposure to 500 nM caused a non-significant increase in mortality after 72 h (26%), which became significant after 96 h (P < 0.05, 84%). A non-significant mortality of 8 % of the fish larvae exposed to 250 nM was observed at 96 h. The LC50 decreased with time, from 1231 nM at 24 h to 377 nM after 96 h (Table 32).

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A production of mucus was observed throughout the 96 h exposure by fish exposed to concentrations of PTX2 above 250 nM. Moreover, most of the fish exhibited an elevated respiratory rate compared to control and turned upside down from several minutes to up to several hours before death (pers. obs.). Finally, both FSW and FSW + ethanol controls showed no mortality of fish larvae throughout the experiment, but after 72 h, fish exposed to the FSW + ethanol produced some mucus.

100 * * * *

80 Conditions 70

% C FSW

y C EtOH

t i

l 60 a

t 50 nM r

o 250 nM

m 50

e 500 nM

i t

a 750 nM l 40

u 1000 nM m u 2000 nM C 30 4000 nM 20

0 24 48 72 96 Time h 1 Figure 35: Cumulative mortality (%) over 96 h in C. variegatus larvae exposed to 50, 250, 500, 750, 1000, 2000, 4000 nM of pectenotoxin-2 (PTX2), filtered sea water and ethanol (0.8 % final) controls (n = 38 for each condition). *significantly different (Pearson’s Chi-square test and post hoc test on residuals)

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-1 Table 32: LC50 (nM and ng mL ) of pectenotoxin-2 (PTX2) based on the Hill coefficient according to exposition time

Time (h) LC50 (nM) LC50 (ng mL-1) 24 1231 1057 48 873 750 72 530 455 96 377 324

III-2-7-3) Histology

Microscopic evaluation of gills in fish exposed to PTX2 and culture of D. acuminata (treatment (i)) revealed severe pathology (Figure 36 A-F). However, control animals exhibited typical gill structure indicative of health at 96 h (Figure 36 A). Exposure to 4000 nM of PTX2 for 24 h resulted in severe disruption of healthy gill structure (Figure 36 C) with swelling, disorganization (loss of normal cell to cell contact) separation, necrosis and sloughing of the respiratory epithelium and the osmoregulatory chloride cells, normally located in the surficial epithelium of the crypts between secondary gill lamellae (Figure 36 B). In the most severely affected individuals, there is extensive hyperplasia (e.g., proliferation) as well as hypertrophy (swelling with hyper-eosinophilic cytoplasmic staining), and redistribution of chloride cells, to a degree where the interlamellar spaces are completely occluded by these cells, many of which are necrotic (Figure 36 D). Similarly, exposure to 5500 cells mL-1 of live D. acuminata culture for 96 h led to milder chloride cell hyperplasia with partial occlusion of the interlamellar spaces in some individuals (Figure 36 F) but only very mild surficial damage in the gill respiratory epithelia (Figure 36 E). In contrast to the extensive and severe pathology observed in fish exposed to moderate to high doses of PTX2, we saw no significant impacts on the epithelial membranes lining the branchial chamber, nor the epidermis of the fish inner and outer body epithelium (Figure 36 C). Observations at lower PTX2 or D. acuminata concentrations and different time as well as TEM images and analyses of other organs (i.e. digestive tracts) are not available yet.

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Figure 36: Gill histopathology in 3-week old Cyprinodon variegatus larvae exposed to a range of pectenotoxin standards over 96 h. A. Control fish exhibiting healthy 1° gill filament (G1) and several healthy 2° gill lamellae (G2) at 96 h, comprised of an outer simple, flattened squamous respiratory epithelium overlying the vascular spaces, lined and maintained by pilaster (pillar) cells and filled with nucleated red blood cells (X40). B. Extensive disruption of gill structure with swelling (Sw), separation, necrosis and sloughing of respiratory epithelial and osmoregulatory chloride cells (Cl) in fish exposed to 4000 nM Ptx2 (X60). C. Low magnification overview of branchial chamber in fish exposed to 4000 nM pectenotoxin 2 (PTX2) showing two gill arches (Ga), damaged (De) surficial tissues of the gill filaments and lamellae, but intact inner chamber lining epithelium (Ie) and outer epidermis (Oe) sampled at 24 h (x4). D. Total occlusion of interlamellar spaces by enlarged eosinophilic chloride cells (Cl) and sloughing respiratory epithelium. Some evidence of cell necrosis (X60). E. Gill lamellae of fish exposed to 5500 cells mL-1 of D. acuminata culture sampled at 96 h and exhibiting largely intact, presumably functional lamellar tissue architecture. (G1) Primary gill filament (X40). F. Gill lamellae of fish exposed to 5500 cells mL-1 of D. acuminata culture sampled at 96 h. Partial occlusion of interlamellar spaces by hypertrophied and necrotic chloride cells (Cl) (X20).

III-2-8) Discussion

This work aimed to determine the effect of a common HAB species, Dinophysis acuminata and one of its toxins, PTX2, on larvae of a common estuarine fish species, Cyprinodon variegatus using a 96 h larval finfish bioassay.

Exposure to PTX2 led to significant and rapid mortalities of fish larvae, from 8-84 % after 96 h to 100 % after 24 h, for low-intermediate (i.e. 250-500 nM) to high (i.e. 2000-4000 nM) concentrations, respectively, as well as severe damage to the gill respiratory epithelia and the osmoregulatory chloride cells. Similar but milder damage to gill tissues was observed when larvae were exposed at the highest cell concentration of a live D. acuminata culture (i.e. 5500 cells mL-1) for 96 h. A significant finding was that no mortality was observed in the exposures of larvae to the live Dinophysis culture.

In a recent study, Rountos et al. (2019) exposed newly hatched and one-week old fish larvae, C. variegatus and M. beryllina to 1000 cells mL-1 of toxic strains of D. acuminata (producer of < 1

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Chapter 3-Part 2: D. acuminata and PTX2 towards fish pg cell-1 of OA and DTX1) and/or Alexandrium catenella (producer of < 2 pg cell-1 of saxitoxins) for six days. While no lethality was observed in larvae exposed only to D. acuminata (i.e. no direct reduction of time to death), a sub-lethal effect was highlighted by a decrease in larval growth rates (Rountos et al., 2019). Our results support their observations, because no lethal effect was observed on 3-week-old larval C. variegatus, even at a higher concentration (i.e. 2500-5000 cells mL-1) of a more toxic strain of D. acuminata (i.e. DAVA01 strain).

Histological observations of both exposure experiments revealed significant hyperplasia and hypertrophy, redistribution and/or increase in the number of chloride cells (= ionocytes), especially at the higher concentrations of PTX2. In fish, chloride cells are located at the base of the secondary gill lamellae and constitute part of the epithelium that lines the primary gill filaments during health (e.g., under non-stress conditions) (Dymowska et al., 2012). These ionocytes play a critical role in regulation of hydro-mineral balance and are involved in ion transport (e.g. sodium and calcium), pH regulation and ammonia excretion (Hwang et al., 2011; Laurent and Perry, 1991; Osborne et al., 2016). However, when gills are exposed to environmental disturbances their morphology has been described to follow a common mechanism that typically results in “branchial chloride cell proliferation” (Laurent and Perry, 1991). For instance, exposure to some xenobiotics can lead to modification of morphology, distribution and number of chloride cells in rainbow trout (Laurent and Perry, 1991).

Such branchial alterations have also been observed with exposure of finfish to OA. Larval Gilt- head bream (Sparus aurata) exposed to OA (µM concentration for 2 to 24 h), displayed enlargement of secondary lamellae in gills after 4 h of exposure (Souid et al., 2018). The authors hypothesized that OA caused collapse of pillar cell and breakdown of the vascular components of the gill respiratory tissues. Similar to our results, exposure of Zebrafish (Danio rerio) and C. variegatus larvae to karlotoxins 1 and 2 (i.e. ichtyotoxins produced by species of the genus Karlodinium; Hallegraeff et al., 2017; Place et al., 2012) for 1-4 h at 50-500 ng mL-1, led to gill damage, including hypertrophy of epithelial cells, fusion of secondary lamellae, and proliferation and redistribution of chloride cells (Deeds et al., 2006). The authors suggested that exposed larvae died from respiratory failure due to gill damage. Similarly, ichtyotoxins produced by the haptophyte Prymnesium parvum (i.e. prymnesins) have been shown to cause gill damage in fish (Ulitzur and Shilo, 1966), in particular, affecting the chloride cells (Terao et al., 1996), and

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Chapter 3-Part 2: D. acuminata and PTX2 towards fish decreased oxygen consumption per breath and oxygen extraction leading to respiratory failure in fish (Bergsson et al., 2019; Svendsen et al., 2018). Thus, morphological/functional alterations of chloride cells and/or gill respiratory lamellar epithelia appear to be a common result of exposure to diverse algal toxins. Laurent and Perry (1991) found that xenobiotics which affect ion transport (e.g. Zn2+ which promotes ion loss from cells), can have a significant impact on chloride cells. While it has been shown that goniodomin A could form a complex with ions (e.g potassium) (Tainter et al., 2020), the formation of similar complexes with PTX2 (i.e. structurally related to goniodomin A) or other DSTs have not been thoroughly studied. We can only hypothesize a non- specific interaction between PTX2 and chloride cells in C. variegatus larvae.

Gill lamellae alterations were also observed, mostly when larvae were exposed to high concentrations of PTX2. In addition, swelling, disruption, separation, necrosis and sloughing of the respiratory epithelial cells lining secondary lamellae, was an important observation in this study. The mode of action of PTX2 is recognized to due to an interaction with, and functional disruption of, the critical cellular protein, actin (Allingham et al., 2007; Hori et al., 1999). The respiratory surface of secondary gill lamellae is essentially composed of two simple flattened epithelial membranes, that allow gas exchange between water and the circulating red blood cells. The integrity of these two membranes is maintained by one of them, the thin specialized vascular endothelium, in the secondary lamellae, called pilaster (pillar) cells (Bettex Galland and Hughes, 1973). Pillar cells are rich in actin fibers, e.g. F-actin (Kudo et al., 2007) that play a critical role in maintaining their very characteristic cellular shape, in their role in maintaining the blood vascular spaces that allow efficient gaseous exchange between the water and secondary lamellae. Thus, PTX2 that binds to pillar cell actin may have interfered with their ability to maintain cell integrity, initiating a cascade of events leading to gill lamellae alterations.

Here we also show that exposure to natural field concentrations of D. acuminata and PTX2 leads to alterations of chloride cells which may modify ion transport and osmotic balance of fish, and may thus be detrimental for larvae. Moreover, we observed damage of fish gill tissue in larvae, which may have affected respiration and decreased acquisition of oxygen, which was also suggested by increased breathing frequency. Thus, these exposures could ultimately have modified energy balance and affected primary metabolism (Van Winkle et al., 1997).

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Grazing rates were not measured in our experiments, but according to Rountos et al. (2019), D. acuminata cells were actively consumed by fish larvae at a rate of 5-10 cells.h-1, which may suggest that D. acuminata cells were also consumed in our experiment by older planktivorous larvae. Moreover, as water enters in contact with gill lamellae for respiration, contact between gills and algal cells may occur (Landsberg, 2002; Laurent and Perry, 1991) which may have played a significant role in the damage to gill epithelia reported here for the highest concentrations of D. acuminata.

Additional questions arise from our observations. (i) Why did Dinophysis exposure not induce any mortality of fish larvae? If the effects (i.e. gill lamella and chloride cell alterations and mortality) were only attributable to PTX2, the extracellular concentration of PTX2 in D. acuminata culture in this study (i.e. less than 3 pg eq. 5500 cells mL-1) was not sufficient to induce mortality. Moreover, even the lysate of D. acuminata which contained in total 140 nM for 5500 cells mL-1 was not lethal. In 1-day old Zebrafish larvae (Danio rerio), the LC50 of OA and DTX1 after 24 h was approx. 12500 and 8500 nM, respectively (Figueroa et al., 2020); these were higher concentrations than were present in our D. acuminata strain used in this study (approx. 1 and 10 nM of OA and DTX1 for 5500 cells mL-1 in total, and < LD in extracellular). These concentrations

(e.g., Figueroa et al., 2020) were also10 times higher than the LC50 obtained for PTX2 (i.e. 1231 nM after 24 h in this study). The results suggest a higher toxicity of PTX2 than DSTs towards fish larvae. The combined effects of combined exposure of both DSTs and PTXs towards fish should be further investigated, since both of these compounds have shown effects towards Caco-2 cells (Alarcan et al., 2019). (ii) How did C. variegatus larvae die? Our preliminary observations of pathologies in gills suggested that fish may have died from respiratory failure with a potential reduction or loss of osmoregulatory ability. This hypothesis is strengthened by the observations of an increase in breathing frequency and mucus production in fish compared to controls, as previously reported for European plaice (Pleuronectes platessa) exposed to harmful concentrations of the ichtyotoxic haptophyte, Prymnesium parvum (Bergsson et al., 2019). (iii) Are PTX2-derived mortalities only caused by the alterations observed in the gills? As the main target of DSTs and PTX2 by oral administration and intraperitoneal injections is the liver (Terao et al., 1993, 1986), mortalities following exposure to PTX2 could be associated with negative effects on hepatocytes and/or other digestive tract tissues, if toxins are ingested. Further observations by light microscopy and TEM will focus on liver and digestive tracts in both exposure experiments.

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Finally, the lethal and sublethal effects observed here (due to both D. acuminata cells and PTX2) towards C. variegatus larvae may have ecological and economic implications. Blooms of Dinophysis, that are known PTX2-producers (Onofrio et al., 2020), are recurrent in the Chesapeake Bay and overlap with the spawning season of C. variegatus (i.e during spring-summer) and numerous other sensitive species (Rountos et al., 2019; Wolny et al., 2020). Indeed, this estuarine fish is preyed on by top predators of economic importance, such as the Striped bass (Morone saxatilis) in the Chesapeake Bay (Gervasi et al., 2019).

Although this study highlighted harmful effects of D. acuminata and PTX2 at relatively high concentrations compared to the usually observed concentrations of Dinophysis blooms of 102 cells L-1 in the Chesapeake Bay and globally 103-104 cells L-1 (Reguera et al., 2012; Wolny et al., 2020), further light microscopy observations and TEM tissue sections will determine the potential harmful effects at more ecologically relevant concentrations. Overall, the reports of harmful effects of the non-diarrheic PTX2 towards marine animals (i.e. early life stages of C. variegatus in this study and C. gigas in Gaillard et al., (2020b)) are novel and should be taken into account to better assess risks associated with Dinophysis blooms.

III-2-9) Acknowledgements

This work was funded by the project CoCliME which is part of ERA4CS, an ERA-NET initiated by JPI Climate, and funded by EPA (IE), ANR (FR), BMBF (DE), UEFISCDI (RO), RCN (NO) and FORMAS (SE), with co-funding by the European Union (Grant 690462). We thank Satoshi Nagai (National Research Institute of Fisheries Science, Japan) for Japanese strains of T. amphioxeia and M. rubrum.

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III-2-10) Supplementary materials

Table 33: Origins and culture conditions for the three organisms required to actively culture Dinophysis acuminata (strain DAVA01) used in this study

Temperature Irradiancea Species Origin Strain ID Medium (°C) (µmol photons m-2 s-1) Teleaulax Inokushi Bay, Oita JATA- f/2 15 ± 1 ~ 100 amphioxeia Prefecture (Japan) AB364287 Mesodinium Inokushi Bay, Oita JAMR- f/12 15 ± 1 ~ 100 rubrum Prefecture (Japan) AB364286 Nassawadow, Dinophysis Chesapeake Bay, VA, DAVA01 f/2 15 ± 1 ~ 100 acuminata USA a Cultures were subjected to light in the PAR domain with a circadian 12 h: 12 h (light: dark) cycle.

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General discussion

Toxic species of the ubiquitous genus Dinophysis are known as public health, economic and societal threats in coastal marine socio-ecosystems. Indeed, although the cellular concentrations of Dinophysis are usually low in most coastal areas (i.e. hundred cells per liters; Reguera et al., 2012), they can synthesize diarrheic toxins and be the origin of DSP outbreaks (Yasumoto et al., 1980, 1978) that annually lead to bans of shellfish farming activities at a worldwide scale.

Before 2006, ecophysiological studies on Dinophysis and chemical analyses (i.e. toxin contents) were performed on cells sampled directly from the natural environment. Park et al., (2006) established the first permanent culture of D. acuminata (i.e. Teleaulax amphioxeia. < M. rubrum < Dinophysis acuminata) based on the mixotrophy of this organism and of its prey, M. rubrum. This scientific advance allowed experiments with Dinophysis using controlled culture conditions, which improved understanding of its toxicity and factors affecting its physiology. Moreover, characteristics of the first prey organism of its food chain, Teleaulax amphioxeia, as well as the second, M. rubrum (Riisgaard and Hansen, 2009), may also affect Dinophysis physiology (i.e. toxin production, growth or photosynthesis), metabolome (García-Portela et al., 2018a) and subsequently its bloom dynamics (Peterson et al., 2013).

This thesis project focused on the potential effects of environmental parameters of global change on the physiology of Dinophysis and its food chain. Indeed, modifications of environmental conditions (i.e. abiotic or biotic factors), which occur at different scales (oceanic or coastal waters), may, in principle, modify Dinophysis bloom dynamics or species distribution and could therefore lead to exhacerbated negative effects on marine organisms and on human health and activities (Hallegraeff, 2010; Wells et al., 2015).

In this context, the effects of temperature, irradiance and pH (i.e. global change key factors; Wells et al., 2015) on growth, pigment contents and Fv/Fm of Teleaulax amphioxeia were determined using a full factorial design to detect direct effects of and interactions between factors. Subsequently, we assessed the effects of contrasting environmental conditions on the physiological status of T. amphioxeia (i.e. after acclimation to two light intensities) and of the quantity of prey

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on Mesodinium rubrum growth, pigment contents and Fv/Fm (Chapter 2-Part 1).

As mentioned in Chapter 1 (section “I-5-3) Salinity”), there are no studies focused on the effects of rapid variations in salinity on the growth and metabolite synthesis in Dinophysis cultures. However, this factor affects Dinophysis distribution and abundance at a regional scale (e.g. Ninčević-Gladan et al. (2008)). In the present thesis, the short-term effects of sudden changes in salinity (i.e. hypo- and hyperosmotic conditions) on the growth, biovolume, toxin and metabolite contents on three French isolates of D. sacculus were assessed (Chapter 2-Part 2).

In the literature, the effects of OA-group and PTXs toxins and their mechanisms of actions in mammalian models and cell lines have been well described (Chapter 1 section “I-6) Toxins of Dinophysis”). Moreover, several studies reported the presence and the accumulation of these toxins in marine organisms (Table 15 online at: https://doi.org/10.17882/76433). However, negative effects of OA-group toxins and live Dinophysis cells (from cultures) were observed on fish and filter-feeding bivalves (section “I-8) Effects of Dinophysis, OA-group toxins and PTXs on marine organisms”), but there is a lack of experiments of the effects of PTX2 producers and PTX2, including on critical early life stages of oysters or fish. To overcome this gap in knowledge, this work aimed at characterizing the effects of D. acuminata and D. sacculus isolates, and OA and PTX2 on the quality of gametes and fertilization success of the Pacific oyster (Crassostrea gigas) (Chapter 3-Part 1). In addition, the effects of D. acuminata and PTX2 on fish larvae (Cyprinodon variegatus) were determined by examining both mortality and histopathological features (Chapter 3-Part 2), in collaboration with Wolfgang Vogelbein from VIMS (Virginia, USA).

The effect of key factors of global change on T. amphioxeia

The model obtained, using a full factorial design experiment with T. amphioxeia (Spanish strain AND-A0710) cultures described an optimal theoretical growth rate of 0.88 d-1 at 17.6 °C, 400 µmol photons m-2 s-1 and a pH of 7.6, with significant direct and combined effects of temperature and irradiance (Chapter 2-Part 1). The tolerance of T. amphioxeia to abiotic factors was observed throughout the experimental domain, i.e. at a temperature of 15 to 22.6°C, a pH of 6.5 to 8.6 and an irradiance of 40 to 646 µmol photons m-2 s-1 (Chapter 2-Part 1). These results suggest that T. amphioxeia may not be negatively affected by modification of environmental conditions expected

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General discussion in the context of climate change, which in Western European Atlantic waters would be, i.e. warmer temperature, higher irradiance and lower pH (Gobler et al., 2017; IPCC, 2013; Orr et al., 2005; Wells et al., 2019, 2015).

Moreover, as this work was first performed in stress conditions, it highlighted the ecophysiological plasticity of a Spanish strain of T. amphioxeia (AND-A0710). Fiorendino et al. (2020) recently observed similar trends in tolerance to temperature (12 to 27 °C) and irradiance (50 to 400 µmol photons m-2 s-1), as well as tolerance to salinities (22 to 34) for a Danish (K-0434) and a Texan (GoMTA) strain of T. amphioxeia. Sala et al. (2015) showed the positive effect of acidification on the abundance of nanophytoplankton (e.g. as T. amphioxeia) in culture, in agreement with observations made in this work. In addition, in a recent study, Altenburger et al. (2020) discovered the digenetic life cycle of T. amphioxeia, which is the diploid stage of a life cycle where Plagioselmis prolonga is the haploid stage of the same species. This life cycle might explain the success of T. amphioxeia, because the two life cycle stages have adaptations for different environmental conditions (e.g. nutrient availability, light, temperature) and thus are able to tolerate a large range of environmental conditions throughout the year. However, results here showed intraspecific variability (as for e.g. macronutrient contents) (Lee et al., 2019), possibly suggesting a cryptic diversity of the cryptophyte (Johnson et al., 2016), thus the generalization of the plasticity of T. amphioxeia should be assessed taking into account the species level in future studies.

The effect of contrasting physiological status and culture density of the prey, T. amphioxeia, on its predator M. rubrum

The tolerance of T. amphioxeia to future ocean conditions would, in principle, lead to more of this prey organisms being available for predators, such as M. rubrum. The ciliate fed with two distinctly photo-acclimated cultures of T. amphioxeia (acclimated to lower or higher levels of light, -2 -1 respectively 100 and 400 µmol photons m s ) did not show any difference in growth rate, Fv/Fm and pigment contents. However, the prey concentration, and thus chloroplast concentration and particulate nutrients from prey, directly affected both growth rate and pigment contents of M. rubrum (Chapter 2-Part 1; also in agreement with Yih et al. (2004)) (Figure 37). Still, the abundance of M. rubrum in a hypothetical scheme of future climate conditions (Figure 37) also

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General discussion depends on several factors, e.g. (i) the effects of global change key factors on M. rubrum and (ii) competition with other grazers for prey organisms.

(i) Danish strains of M. rubrum have been shown to grow at high temperature (up to 24 °C) and irradiance (up to 400 µmol photons m-2 s-1) (Fiorendino et al., 2020), and tolerate salinity (from 22 to 34) and pH (from 7.8 to 8.8) fluctuations (Smith and Hansen, 2007). Moreover, a Spanish strain of M. rubrum is able to grow in a temperature range from 15 to 25 °C (Rodríguez et al., 2012) and light intensity from 10 to 650 µmol photons m-2 s-1 (García-Portela et al., 2018b) suggesting that future climate conditions would not be detrimental for the intermediate level of the food chain of Dinophysis species.

(ii) Other phytoplankton organisms, including several dinoflagellate genera as well as the ciliates, are heterotrophic or mixotrophic and feed T. amphioxeia (e.g. Karlodinium armiger, Mesodinium spp.; (Berge et al., 2008; Garcia-Cuetos et al., 2012; Moestrup et al., 2012) but are not prey specialists. Moreover, they can grow at a higher rate than the Spanish M. rubrum (0.31 d-1 Chapter 2-Part 1 and 0.39 d-1 García-Portela (2018b). For example, M. pulex have shown a growth rate of 0.8 d-1 for (Tarangkoon and Hansen, 2011)) and thus appears to be more competitive than M. rubrum (Figure 37). Competition between M. rubrum and other organisms, due to allelopathy or behavioral competition for prey (e.g. tolerance to prey diversity, difference in ingestion rate, efficiency of feeding mechanisms), should be explored in vitro, e.g. in cultures (e.g. mixotrophic ciliates) acclimated to future abiotic conditions.

Dinophysis in future climate conditions

Different questions remain concerning Dinophysis in the theoretical scheme of future climate conditions (Figure 37). How would (i) growth and (ii) toxin production of Dinophysis be affected by global change abiotic and/or biotic factors?

(i) Experiments on the effects of abiotic factors on Dinophysis (Chapter 1 section “I-5) Ecophysiological experiments with Dinophysis species”) mostly showed an increase of growth rate with increasing temperature (e.g. Kamiyama et al., 2010), irradiance (e.g. Kim et al., 2008) and nitrogen (i.e. regenerated forms; see García-Portela et al. (2020)), as well as a tolerance to salinity fluctuations (this study from at least 25 to up to 42 for three strains of D. sacculus; Chapter 2-Part

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2 and Fiorendino et al. (2020)). This suggests that Dinophysis cf acuminata can perform well in a wide range of abiotic conditions, including various future climate scenarios. However, as Dinophysis is an obligate non-constitutive mixotroph (Kim et al., 2008), both light and prey availability are required for its growth (Chapter 1 section “I-3-3) Mixotrophy of Dinophysis”) (Figure 37).

Competition for prey (i.e. M. rubrum) between Dinophysis and other grazers (e.g. dinoflagellates) may happen in the environment. Competition for M. rubrum is enhanced by the fact that it is usually found at concentrations as low as 1 cell mL-1 in the environment (Smith and Hansen, 2007). These low concentrations may limit the heterotrophic growth of its predators (Park et al., 2013). In a field study,, the dinoflagellates Amylax buxus and Amylax tricantha were found to retain organelles, e.g. plastids, mitochondria and nuclei, from a cryptophytic origin (Koike and Takishita, 2008). In culture, A. tricantha has been shown to feed only on M. rubrum (Park et al., 2013) and this species can outgrow D. acuminata due to its higher growth rate, thus has a competitive advantage (Park et al., 2013).

In addition, Dinophysis appears to be less competitive compared to generalist constitutive mixotrophs (e.g. Alexandrium pseudogonyaulax, Karlodinium armiger) (Blossom et al., 2012; Hansen et al., 2019; Hansen and Tillmann, 2020). Indeed, K. armiger is a type 2 constitutive mixotroph which is, by definition, able to largely increase (5-10 fold) its growth rate in the presence of prey (i.e. carbon uptake from prey is high, thus requiring an important prey concentration) (Berge et al., 2008) as compared with type 1, which only marginally increase their growth rate in the presence of prey (Hansen and Tillmann, 2020). However, Dinophysis species may, in part, compensate their disadvantage compared to type 2 constitutive mixotrophs by their capacity to be mainly photosynthetic in prey-limited conditions (Hansen et al., 2013). Overall, it could be interesting to compare growth of Dinophysis with other constitutive and non-constitutive mixotrophic organisms in different abiotic environmental conditions as such factors may influence feeding rate and photosynthetic performance (Hansen and Tillmann, 2020).

(ii) No abiotic or biotic factors that directly modulated toxin production in Dinophysis have been clearly identified (Chapter 1 section “I-5) Ecophysiological experiments with Dinophysis species”) (Figure 37), despite a few exceptions (e.g. higher PTX2 content at low temperature; Kamiyama et al., 2010; Rodríguez et al., 2012). However, the toxic potential of a culture (i.e.

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General discussion amount of DSTs per volume of culture) increases with the increase of cellular concentration (e.g. Basti et al. (2018)), the latter showing a positively correlation with temperature, irradiance, nutrients and/or prey concentration. Thus, if global change led to higher Dinophysis concentrations and abiotic and/or biotic factors did not decrease toxin production, this scenario in principle would lead to an increase of overall toxicity (Figure 37). In future studies, it could be interesting to assess which abiotic/biotic factors directly modulate toxin synthesis of Dinophysis in order to determine e.g. potential trends in toxin production in future ocean conditions and the biosynthesis pathway of these metabolites. In addition, considerations of the combined effects of such factors at all levels of the Dinophysis food chain, may shed light on the potential complexity of toxin production by Dinophysis species.

 temperature + Teleaulax  irradiance + ± salinity amphioxeia ± pH Grazers competition +  temperature + Mesodinium +  irradiance rubrum ± pH

Grazers ± salinity competition  temperature + + ± salinity Dinophysis +  irradiance + Toxicity?  nutrients

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Part 1 and Chapter 2-Part 2) and grey arrows represent abiotic factors tested in the literature. Dotted arrows indicate putative effects. Arrowheads with “+” mean growth enhancement. Factors preceded by “±” mean tolerance to increase and decrease. The arrows size is not informative.

In order to better understand Dinophysis distribution and bloom dynamics (e.g. onset, duration) in coastal waters under future climate conditions, it is interesting to study current and past trends of the effect of environmental factors on Dinophysis species. Such work was made regionally as part of the CoCliME project (see Gaillard et al. 2019 EPC7 Zagreb for the French Atlantic and English Channel coasts in an analysis of data collected from the French monitoring database (REPHY)). Over the last 20 years, neither statistical correlations nor global trends were observed either in Dinophysis abundances and bloom durations, nor in temperature and salinity changes. Additionally, no correlation was observed between Dinophysis abundance and environmental factors. However, strong variability between regions was observed, which could explain the absence of global trends in France. In addition, the identification of Dinophysis in the REPHY database is made at the genus level, although it is known that species (e.g. D. acuminata vs. D. acuta) have different seasonal and spatial patterns (spring-summer blooms in turbulent areas vs. late summer blooms in stratified areas) (see references in García-Portela et al. (2019)), with patchy distribution at usually low concentration (ca. 1 cell L-1) implying sampling and counting are challenging and may bias statistical analyses from the database(s) (similarly, Australian monitoring database Farrell et al. (2018)). In contrast, a global (worldwide) modelisation study showed an increase of bloom duration (+ 3 weeks) and growth rate (+ 0.01 d-1) of D. acuminata with increasing sea surface temperature over the last 30 years in the North Atlantic and Pacific regions (Gobler et al., 2017). In the range of French Atlantic coasts, we hypothesize that the stability of current and past (20 years) environmental abiotic conditions measured were neither more detrimental nor more favorable to Dinophysis, M. rubrum and T. amphioxeia. This could partly explainwhy no trends in distribution and bloom dynamic of Dinophysis were observed regionally in French coastal waters over the last two decades.

Dinophysis is a globally distributed genus (e.g. D. acuminata is cosmopolitan; Chapter 1 section “I-10) DSTs distribution”) and is present throughout the year but not always forming blooms (Chapter 3-Part 1). This suggests a good adaptation to its environment, as reflected by the wide

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General discussion range of tolerance to e.g. salinity and temperature of several toxic species (Nagai et al., 2020) and by the fact that no resting forms (e.g. cysts) have been clearly identified for Dinophysis (Reguera et al., 2012).

In order to improve knowledge on the regional distribution of Dinophysis, a focus was made on the effect of salinity on Dinophysis. In environmental studies, Dinophysis distribution and abundance were often correlated with salinity variations (e.g. Caroppo et al. (2001)). Moreover, D. sacculus was found to perform well in waters subjected to quick salinity variations, such as closed, semi- enclosed and estuarine areas (Zingone et al., 1998), but physiological studies of the effects of a salinity stress on Dinophysis were lacking.

The osmoregulation process in D. sacculus

Salinity stress may affect microalgae due to differences in osmolarity between the internal and external environments. In order to maintain an equilibrium, microalgae can rely on several physiological mechanisms, such as the osmoregulation process, which consists in adjustments in turgor pressure and ion concentrations, and synthesis and accumulation of osmolytes (Kirst, 1989).

Three strains of D. sacculus from French Atlantic and Mediterranean coasts had a good tolerance after 24 h of sudden hypoosmotic (25) and hyperosmotic (42) stress, as shown by a stable growth rate, biovolume, toxin content and metabolomic profiles compared to the control (35) (Chapter 2- Part 2).

The salinity tolerance of three strains of this species (or ecotype of the D. acuminata-complex), whatever their origin, may in part be explained by the synthesis of osmolytes, i.e. proline, glycine betaine and DMSP. However, significant increase of osmolyte (i.e. glycine betaine) from the control to the hyperosmotic condition, was observed only for one strain, thus suggesting the role of other potential osmolytes and/or mechanisms (e.g. modification of ion ratios) of osmoregulation (Chapter 2-Part 2). The osmoregulation process of Dinophysis could be better investigated by performing similar experiments with other species outside of the D. acuminata-complex which have been observed in a broader spatial range, e.g. D. acuta.

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The identification of possible unknown osmolytes should be conducted using more contrasting salinity levels on cultures with significantly higher cell densities. Even if it was not successful here, metabolomics (after the development of a more appropriate LC method for osmolytes - small and polar metabolites - based on HILIC) is still the best approach to detect metabolites ot known a priori. A bottleneck of this approach is the annotation (i.e. identification of these metabolites) as classical dereplication strategies rely on databases which are currently limited but might be more comprehensive in the future. Hopefully, alternative and more efficient dereplication tools will emerge (Wolfender et al., 2019), facilitating the identification of metabolites such as the osmolytes of Dinophysis species.

Intraspecific variability in D. sacculus

The next section discusses intraspecific variability observed in the metabolomic profiles of three strains of D. sacculus, as well as variability in toxin content, during the salinity stress experiment, and develops hypotheses about the origin of the populations. The three D. sacculus strains had been kept for 2-5 years with the same culture conditions (abiotic factors and prey), thus reducing changes in metabolomic profiles related to prey origin (as described in(García-Portela et al., 2018a). The metabolome differences between the Mediterranean strain and the two Atlantic strains are likely to be explained by their different geographical origin and suggest that strains may evolve keeping their population separated. However, metabolomes of the two Atlantic strains, although geographically closer, could also be differentiated.

Batifoulier et al. (2013) showed that Dinophysis (genus level identification) in Arcachon Bay did not develop directly in the Bay but originated from Capbreton (Southern France, near the Spanish border) and were advected (i.e. transported) into the bay from the open ocean. This suggests that a pool of a similar population (i.e. similar species occurring in the same place in the same time) may be transported across several locations, i.e. Dinophysis pool originated from stochastic introduction by advection (see Rengefors, 2020).

Metabolomic observations made in the present work could suggest that the two Atlantic D. sacculus are ecologically different populations (i.e. each developing in its location and may be genetically and phenotypically different). However, differences in terms of gene expression (Le

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General discussion

Gac et al., 2016), physiology (e.g. growth) (Brandenburg et al., 2018) and qualitative and quantitative toxin profiles (Martens et al., 2017) were observed between a high number (> 15) of strains of another dinoflagellate genus, Alexandrium, sampled in the same population for,. Thus, this variability suggests that differences in metabolite expression happens in the same dinoflagellate population as could be the case for Dinophysis.

Further studies are needed to explain the distribution of Dinophysis species, including D. sacculus in French Atlantic coastal waters and to identify different populations. Nevertheless, this would require different tools than those used in this work, metabolomics analysis, which is but a “snapshot” of the physiological status.

Roles and implications of the OA-group toxins and PTXs

The roles of the OA-group toxins and PTXs are still unknown (Reguera et al., 2012), but they are discussed here from different perspectives, i.e. (i) physiological and cellular perspectives, (ii) population and competition perspectives and (iii) environmental and societal perspectives (Figure 38).

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Roles and implications of Dinophysis toxins Physiology and Population and Environmental intracellular extracellular and societal perspectives perspectives perspectives Osmoregulation? Allelopathy Sanitary Complexation Growth DSP between toxins inhibition of OA- and ions group toxins Photosynthesis? Defense Initiation of Grazing by synthesis copepods Marine … ressources Prey capture Bioactive Effects in vivo? compounds and in vitro towards fish and PUFA? bivalves … …

Figure 38: The roles and implications of DSTs and PTXs on several aspects at different scales, from cellular to societal levels.i.e. physiological and intracellular, population and extracellular, and environmental and societal. Blue font represents aspects tested in this thesis (Chapter 2-Part 2, Chapter 3-Part 1 and Chapter 3-Part 2) and grey font represents aspects observed, tested and/or hypothesized in the literature.

(i) The physiological role of intracellular toxins of Dinophysis as osmolytes has been investigated in Chapter 2-Part 2 (Figure 38). Errera and Campbell (2011) suggested that brevetoxins could facilitate osmoregulation by interaction with ion channels. In this experiment OA and PTX2 were neither accumulated nor released, thus did not act like compatible solutes, in contrast to proline, glycine betaine and DMSP. However, OA and PTX2 may interact (e.g. by complexation) with ion/ions channels, that accounts for an osmoregulation mechanism (Hagemann, 2011; Kirst, 1989)

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General discussion as they have been shown to interact with sodium, calcium and potassium ions (Blaghen et al. 1997, Norte et al. 1998, Tainter et al. 2020) (Figure 38), thus, may modulate ion fluxes in the algal cells. However, such interaction in toxic microalgae is unknown and should be explored as suggested in Chapter 2-Part B with epifluorescence, NMR or radiotracer methods.

The biosynthetic pathways of OA-group toxins and PTXs are still unknown, this explains why hypotheses about the physiological roles of these toxins in literature are scarce. Okadaic acid has been shown to be located in the periphery of Prorocentrum chloroplasts, and suggested to be “a product of photosynthesis machinery” (Zhou and Fritz, 1994). Although this hypothesis has never been proven, it is unlikely in view of the current knowledge on Dinophysis mixotrophy. However, Jia et al. (2019) showed that toxin production was indeed initiated by light during the G1 cell cycle phase, which appeared to be an important factor for the initiation of the synthesis of algal toxins (i.e. DTX in Prorocentrum) (see Cembella and John, 2006) (Figure 38). Future experiments should go deeper in the determination of factors which trigger or inhibit toxin production, and perform transcriptomic analyses to determine the mechanism of toxin synthesis by Dinophysis. That was the way followed for the discovery of domoic acid biosynthetic pathway (Brunson et al., 2018).

(ii) The role of extracellular DSTs and PTXs in terms of population and competition aspects towards other marine organisms (i.e. allelopathy) is discussed here. The effects could be classified as competition with other phytoplankters, defense against grazers and prey capture (Figure 38).

In a preliminary experiment (data not presented), we used a fluorescence-based bioassay (Long et al., 2018a), which determined the allelochemical potential of a microalgae towards a target diatom,

Chaetoceros muelleri (based on the modification of Fv/Fm). But no allelochemical interactions could be observed after 2 h exposure of the same diatom to D. sacculus (500 cells mL-1; strain used in Chapter 3-Part 1). This absence of effects could suggest that i) D. sacculus did not synthesize allelochemicals, or ii) that 500 cells mL-1 were not enough to lead to allelopathic effects, and/or iii) that the allelochemicals from D. sacculus are different to the bioactive extracellular compounds produced by Alexandrium that affect both C. muelleri and the oyster C. gigas (Castrec et al., 2018; Long et al., 2018a). Allelopathic effects of Dinophysis acuminata on its ciliate prey have been reported during the prey capture process (Mafra et al., 2016; Ojamäe et al., 2016) and Mafra et al. (2016) suggested that the allelopathic substance could be polyunsaturated fatty acids (PUFAs) (Chapter 1 section “I-8-1) Allelopathic effects”) (Figure 38). However, the effects of toxins of the

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General discussion

OA-group towards marine phytoplankton growth have been observed, also suggesting an allelopathic effect (e.g. Windust et al., 1996) (Chapter 1 section “I-8-1) Allelopathic effects”), but high µM concentrations of toxin were used in the experiment (Figure 38). In addition, experiments on the feeding behavior of copepods on D. acuminata and D. norvegica (Carlsson et al., 1995; Jansen et al., 2006) showed that copepods may preferentially feed (or not) on Dinophysis. The authors also highlighted toxic effects towards copepods (lower survival), again suggesting allelopathic effects (Figure 38). Thus, these allelopathic effects should be further investigated, for instance in co-culture experiments with toxic species of Dinophysis versus competitor, prey (i.e. M. rubrum) or predator (e.g. copepods of the genus Acartia or Calanus). Such experiments could be performed using cell culture inserts for well plates or in co-culture chambers with adapted net mesh size allowing exchange of metabolites without direct contact (Bojadzija Savic et al., 2020).

(iii) The implications of Dinophysis toxins from an environmental point of view have been mainly studied for the sanitary, economic and social issues related to the accumulation of DSTs in mollusks and leading to DSP outbreaks and bans of shellfish harvesting, transport and marketing (Figure 38) and are not discussed here. However, much less attention has been paid to the direct effects of Dinophysis toxins on marine organisms (Chapter 1 see section “I-8 Effects of Dinophysis, OA- group toxins and PTXs on marine organisms”; Chapter 3-Part 1 and Chapter 3-Part 2) and are discussed in the next sections.

Effects of Dinophysis and its toxins on marine organisms

Accumulation of DSTs has been observed in several marine organisms from zooplankton and mollusks to crustaceans, fish and marine mammals (Chapter 1 section “I-7) Accumulation of Dinophysis, OA-group toxins and PTXs in marine organisms”) (Figure 39). Although direct effects have been suggested in turtles (Landsberg et al., 1999) and mammals (Fire et al., 2011) (Figure 39), most of the observations concerned the accumulation of DSTs and the effects of both DSTs and to a lesser extent Dinophysis cells towards filter-feeding bivalves and fish. In addition to accumulation, negative effects have been mainly observed towards fish (in vitro) and shellfish (both in vitro and in vivo) and, in this context, early life stages of these organisms are more sensitive to toxins from HAB species (Figure 39).

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Fish larvae  Mortality  Gill pathologies

Accumulation - Effects? Recruitment? Dinophysis Pediveliger depends of: spp.  mortality Grazers OA group- D-shaped defense toxins Abnormal Co-occurrence PTX2 development Competition Environmental  Fertilization concentrations Oocytes Prey capture  Mortality  ROS Route of exposure Hemocytes Accumulation -  ROS Effects?

Figure 39: Schematic representation of the effects of Dinophysis and their toxins on marine organisms: from in vivo experiments to the environment, with a focus on experiments performed in this thesis (light blue font) and their potential effects on fish and oyster recruitment, depending on environmental co-occurrence and concentrations, and routes of exposure (Chapter 3-Part 1, Chapter 3-Part 2 and general discussion). Dotted arrows represent literature observations. In vivo effects towards fish and filter-feeding bivalves from the literature are not presented here but in Chapter 1 section “I-8) Effects of Dinophysis, OA-group toxins and PTXs on marine organisms”. Drawings are not to scale; arrows size is not informative.

The (i) co-occurrence of Dinophysis and its toxins with other marine organisms, as well as the (ii) natural concentration of Dinophysis and the (iii) routes of exposure (cells or dissolved toxins), are important factors to consider when assessing all the potential environmental effects of Dinophysis (Figure 39).

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General discussion

(i) The analysis of databases in Chapter 3-Part 1, showed that the co-occurrence of oyster larvae (1st stage) with Dinophysis happens in several locations on the French Atlantic coast (Bay of Brest, Bay of Bourgneuf, Marennes-Oléron and Bay of Arcachon) from late spring to late summer. As described in Chapter 1 (section I-9-1) C. gigas), early life stages of oyster, from gametes to pediveliger larvae represent ca. 15 days of their life, and overlaps between stages may exist in the environment (see: https://wwz.ifremer.fr/velyger_eng/Acces-aux-Donnees/Rade-de-Brest/Stade- larvaire), thus increasing the probability of co-occurrence for weeks to months. Moreover, the geographical distribution of C. variegatus, i.e. East coast of South, Central and North America, and its habitat (i.e. benthic in coastal and estuarine waters) coincide with observations of Dinophysis blooms in these areas and especially in the US (Table 15 available online at: https://doi.org/10.17882/76433 and Chapter 1 section I-9-1) C. gigas). In addition, adults may also be exposed, in particular oysters, which are sessile organisms. Moreover, the well known co- occurrence with other mollusks of interest, such as mussels or clams, and fish in areas where Dinophysis blooms, e.g. estuarine, coastal waters (Reguera et al., 2014) stresses the need to assess the effects of Dinophysis and its toxins on these organisms as well.

(ii) Although Dinophysis is usually present at low concentrations in the water, exceptional blooms of 105-107 cells L-1 have been reported (Reguera et al., 2012) and were also observed in France (REPHY, 2019). According to Chapter 3-Part 1, a concentration of 0.5 to 5 cells mL-1 (i.e. 500 to 5000 cells L-1) of Dinophysis decreased by half the fertilization success of C. gigas, and such concentrations were often (ca. 10 times per year, according to the location; Chapter 2-Part 1) reached in the four described areas on the French Atlantic coast, from spring to fall. Exposure of C. variegatus larvae to natural concentrations of D. acuminata in the US (i.e. hundreds of cells per liter) (Wolny et al., 2020) did not cause mortality, but histological analyses are ongoing. Future studies should focus on long-term effect of chronic exposure to low concentrations of Dinophysis on adults when early life stages were exposed, as well as directly on adults and effects on the next generation.

(iii) According to the animal species and the life stages, the routes of exposure may be different. Indeed, Dinophysis cells may affect gametes, eggs, newly hatched eggs and first stage of larvae. In addition, Dinophysis cells and their contents (e.g. toxins, bioactive compounds) could negatively impact filter-feeding bivalves and planktivorous fish by exposure during ingestion. Moreover,

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General discussion native toxins and products of metabolization in bivalves could affect higher trophic levels (i.e. predators) by consumption of contaminated organisms. Finally, dissolved toxins passively released during the senescent growth phase at the end of a Dinophysis bloom may affect most marine organisms. These dissolved toxins may be a threat for these organisms, due to the fact that lipophilic OA has been shown to pass easily through the mollusks membranes (Blanco and Rossignoli, 2019; Daranas et al., 2007).

In this thesis, initial experiments focused on the exposure of gametes of C. gigas to D. sacculus and D. acuminata. This exposure resulted in a decline of fertilization success at low to high concentrations (from 0.5 to 500 cells mL-1) of Dinophysis, mainly due to the effect (i.e. increase of mortality) towards oocytes. Nevertheless, a cumulative effect was observed when both oocytes and spermatozoa were pre-exposed (Chapter 3-Part 1). The effects may come from the cells (i.e. mechanical damages by contact), known toxins or other bioactive compounds, but apparently not from the extracellular compartment (since supernatants did not show effects). While OA did not show any effect, PTX2 (from 5 to 50 nM) increased ROS production of oocytes and decreased the fertilization success, but did not explain all the observed effects (Chapter 3-Part 1) (Figure 39). The observed effects for oocytes of C. gigas were different when exposed to Dinophysis (increase in oocytes mortality) or PTX2 standard (increase in ROS production). Moreover, low concentrations of Dinophysis also produce negative effects towards oyster gametes (i.e. resulting in a decline of fertilization rate). This effect occurred, even though Dinophysis cells were intact (i.e. not detectable amount of toxins were released into extracellular compartment at low cellular concentration of Dinophysis, i.e. 0.5 or 5 cell(s) mL-1), suggesting different that different routes of exposure affect gametes in two different ways, both resulting in a decline in fertilization rate.

As far as we know, no other study has focused on the effects of Dinophysis and its toxins towards early life stages of oyster or other filter-feeding bivalves. However, in unpublished data, we observed developmental problems of D-shaped larvae in the presence of, or after exposure of gametes to D. sacculus. Furthermore, we exposed pediveliger larvae to PTX2 standard and observed 8, 43 and 100% of mortality after 24 h when exposed to 10, 20 and 50 nM concentrations of the toxin (Figure 39). Thus, our in vitro experiments suggested negative effects of Dinophysis and its toxins and towards several early life stages of the Pacific oyster (Chapter 1; I-9-1) C. gigas).

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General discussion

According to literature available, only a few experiments on the effects of Dinophysis on adult oysters (in vivo and in vitro) have been performed. Results from these experiments reported negative effects of Dinophysis (natural blooms or cultures), such as physiological modifications, including damage to digestive organs and mortality, as well as hemocytes damage (e.g. (Basti et al., 2015b; Mello et al., 2010); Chapter 1 section “I-8-2) Effect of filter-feeding bivalves exposure to Dinophysis or DSTs”). Hemocytes are contained within the hemolymph of bivalves, and initiate physiological response of bivalves exposed to environmental or biological stresses, thus modulating their immune responses (Hégaret et al., 2003b). Exposure of hemocytes from several bivalve species to OA showed cytotoxic, genotoxic and immunological effects, suggesting that OA could be detrimental for adult bivalves. Interestingly, we observed (unpubl. data) that, ROS production by C. gigas hemocytes increased when exposed to 50 nM of PTX2 (Figure 39), suggesting a potential threat posed by both, OA and PTX2, to adult oysters. These experiments showed negative effects of both Dinophysis and PTX2 towards several oyster life stages and type of cells.

Secondly, the effects of PTX2 and D. acuminata towards larvae of sheepshead minnow (C. variegatus) were determined. This toxin induced mortality after 96 h of exposure at intermediate to high concentrations (500 to 4000 nM) and damaged gill lamellae (Chapter 3-Part 2). Moreover, while D. acuminata did not cause fish mortality, damage similar to the effects after exposure to PTX2was observed in gills at high cellular concentration (5500 cells mL-1), (Chapter 3-Part 2) (Figure 39). Moreover, one experiment showed harmful effects (i.e. decreased growth rate) of D. acuminata, and genotoxicity was observed in adult fish exposed to OA and DTX1 (Rountos et al., 2019). Other in vitro studies showed negative effects of OA-group toxins on early life stages of fish, including embryos and larvae, such as damage in the digestive tract, genotoxicity, developmental disorders and mortalities (e.g. (Souid et al. (2018); Chapter 1 section “I-8-3) Effects of fish exposure to Dinophysis or DSTs”), suggesting toxic effects on several life stages

Thus, experiments on fish and oyster suggest that Dinophysis cells and one of its toxins alone, i.e. PTX2 have different mechanisms of toxicity of, which could be different towards fish or mollusks. Finally, active release of PTX2 (and OA-group toxins) has not been observed in Dinophysis and suggests that the negative effects of Dinophysis and its toxins should be faced as two different mechanisms.

238

General discussion

Overall, negative effects of DSTs and/or Dinophysis have been highlighted in this thesis towards almost all the oyster life cycle stages. These included from gametes and fertilization process, embryos and different larval development stages (with different feeding mechanisms, i.e. vellum or gills) and metamorphosis, to adults with physiological and pathological effects and especially on hemocytes. These observations raised the interest in studying the effects of HAB species, and Dinophysis, on all life stages of marine organisms and suggest that even if adults are not negatively affected by toxins (e.g. crab) (Torgersen et al., 2005), early life stages may be differentially affected. In addition, different responses may exist in different species and between life stages due to differences in: feeding behavior, protection (i.e. naked or surrounded by shell), motility (pelagic, benthic, sessile), habitat, sensitivity either to dissolved toxins or to Dinophysis cells.

Previous results from the literature and from this thesis (in vitro and in vivo tests on hemocytes) suggest that Dinophysis blooms may lead to recruitment failure of both fish and shellfish. Consequently, sanitary, economic, social and environmental problems may arise from these blooms, due to DSP events affecting human health and activities, e.g. bans of shellfish harvesting (professional and recreative), and effects on natural fish and shellfish populations. Further studies should focus on the effects on other marine organisms.

Environmental risks posed by PTX2-producers

Species of the genus Dinophysis, e.g. D. caudata, D. fortii, D. acuminata and D. sacculus (Table 15 available online at: https://doi.org/10.17882/76433) (Reguera et al., 2014) are the only known producers of pectenotoxins and according to the results of this thesis, dissolved PTX2 has negative effects towards gametes, and to several stages of larvae and hemocytes of C. gigas and larvae of C. variegatus. To date, such effects have not been studied in in vitro experiment on marine organisms. Pectenotoxins are not diarrheic toxins, a fact that may have limited the interest for studying the effects of PTX-producers. A possible hypothesis is PTX2 causes impairment of the actin filaments of gamete membranes or gill lamellae (i.e. pillar cells). The negative effect may come from the macrocyclic polyether structure of the PTX2, because OA, which is a linear polyether, did not induce toxic effects in oyster gametes (Chapter 3-Part 1) and PTX2-sa (opening of the lactone ring also results in a linear polyether) did not interfere with actin polymerization (Ares et al., 2007; Butler et al., 2012). However, goniodomin A, another macrocyclic polyether 239

General discussion structurally close to PTX2 which interferes with actin filament (Tainter et al., 2020), also negatively affects C. variegatus gill epithelia by an unknown mechanism (Chapter 3-Part 2). Cross experiments between PTX2 and goniodomin A and their producers, respectively Dinophysis and e.g. Alexandrium monilatum, could be performed in vitro on model organisms, such as cell lines (i.e. macrophages), fish or bivalves. Indeed, it could be interesting to study a potential additive effect of these toxins and/or the Dinophysis producers. Blooms of the latter usually co-occur with C. variegatus in different parts of the world (e.g. the Chesapeake Bay).

To date, the risk assessment of marine biotoxins for human consumers has been established by FAO (FAO/IOC/WHO, 2004; FAO/WHO, 2016). Pectenotoxins are not diarrheic and are not regulated in several countries (e.g. in USA) (Trainer et al., 2013). Nevertheless, cytotoxicity, genotoxicity and apoptotic effects , in addition to the effect on actin filaments towards mammals, fish, mollusks and cell lines (e.g. human neuroblastoma; Leira et al. (2002) have been observed; Chapter 1 section “I-6-2-3) Elucidation of action mode of PTXs”).

Most PTX analogues come from the PTX2, which undergoes acid-catalyzed hydrolysis in aqueous media to form its seco-acid (opening of the macrocycle) derivative or can be biotransformed by mollusks (Blanco, 2018). Recently, Krock et al. (2020) reported the presence at low background concentrations of PTX2 in SPATT in Antarctica, and mentioned that this toxin can persist for a long time in Antarctic waters, as well as in dissolved fractions in Galician waters, weeks after no detection of Dinophysis (Pizarro et al., 2009). Presence of PTX2 and OA-group toxins was also noticed in French and Nigerian waters in SPATT (Zendong et al., 2016a, 2016b). In addition, the metabolization of PTXs and the depuration time varied among filter-feeding bivalve species, from hours to weeks (see references in Blanco et al., 2018). These two aspects suggest that PTX2 and its hydrolyzed analogues (i.e. seco acid and ester products) may be found in both the environment and in filter-feeding bivalves. Unlike PTX2, the toxicity of its analogues, including to marine organisms, is unknown and yet they are exposed to (traces of) dissolved PTX2 (and at least PTX2sa) and PTX2-producers. Moreover, as mentioned in Chapter 1 (section “I-6-3) Effects of OA-group toxins and PTXs on humans”), to date, no toxicity of PTX2 or analogues towards humans have been reported.

Thus, a risk of toxicity to marine animals may exist if seco acid and ester products of PTXs have toxic effects or if there is an additive/synergistic effect of OA-group toxins, PTX native forms and

240

General discussion

PTX analogues towards a common target (e.g. actin) (Alarcan et al., 2019). Thus, further experiments should be carried out first for a better characterization of the toxic effects of PTXs and then determine the potential effects of these “non-toxic” products on marine organisms.

241

Conclusion

The PhD thesis first aimed to determine the effects of environmental factors on the physiology of the organismsm conforming the food chain of Dinophysis.

Dinophysis and its prey M. rubrum are obligate mixotrophs, sequester chloroplasts from prey and need light in order to grow. T. amphioxeia is at the basis of this food chain, thus its plastids are found in both M. rubrum and Dinophysis. This complex relationship may be important in order to explain and understand the bloom dynamics of the two predators.

The effects of temperature, irradiance and pH on the growth, the Fv/Fm and the pigment contents of T. amphioxeia, were assessed using a full factorial design. Then, two physiologically contrasting cultures of T. amphioxeia were obtained by photoacclimation in semi-continuous cultures at two light intensities, and used for feeding M. rubrum to determine the potential effects of prey quality and quantity on its growth, Fv/Fm and pigment content (Chapter 2-Part 1). Optimal growth of T. amphioxeia was attained at a temperature of 17.6 °C, an irradiance of 400 µmol photons m-2 s-1 and a pH of 7.6. This experiment highlighted the tolerance of T. amphioxeia to the tested abiotic factors and could suggest that the cryptophyte may not be affected by modifications of such factors in future ocean conditions. Moreover, the quantity of prey seemed an important parameter for M. rubrum growth, Fv/Fm and pigment contents, which may favor the ciliate in future climate conditions (Chapter 2-Part 1).

In future experiments, it could be interesting to test our models on several strains of T. amphioxeia to study the intraspecific variability of this species. In addition, to verify our hypothesis about M. rubrum and Dinophysis dynamics in future climate conditions (i.e. abundance of prey which could favor Dinophysis growth), integrative ecophysiological experiments may be implemented with competitors under potentially future (e.g. ocean temperature, pH or irradiance) conditions. This hypothesis could also be verified in situ, for instance in French Atlantic coastal waters, by monitoring T. amphioxeia, M. rubrum and Dinophysis at the species level, in order to determine a potential natural succession of these organisms.

242

Conclusion and perspectives

Species of the D. acuminata-complex and especially D. sacculus, have been shown to perform well in closed, semi-enclosed and estuarine areas, where rapid salinity variations are likely to happen. The effect of sudden salinity variations on the growth, biovolume, and toxin, osmolyte and other metabolite contents were tested on three French strains of D. sacculus (Chapter 2-Part 1). Osmoregulation took place in D. sacculus which seemed to be tolerant to the salinity stress, highlighted at least by the accumulation of osmolytes (i.e. proline, glycine betaine and DMSP) at the highest tested salinity compared to the lowest. In addition, growth, biovolume and metabolite contents in cells from the control and the hypo- hyperosmotic treatments were similar.

More generally, ecophysiological experiments should focus on both abiotic and biotic factors which are known to modulate toxin production in several species of Dinophysis. The aim of these studies would be to determine the potential toxicity of Dinophysis cells under future climate conditions, and to understand the role and discover the biosynthetic pathway of Dinophysis toxins.

The second part of the PhD thesis aimed to determine the effects of Dinophysis and its toxins on marine animals (oyster and fish).

Dinophysis toxins have been observed to accumulate in several marine organisms, including zooplankton, mollusks, fish and mammals. Moreover, toxicity of the OA-group toxins was observed during in vitro experiments on several model organisms including fish and mollusks. In addition, in vivo experiments showed toxic effects towards filter-feeding bivalves. However, few studies focused on the effect of PTX2, Dinophysis and especially PTX-producers.

The effect of Dinophysis, OA and PTX2 were determined in vitro on two marine animal models, the Pacific oyster (Crassostrea gigas) and the teleost fish sheepshead minnow (Cyprinodon variegatus). Toxic effects of D. sacculus and D. acuminata on oocytes (i.e. increases mortality) as well as PTX2 (i.e. increased ROS production) have been observed after 2 h of exposure and may have led to the decrease in fertilization rate, at low and natural field concentrations (i.e. 0.5 to 5 cell(s) mL-1) of Dinophysis and low concentration of PTX2 (5 nM). Moreover, fish exposed for 96 h to high concentration of D. acuminata (i.e. 5500 cells mL-1) showed pathology in gills, and exposure from medium to high concentration of PTX2 (i.e. 500 to 4000 nM) led to, apparently, similar pathology in gill and mortality of larvae. These experiments highlighted that the toxic effects of Dinophysis cells and PTX2 standards may be different. Effects on different targets (i.e.

243

Conclusion and perspectives bivalve gametes, fish or bivalve larvae) were also different and may be due to other bioactive compounds.

To further characterize the toxic effects of Dinophysis towards marine organisms and the possible environmental/ecological implications, it could be interesting to perform experiments on several life stages of fish and bivalve species and to include different species of Dinophysis, producing different toxins, as well as non-toxin producers. Moreover, such implications may be further investigated by studying environmental co-occurrence, concentrations and routes of exposure of Dinophysis towards marine organisms. Overall, the effects of PTXs towards recruitment of marine organisms is of particular interest and should be more intensively studied.

244

Scientific publications and communications

Published:

1-) Gaillard, S., Charrier, A., Malo, F., Carpentier, L., Bougaran, G., Hégaret, H., Réveillon, D., Hess, P., Séchet, V., 2020a. Combined effects of temperature, irradiance, and pH on Teleaulax amphioxeia (Cryptophyceae) physiology and feeding ratio for its predator Mesodinium rubrum (Ciliophora). Journal of Phycology. 56, 775-783. https://doi.org/10.1111/jpy.12977. Open access: https://hal.archives-ouvertes.fr/hal-02880016

2-) Gaillard, S., Le Goïc, N., Malo, F., Boulais, M., Fabioux, C., Zaccagnini, L., Carpentier, L., Sibat, M., Réveillon, D., Séchet, V., Hess, P., Hégaret, H., 2020b. Cultures of Dinophysis sacculus, D. acuminata, and pectenotoxin 2 affect gametes and fertilization success of the Pacific oyster, Crassostrea gigas. Environ. Pollut. 265, 114840. https://doi.org/10.1016/j.envpol.2020.114840. Open access: https://hal.archives-ouvertes.fr/hal-02880050/

In revision:

3-) Gaillard, S., Réveillon, D., Danthu, C., Hervé, F., Sibat, M., Carpentier, L., Hégaret, H., Séchet, V., Hess, P., 2020c. Effect of a 24 h salinity stress on the growth, biovolume, toxins, osmolytes and metabolite profiles on three strains of the Dinophysis acuminata-complex (“D. sacculus”). In revision in Harmful Algae (2020-09-01)

4-) Séchet, V., Sibat, M., Billien, G., Carpentier, L., Rovillon, G.A., Raimbault, V., Malo, F., Gaillard, S., Perrière-Rumebe, M., Hess, P., Chomérat, N., 2021. Characterization of toxin- producing strains of Dinophysis spp. (Dinophyceae) from French coasts, with a particular focus on the D. acuminata-complex. In revision in Harmful Algae (2020-08-01).

In preparation:

5-) Gaillard, S., Mason, P.L., Ayache, N., Sanderson, M., Smith, J. L., Giddings, S., McCarron, P., Séchet, V., Hégaret, H., Réveillon, D., Hess, P., Vogelbein, W. K., 2021. Mortality and histopathological features of sheepshead minnow (Cyprinodon variegatus) larvae exposed to Dinophysis acuminata and pectenotoxin 2 during a 96 h bioassay. In preparation for submission.

245

Oral presentation:

Culture of prey organisms of Dinophysis sp. helps explain why its occurrence along the French Atlantic coast does not shown trends over a 20-years period. Gaillard, S., Malo, F., Carpentier, L., Charrier, A., Bougaran, G., Irisson, J.O., Sourisseau, M., Réveillon, D., Séchet, V., Hégaret, H., Hess, P., August 2019. 7th European Phycological Congress (EPC), Zagreb, Croatia.

Poster:

Influence of temperature, irradiance and pH on the physiology of Teleaulax amphioxeia. Gaillard, S., Charrier, A., Malo, F., Carpentier, L., Bougaran, G., Hégaret, H., Réveillon, D., Hess, P., Séchet, V., October 2018. 18th International Conference on Harmful Algae (ICHA), Nantes, France.

246

Appendices

I (µmol Culture T photons L: D Species Strain Origin media (°C) m-2 s-1) pH Salinity (h) Reference CR- MAL0 Teleaulax sp. 1 Korea f/2 n.a. n.a. n.a. n.a. n.a. (Park et al., 2006) (Smith and T. amphioxeia n.a. Denmark f/2 15 100 n.a. 30 16: 8 Hansen, 2007) Teleaulax sp. n.a. Korea n.a. n.a. n.a. n.a. n.a. n.a. (Kim et al., 2008) TA- 100 to T. amphioxeia INO01 Japan f/2 18 150 n.a. 30 12: 12 (Nagai et al., 2008) AB36 100 to (Nishitani et al., T. amphioxeia 4287 Japan f/2 18 150 n.a. 30 12: 12 2008a) AB36 (Nishitani et al., T. amphioxeia 4287 Japan f/2 16 100 n.a. 30 12: 12 2008b) CCMP (Hackett et al., G. cryophila 2564 Antarctica f/2 3 100 n.a. n.a. 14: 10 2009) Chry- (Kamiyama and Teleaulax sp. S0606 Japan SWM3 15 15 n.a. n.a. 12: 12 Suzuki, 2009) K- (Riisgaard and T. amphioxeia 0434 Denmark f/2 21 100 n.a. 32 14: 10 Hansen, 2009) K- (Garcia-Cuetos et T. amphioxeia 0434 Denmark f/2 15 100 n.a. 32 14: 10 al., 2010) CCMP G. cryophila 2564 Antarctica f/2 6 50 n.a. n.a. 14: 10 (Tong et al., 2010) AB36 modified 100 to (Kamiyama et al., T. amphioxeia 4287 Japan f/2 18 150 n.a. 30 12: 12 2010) CCMP G. cryophila 2564 Antarctica f/2 4 65 n.a. n.a. 14: 10 (Fux et al., 2011) TA- T. amphioxeia INO01 Japan f/2 18 100 n.a. n.a. 12: 12 (Nagai et al., 2011) CCMP G. cryophila 2564 Antarctica f/2-Si 4 n.a. n.a. n.a. n.a. (Tong et al., 2011) CR- MAL0 Teleaulax sp. 1 Korea f/2-Si 20 10 n.a. 30 14: 10 (Kim et al., 2012b) K- (Nielsen et al., T. amphioxeia 0434 Denmark f/2 15 130 8 32 16: 8 2012) 15 - AND- 19 - (Rodríguez et al., T. amphioxeia A0710 Spain L1/20 25 100 n.a. 32 12: 12 2012) TA- T. amphioxeia INO01 Japan f/2 18 100 n.a. n.a. 12: 12 (Nagai et al., 2013) 247

Chry- Teleaulax sp. S0606 Japan f/2 18 100 n.a. n.a. 12: 12 K- (Nielsen et al., T. amphioxeia 0434 Denmark f/2 15 130 8 32 16: 8 2013) AND- T. amphioxeia A0710 Spain L1/20 n.a. 150 n.a. 32 12: 12 (Raho et al., 2013) AND- T. amphioxeia A0710 Spain f/2 16 100 n.a. 30 12: 12 (Rial et al., 2013) AND- T. amphioxeia A0710 Spain L1/20 15 150 n.a. 32 12: 12 (Riobó et al., 2013) AB36 T. amphioxeia 4287 Japan f/2 22 120 n.a. 30 12: 12 (Mafra et al., 2014) AB36 (Basti et al., T. amphioxeia 4287 Japan f/2 18 100-150 n.a. 30 12: 12 2015c) (Hattenrath- K- Lehmann et al., T. amphioxeia 0434 Denmark f/2 18 70 n.a. 25 14: 10 2015) CCMP (Tong et al., G. cryophila 2564 Antarctica f/2 4 50 n.a. n.a. 14: 10 2015b) AB36 modified (Tong et al., T. amphioxeia 4287 Japan f/6 15 65 n.a. n.a. 14: 10 2015a) K- (Hansen et al., T. amphioxeia 1837 Denmark f/2 15 100 8 32 16: 8 2016) AB36 T. amphioxeia 4287 Japan f/2 20 100 n.a. 30 12: 12 (Mafra et al., 2016) K- (Nielsen et al., T. amphioxeia 0434 Denmark f/2 15 130 8 30 16: 8 2016) K- (Ojamäe et al., T. amphioxeia 1837 Denmark f/2 15 100 n.a. 35 14: 10 2016) TA- (Papiol et al., T. amphioxeia INO01 Japan f/2 18 70-100 n.a. n.a. 12: 12 2016) AND- T. amphioxeia A0710 Spain f/6 15 ≈50 n.a. n.a. 14: 10 (Gao et al., 2017) K- (Rusterholz et al., T. amphioxeia 0434 Denmark f/2 15 n.a. 8.2 30 16: 8 2017) AB36 T. amphioxeia 4287 Japan f/2 18 100 n.a. n.a. 12: 12 (Basti et al., 2018) AND- T. amphioxeia A0710 Spain f/6 15 ≈50 n.a. n.a. 14: 10 (Gao et al., 2018) AND- (García-Portela et T. amphioxeia A0710 Spain L1/20 15 270 8 32 12: 12 al., 2018b) AB36 modified T. amphioxeia 4287 Japan f/2 15 65 n.a. 32 14: 10 (Jiang et al., 2018) AB36 T. amphioxeia 4287 Japan f/2 15 50 n.a. n.a. n.a. (Smith et al., 2018) AND- T. amphioxeia A0710 Spain f/2 19 50 n.a. n.a. n.a. CCMP G. cryophila 2564 Antarctica f/2 4 50 n.a. n.a. n.a. USGC (=GC G. cryophila EP02) USA f/2 15 50 n.a. n.a. n.a. AB36 T. amphioxeia 4287 Japan f/6 15 20 n.a. 30 14: 10 (Gao et al., 2019)

248

Plagioselmis CR10 (García-Portela et prolonga EHU Spain K 19 160-180 n.a. 32 16: 8 al., 2019) AB36 (Hongo et al., T. amphioxeia 4287 Japan f/2 18 100 n.a. n.a. 12: 12 2019) AND- T. amphioxeia A0710 Spain f/6 15 54 n.a. n.a. 14: 10 (Jia et al., 2019) K- (Rountos et al., T. amphioxeia 0434 Denmark f/2 18 100 n.a. 25 12: 12 2019) CR- MAL0 T. amphioxeia 4 Korea f/2 20 50 n.a. n.a. n.a. (Lee et al., 2019) CR- MAL0 T. amphioxeia 5 Korea f/2 20 50 n.a. n.a. n.a. CR- MAL0 T. amphioxeia 6 Korea f/2 20 50 n.a. n.a. n.a. CR- MAL0 T. amphioxeia 7 Korea f/2 20 50 n.a. n.a. n.a. CR- MAL0 T. amphioxeia 8-1 Korea f/2 20 50 n.a. n.a. n.a. CR- MAL0 T. amphioxeia 9-2 Korea f/2 20 50 n.a. n.a. n.a. CR- LOHA T. amphioxeia BE01 Korea f/2 20 140 n.a. 30 14: 10 (Park et al., 2019) AND- (Gaillard et al., T. amphioxeia A0710 Spain L1 17.8 100 8 35 12: 12 2020a) AND- (Gaillard et al., T. amphioxeia A0710 Spain L1 17.8 100 8 35 12: 12 2020b) AND- (Gaillard et al., T. amphioxeia A0710 Spain L1 17.8 100 8 35 12: 12 2020c) Plagioselmis CR10 (García-Portela et prolonga EHU Spain L1/20 15 200 8 32 12: 12 al., 2020) K- (Nielsen et al., R. salina 0294 Denmark f/2 15 130 8 32 16: 8 2020) K- T. amphioxeia 0434 Denmark f/2 15 130 8 32 16: 8 AB36 (Wolny et al., T. amphioxeia 4287 Japan f/6 15 65 n.a. n.a. 14: 10 2020) AND- (Séchet et al., T. amphioxeia A0710 Spain L1 17 90 8.2 35 12: 12 2020) K- (Fiorendino et al., T. amphioxeia 0434 Denmark L1 20 100 n.a. 30 14: 10 2020) GoMT A USA L1 20 100 n.a. 30 14: 10

249

I (µmol L: Culture T photons D predator: Nutrition Species Strain Origin media (°C) m-2 s-1) pH Salinity (h) prey frequency Reference M. MR- (Park et al., rubrum MAL01 Korea f/2 n.a. n.a. n.a. n.a. n.a. n.a. n.a. 2006) (Smith and M. 12: Hansen, rubrum n.a. Denmark f/2 15 100 n.a. 30 12 n.a. n.a. 2007) M. MR- (Kim et al., rubrum MAL01 Korea n.a. n.a. n.a. n.a. n.a. n.a. n.a. once/week 2008) M. 100 to 12: (Nagai et rubrum MR-INO01 Japan f/2 18 150 n.a. 30 12 1: 7.5 n.a. al., 2008) (Nishitani M. 100 to 12: et al., rubrum AB364286 Japan f/2 18 150 n.a. 30 12 1: 1 n.a. 2008a) (Nishitani M. 12: et al., rubrum AB364286 Japan f/2 16 100 n.a. 30 12 1: 25 n.a. 2008b) M. 14: (Hackett et rubrum CCMP2563 Antarctica f/2 3 100 n.a. n.a. 10 n.a. n.a. al., 2009) Sea water + EDTA + (Kamiyama M. Meso- trace 12: and Suzuki, rubrum S0606A4 Japan metals 15 15 n.a. n.a. 12 n.a. n.a. 2009) (Riisgaard and M. 14: Hansen, rubrum Mr-DK2007 Denmark f/20 21 100 n.a. 16 10 n.a. n.a. 2009) (Garcia- M. 14: Cuetos et rubrum Mr-DK2007 Denmark f/20 20 100 n.a. 16 10 n.a. n.a. al., 2010) M. 14: (Tong et al., rubrum CCMP2563 Antarctica f/2 6 50 n.a. n.a. 10 3: 1 n.a. 2010) M. modified 100 to 12: (Kamiyama rubrum AB364286 Japan f/2 18 150 n.a. 30 12 85: 1 n.a. et al., 2010) M. 14: (Fux et al., rubrum CCMP2563 Antarctica f/2 4 65 n.a. n.a. 10 n.a. n.a. 2011) M. 12: (Nagai et rubrum MR-INO01 Japan f/2 18 100 n.a. n.a. 12 1: 1 n.a. al., 2011) M. (Tong et al., rubrum CCMP2563 Antarctica f/2 4 n.a. n.a. n.a. n.a. 1: 10 n.a. 2011) M. MR- 14: (Kim et al., rubrum MAL01 Korea f/2 20 10 n.a. 30 10 n.a. n.a. 2012b) M. MBL- 16: (Nielsen et rubrum DK2009 Denmark f/2 15 130 8 32 8 1: 10 n.a. al., 2012) 15 - M. AND- 19 - 12: (Rodríguez rubrum A0711 Spain L1/20 25 100 n.a. 32 12 n.a. n.a. et al., 2012)

250

M. 12: (Nagai et rubrum MR-INO01 Japan f/2 18 100 n.a. n.a. 12 n.a. n.a. al., 2013) M. Meso- 12: rubrum S0606A4 Japan f/2 18 100 n.a. n.a. 12 n.a. n.a. M. MBL- 16: (Nielsen et rubrum DK2009 Denmark f/2 15 130 8 32 8 1: 10 twice/week al., 2013) M. AND- 12: (Raho et al., rubrum A0711 Spain L1/20 n.a. 150 n.a. 32 12 n.a. n.a. 2013) M. AND- 12: (Rial et al., rubrum A0711 Spain f/2 16 100 n.a. 30 12 3: 1 n.a. 2013) M. AND- 12: (Riobó et rubrum A0711 Spain L1/20 15 150 n.a. 32 12 n.a. n.a. al., 2013) M. 12: once to (Mafra et rubrum n.a. Brazil f/2 22 120 n.a. 30 12 1: 10 twice/week al., 2014) M. (Basti et al., rubrum AB364286 Japan n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. 2015b) M. 100- 12: (Basti et al., rubrum AB364286 Japan f/2 18 150 n.a. 30 12 n.a. once/week 2015c) (Hattenrath- M. MBL- 14: Lehmann et rubrum DK2009 Denmark f/2 18 70 n.a. 25 10 1: 10 n.a. al., 2015) M. 14: (Tong et al., rubrum CCMP2563 Antarctica f/2 4 50 n.a. n.a. 10 1: 10 n.a. 2015b) M. modified 14: every two (Tong et al., rubrum AB364286 Japan f/6 15 65 n.a. n.a. 10 1: 14 weeks 2015a) M. MBL- 16: (Hansen et rubrum DK2009 Denmark f/2 15 100 8 32 8 1: 10 twice/week al., 2016) M. 12: (Mafra et rubrum AB364286 Japan f/2 20 100 n.a. 30 12 1: 10 n.a. al., 2016) M. MBL- 16: (Nielsen et rubrum DK2009 Denmark f/2 15 130 8 30 8 1: 10 twice/week al., 2016) M. MBL- 14: (Ojamäe et rubrum DK2009 Denmark f/2 15 100 n.a. 35 10 1: 3 n.a. al., 2016) M. New 12: (Papiol et rubrum n.a. Zeland f/2 18 70-100 n.a. n.a. 12 n.a. n.a. al., 2016) M. 14: (Gao et al., rubrum AB364286 Japan f/6 15 ≈50 n.a. n.a. 10 1: 1 once/week 2017) M. AND- 14: rubrum A0711 Spain f/6 15 ≈50 n.a. n.a. 10 1: 1 n.a. every week M. MBL- 16: or two (Rusterholz rubrum DK2009 Denmark f/2 15 n.a. 8.2 30 8 1: 5 weeks et al., 2017) M. 12: (Basti et al., rubrum AB364286 Japan f/2 18 100 n.a. n.a. 12 1: 1 n.a. 2018) M. 14: (Gao et al., rubrum AB364286 Japan f/6 15 ≈50 n.a. n.a. 10 1: 1 once/week 2019) (García- M. AND- 12: Portela et rubrum A0711 Spain L1/20 15 270 8 32 12 n.a. n.a. al., 2018b) M. 14: (Jiang et al., rubrum AB364286 Japan f/6 15 65 n.a. 32 10 1: 1 n.a. 2018) M. (Smith et rubrum CCMP2563 Antarctica f/2 4 50 n.a. n.a. n.a. 1: 10 n.a. al., 2018) M. rubrum AB364286 Japan f/12 15 50 n.a. n.a. n.a. n.a. n.a.

251

M. AND- rubrum A0711 Spain f/12 19 50 n.a. n.a. n.a. n.a. n.a. M. 14: (Gao et al., rubrum AB364286 Japan f/6 15 20 n.a. 30 10 n.a. n.a. 2019) (García- M. AND- 160- 16: Portela et rubrum A0711 Spain K 19 180 n.a. 32 8 n.a. n.a. al., 2019) M. 12: (Hongo et rubrum AB364286 Japan f/2 18 100 n.a. 30 12 14: 1 once/week al., 2019) M. AND- 14: (Jia et al., rubrum A0711 Spain f/6 15 54 n.a. n.a. 10 1: 1 once/week 2019) M. MBL- 12: (Rountos et rubrum DK2009 Denmark f/2 18 100 n.a. 25 12 1: 5 n.a. al., 2019) M. MR- 14: (Park et al., rubrum LOHABE01 Korea f/2 20 140 n.a. 30 10 n.a. n.a. 2019) M. AND- 12: (Gaillard et rubrum A0711 Spain Seawater 17.8 100 8 35 12 1: 1 thrice/week al., 2020a) M. MBL- 12: (Gaillard et rubrum DK2009 Denmark Seawater 17.8 100 8 35 12 1: 1 thrice/week al., 2020b) M. MBL- 12: (Gaillard et rubrum DK2009 Denmark Seawater 17.8 100 8 35 12 1: 1 thrice/week al., 2020c) (García- M. AND- 12: Portela et rubrum A0711 Spain L1/20 15 200 8 32 12 n.a. n.a. al., 2020) M. MBL- 16: (Nielsen et rubrum DK2009 Denmark f/2 15 130 8 32 8 1: 10 twice/week al., 2020) M. 14: (Wolny et rubrum AB364286 Japan f/6 15 65 n.a. n.a. 10 n.a. n.a. al., 2020) M. MBL- 12: (Séchet et rubrum DK2009 Denmark L1 17 90 8.2 35 12 n.a. twice/week al., 2020) M. MBL- 14: (Fiorendino rubrum DK2009 Denmark L1 20 100 n.a. 30 10 1: 10 n.a. et al., 2020)

252

I (µmol predat Culture T photons Salinit L: D or: Nutrition Species Strain Origin media (°C) m-2 s-1) pH y (h) prey frequency Reference D. (Park et al., acuminata Korea f/2 20 60 n.a. 30 24: 0 n.a. n.a. 2006) D. DA- (Kim et al., acuminata MAL01 Korea f/2 20 60 n.a. 30 24: 0 n.a. twice/week 2008) D. DAEP0 14: (Hackett et acuminata 1 USA f/2 3 100 n.a. n.a. 10 1: 7 n.a. al., 2009) Sea water + EDTA + (Kamiyama D. trace 12: and Suzuki, acuminata n.a. Japan metals 15 15 n.a. n.a. 12 n.a. n.a. 2009) (Riisgaard D. Da- Denmar 14: and Hansen, acuminata DK2007 k f/20 21 100 n.a. 22 10 n.a. n.a. 2009) (Garcia- D. Da- Denmar 14: Cuetos et acuminata DK2007 k f/20 18 100 n.a. 22 10 n.a. n.a. al., 2010) D. DAEP0 14: (Tong et al., acuminata 1 USA f/2 6 300 n.a. n.a. 10 1: 20 n.a. 2010) D. modifie 100 to 12: (Kamiyama acuminata Japan Japan d f/2 18 150 n.a. 30 12 1: 16 n.a. et al., 2010) D. DAEP0 14: (Fux et al., acuminata 1 USA f/2 6 65 n.a. n.a. 10 1: 5 n.a. 2011) D. DAMV 14: acuminata 01 USA f/2 6 65 n.a. n.a. 10 1: 5 n.a. D. DABOF 14: acuminata 01 Canada f/2 6 65 n.a. n.a. 10 1: 5 n.a. D. acuminata DARE0 14: (or ovum) 1 Chile f/2 6 65 n.a. n.a. 10 1: 5 n.a. D. acuminata DAPA0 14: (or ovum) 1 USA f/2 6 65 n.a. n.a. 10 1: 5 n.a. D. DA0810 (Nagai et acuminata HAR03 Japan n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. al., 2011) D. DAEP0 (Tong et al., acuminata 1 USA f/2 6 n.a. n.a. n.a. n.a. 1: 4 n.a. 2011) DC- D. LOHAB 14: (Kim et al., acuminata E01 Korea f/2 20 10 n.a. 30 10 n.a. twice/week 2012b) D. (Nielsen et acuminata n.a. n.a. f/2 15 130 8 32 16: 8 10: 1 n.a. al., 2012) (Hattenrath- D. Lehmann et acuminata n.a. USA n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. al., 2013)

253

D. 12: (Mafra et acuminata n.a. Brazil f/2 22 120 n.a. 30 12 1: 3 n.a. al., 2014) (Hattenrath- D. 14: Lehmann et acuminata n.a. USA f/2 18 70 n.a. 25 10 1: 10 n.a. al., 2015) D. DAEP0 14: (Tong et al., acuminata 1 USA f/2 6 65 n.a. n.a. 10 1: 20 n.a. 2015a) D. DAMV 14: acuminata 01 USA f/2 6 65 n.a. n.a. 10 1: 20 n.a. D. DABOF 14: acuminata 01 Canada f/2 6 65 n.a. n.a. 10 1: 20 n.a. D. DAPA0 14: acuminata 1 USA f/2 6 65 n.a. n.a. 10 1: 20 n.a. D. DAMV modifie 14: (Tong et al., acuminata 01 USA d f/6 15 65 n.a. n.a. 10 1: 6 once/week 2015b) D. FR1010 Denmar 14: (Ojamäe et acuminata 09 k f/2 15 100 n.a. 35 10 1: 10 n.a. al., 2016) 1: 10 D. DAYS0 14: then 1: (Gao et al., acuminata 1 China f/6 15 100 n.a. n.a. 10 5 n.a. 2017) D. Da- Denmar 1: 5- (Rusterholz acuminata DK2007 k f/2 15 n.a. 8.2 30 16: 8 10 twice/week et al., 2017) D. DA0810 12: (Basti et al., acuminata HAR03 Japan f/2 18 100 n.a. n.a. 12 n.a. n.a. 2018) D. DAYS0 14: (Gao et al., acuminata 1 China f/6 15 ≈50 n.a. n.a. 10 n.a. n.a. 2019) (García- D. VGO13 12: Portela et acuminata 49 Spain L1/20 15 270 8 32 12 n.a. n.a. al., 2018b) D. DAMV Seawate 14: (Jiang et al., acuminata 01 USA r 15 65 n.a. 32 10 n.a. once/week 2018) D. DAEP0 Seawate (Smith et acuminata 1 USA r 6 65 n.a. n.a. n.a. 1: 10 n.a. al., 2018) D. DAEP0 14: (Gao et al., acuminata 1 USA f/6 15 20 n.a. 30 10 n.a. n.a. 2019) (García- D. VGO13 Portela et acuminata 49 Spain K 19 160-180 n.a. 32 16: 8 n.a. n.a. al., 2019) D. DAYS0 14: (Jia et al., acuminata 1 China f/6 15 54 n.a. n.a. 10 1: 5 twice/week 2019) D. 12: (Rountos et acuminata n.a. USA f/2 18 100 n.a. 25 12 1: 2 n.a. al., 2019) D. (Diaz et al., acuminata n.a. Spain n.a. n.a. n.a. n.a. n.a. n.a. n.a. n.a. 2019) D. acuminata 54 14: (Park et al., complex* strains Korea f/2 20 140 n.a. 30 10 n.a. n.a. 2019) IFR- D. DAU- L1/20 + 12: 1: 1 to (Gaillard et acuminata 02Ar France K/2 17.8 100 8 35 12 1: 10 thrice/week al., 2020b) (García- D. VGO13 12: Portela et acuminata 49 Spain L1/20 15 200 8 32 12 1: 10 once/week al., 2020) D. DADE0 14: (Wolny et acuminata 1 USA f/6 15 65 n.a. n.a. 10 1: 10 n.a. al., 2020)

254

IFR- D. DAU- L1/20 + 12: (Séchet et acuminata 03An France K/2 17 90 8.2 35 12 1: 1 n.a. al., 2020) IFR- D. DAU- L1/20 + 12: acuminata 01Es France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DAU- L1/20 + 12: acuminata 01Ca France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DAU- L1/20 + 12: acuminata 01Ke France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DAU- L1/20 + 12: acuminata 01Du France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DAU- L1/20 + 12: acuminata 02Bm France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DAU- L1/20 + 12: acuminata 01Bo France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DAU- L1/20 + 12: acuminata 02Ar France K/2 17 90 8.2 35 12 1: 1 n.a. D. DAVA0 14: (Fiorendino acuminata 1 USA L1 20 100 22 10 1: 10 n.a. et al., 2020)

I (µmol L: Culture T photons D predator Nutrition Species Strain Origin media (°C) m-2 s-1) pH Salinity (h) : prey frequency Reference 14: (Jaén et al., D. acuta n.a. n.a. f/2 15 20 n.a. n.a. 10 n.a. n.a. 2009) DAN North A- Atlantic 16: (Nielsen et D. acuta 2010 Ocean f/2 15 130 8 32 8 1: 10 twice/week al., 2013) VGO1 n.a. 14: (Rial et al., D. acuta 065 Spain L1/20 n.a. 70-200 n.a. 32 10 1: 20 n.a. 2013) DAN North A- Atlantic 16: (Hansen et D. acuta 2010 Ocean f/2 15 100 8 32 8 1: 10 twice/week al., 2016) DAN North A- Atlantic 16: (Nielsen et D. acuta 2010 Ocean f/2 15 130 8 30 8 1: 10 twice/week al., 2016) DAN North A- Atlantic 14: (Ojamäe et D. acuta 2010 Ocean f/2 15 100 n.a. 35 10 1: 10 n.a. al., 2016) New 12: (Papiol et D. acuta n.a. Zeland f/2 18 70-100 n.a. n.a. 12 n.a. n.a. al., 2016) 255

DAN North A- Atlantic 16: (Rusterholz D. acuta 2010 Ocean f/2 15 n.a. 8.2 30 8 1: 5-10 twice/week et al., 2017) (García- VGO1 12: Portela et D. acuta 065 Spain L1/20 15 270 8 32 12 n.a. n.a. al., 2018) (García- VGO1 16: Portela et D. acuta 065 Spain K 19 160-180 n.a. 32 8 n.a. n.a. al., 2019) n.a (Diaz et al., D. acuta n.a. Spain n.a. n.a. n.a. n.a. n.a. . n.a. n.a. 2019) (García- VGO1 12: Portela et D. acuta 065 Spain L1/20 15 200 8 32 12 1: 10 once/week al., 2020) DAN North A- Atlantic 16: (Nielsen et D. acuta 2010 Ocean f/2 15 130 8 32 8 1: 10 twice/week al., 2020) IFR- DAC- L1/20 + 12: (Séchet et D. acuta 03Ke France K/2 17 90 8.2 35 12 1: 1 n.a. al., 2020) IFR- DAC- L1/20 + 12: D. acuta 02Lc France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- DAC- L1/20 + 12: D. acuta 03Lc France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- DAC- L1/20 + 12: D. acuta 04Lc France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- DAC- L1/20 + 12: D. acuta 01Ar France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- DAC- L1/20 + 12: D. acuta 01Po France K/2 17 90 8.2 35 12 1: 1 n.a.

I (µmol L: Culture T photons D predator Nutrition Species Strain Origin media (°C) m-2 s-1) pH Salinity (h) : prey frequency Reference D. VGO1 12: (Raho et al., sacculus 132 Spain L1/20 n.a. 150 n.a. 32 12 n.a. n.a. 2013) D. VGO1 12: (Riobo et sacculus 132 Spain L1/20 15 150 n.a. 32 12 n.a. n.a. al., 2013) IFR- D. DSA- Seawate 12: 1: 1 to (Gaillard et sacculus 01Lt France r 17.8 100 8 35 12 1: 10 thrice/week al., 2020b)

256

IFR- D. DSA- Seawate 12: 1: 1 to 1: (Gaillard et sacculus 01Lo France r 17.8 100 8 35 12 10 thrice/week al., 2020c) IFR- D. DSA- Seawate 12: 1: 1 to 1: sacculus 01Me France r 17.8 100 8 35 12 10 thrice/week IFR- D. DSA- Seawate 12: 1: 1 to 1: sacculus 02Th France r 17.8 100 8 35 12 10 thrice/week IFR- D. DSA- L1/20 + 12: (Séchet et sacculus 01Co France K/2 17 90 8.2 35 12 1: 1 n.a. al., 2020) IFR- D. DSA- L1/20 + 12: sacculus 01Lo France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DSA- L1/20 + 12: sacculus 01Me France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DSA- L1/20 + 12: sacculus 01Lt France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DSA- L1/20 + 12: sacculus 01Th France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DSA- L1/20 + 12: sacculus 02Th France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DSA- L1/20 + 12: sacculus 03Th France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DSA- L1/20 + 12: sacculus 01Vp France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DSA- L1/20 + 12: sacculus 01Po France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DSA- L1/20 + 12: sacculus 01Ma France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DSA- L1/20 + 12: sacculus 01Lj France K/2 17 90 8.2 35 12 1: 1 n.a.

I (µmol L: Culture T photons D predator Nutrition Species Strains Origin media (°C) m-2 s-1) pH Salinity (h) : prey frequency Reference

257

(Nishitani D. 100 to 12: et al., caudata n.a. n.a. f/2 18 150 n.a. 30 12 1: 3 n.a. 2008a) D. VGO1 12: (Raho et al., caudata 064 Spain L1/20 n.a. 150 n.a. 32 12 n.a. 2013) D. VGO1 12: (Rial et al., caudata 064 Spain L1/20 n.a. 70-200 n.a. 32 12 1: 20 n.a. 2013) D. 12: (Mafra et caudata n.a. Brazil f/2 22 120 n.a. 30 12 1: 3 n.a. al., 2014) D. 21- n.a (Basti et al., caudata n.a. Japan n.a. 23 n.a. n.a. n.a. . n.a. n.a. 2015a) D. 21- 12: (Basti et al., caudata n.a. Japan f/2 23 100-150 n.a. 30 12 n.a. n.a. 2015b) IFR- D. DCA- L1/20 + 12: (Séchet et caudata 01Ke France K/2 17 90 8.2 35 12 1: 1 n.a. al., 2020) IFR- D. DCA- L1/20 + 12: caudata 03Ke France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DCA- L1/20 + 12: caudata 02Lo France K/2 17 90 8.2 35 12 1: 1 n.a. IFR- D. DCA- L1/20 + 12: caudata 04Tr France K/2 17 90 8.2 35 12 1: 1 n.a. 100 to 12: (Nagai et D. fortii n.a. f/2 18 150 n.a. 30 12 1: 10 n.a. al., 2008) DF070 5TTR 12: (Nagai et D. fortii 24 Japan f/2 18 100 n.a. n.a. n.a. 12 n.a. al., 2011) 12: (Hongo et D. fortii n.a. Japan f/2 18 100 n.a. 30 12 1: 10 thrice/week al. 2019) D. (Nishitani infundib 12: et al., ulus n.a. Japan f/2 16 100 n.a. 30 12 1: 20 n.a. 2008b) D. acumina ta (or DARE 14: (Fux et al., ovum) 01 Chile f/2 6 65 n.a. n.a. 10 1: 5 n.a. 2011) D. acumina ta (or DAPA 14: (Fux et al., ovum) 01 USA f/2 6 65 n.a. 10 1: 5 n.a. 2011) DAC- 12: (Mafra et D. ovum BR2 Brazil f/2 20 100 n.a. 30 12 1: 3 n.a. al., 2016) 14 DoSS3 : (Fiorendino D. ovum 195 USA L1 20 100 n.a. 22 10 1 : 10 n.a. et al., 2020) 15 - D. VGO1 19 - 12: (Rodriguez tripos 062 Spain L1/20 25 100 n.a. 32 12 n.a. n.a. et al., 2012) D. DT080 12: (Nagai et tripos 4S01 Japan f/2 18 100 n.a. n.a. 12 n.a. n.a. al., 2013) D. VGO1 12: (Raho et al., tripos 062 Spain L1/20 n.a. 150 n.a. 32 12 n.a. n.a. 2013)

258

D. VGO1 12: (Rial et al., tripos 062 Spain L1/20 n.a. 70-200 n.a. 32 12 1: 20 n.a. 2013) IFR- D. DTR- L1/20 + 12: (Séchet et tripos 01Ar France K/2 17 90 8.2 35 12 1: 1 n.a. al., 2020)

259

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Titre : Études écophysiologiques sur Dinophysis et sa chaîne alimentaire, et effets in vitro du dinoflagellé et de ses toxines sur les premiers stades de vie de deux modèles d'animaux marins (l’huître et le poisson)

Mots clés : Changement global, Crassostrea gigas, Cyprinodon variegatus, Dinophysis, Efflorescences, Mesodinium, Pecténotoxines, Teleaulax, Toxines diarrhéiques

Résumé : Les efflorescences algales nuisibles de rapides de la salinité sur D. sacculus a été étudié. Ce Dinophysis sont responsables de l'intoxication diarrhéique facteur a été rapporté comme pouvant potentiellement par les mollusques chez les consommateurs humains de déterminer la distribution de Dinophysis. D. sacculus mollusques contaminés après bioaccumulation de leurs présente une grande tolérance aux stress de salinité, au toxines. Dinophysis est un organisme mixotrophe qui moins en partie due à la synthèse d'osmolytes, ce qui séquestre les chloroplastes d'une proie unique, explique ainsi sa distribution environnementale dans des Mesodinium rubrum, pour effectuer sa propre habitats côtiers soumis à des fluctuations rapides de photosynthèse, lui-même mixotrophe se nourrissant d’un salinité. Bien que la toxicité et les modes d'action des cryptophyte, Teleaulax amphioxeia. La relation entre ces toxines de Dinophysis soient connus sur les mammifères trois organismes est fondamentale afin de comprendre la terrestres, les effets directs de Dinophysis et de ses distribution et la dynamique des efflorescences de toxines sur les organismes marins sont encore mal Dinophysis. Dans un premier temps, l'effet de la étudiés. Il a donc été montré expérimentalement que température, de l'irradiance et du pH sur la physiologie de Dinophysis et la pecténotoxine 2 (PTX2) altèrent les T. amphioxeia a été déterminé en utilisant un plan factoriel ovocytes d'huîtres Crassostrea gigas, affectant ainsi le complet, avant d'évaluer l'effet d'états physiologiques taux de fécondation, et causent des dommages aux contrastés après l'acclimatation à deux intensités branchies des poissons Cyprinodon variegatus et la lumineuses de T. amphioxeia et l’effet de la quantité de mortalité des larves. En conclusion, ce travail met en proies pour M. rubrum. La plasticité physiologique de T. évidence l'importance d'une meilleure compréhension de amphioxeia a été mise en évidence et suggère une la physiologie de la chaîne trophique de Dinophysis, et tolérance importante aux conditions futures de l'océan, suggère un effet négatif du dinoflagellé et de la PTX2 sur ainsi que l'importance de l’abondance des proies pour la les ressources aquatiques, jusqu’alors inconnu croissance de son prédateur. Ensuite, l'effet des variations

Title: Ecophysiological studies on Dinophysis and its food chain, and in vitro effects of the dinoflagellate and its toxins on early life stages of two models of marine animals (oyster and fish)

Keywords: Crassostrea gigas, Cyprinodon variegatus, Diarrheic toxins, Dinophysis, Global change, Harmful algal blooms, Mesodinium, Pectenotoxins, Teleaulax

Abstract: Harmful algal blooms of Dinophysis are of its predator. Subsequently, the effect of rapid salinity responsible for diarrheic shellfish poisoning in human variations on D. sacculus were assessed, as such consumers of contaminated mollusks after bioaccumulation variations may be a potential driver of its distribution. D. of their toxins. Dinophysis is a mixotrophic organism, which sacculus present a large tolerance to salinity stresses, at sequesters chloroplasts from a unique prey Mesodinium least partly due to the synthesis of osmolytes, explaining rubrum to perform its own photosynthesis, itself its coastal disitrubution in areas subjected to rapid salinity mixotrophic and feeds a cryptophyte, Teleaulax fluctuations. Although toxicity and modes of actions of amphioxeia. The relationship between these three Dinophysis toxins are known on terrestrial mammals, the organisms is fundamental to understand the distribution direct effects of Dinophysis and its toxins on marine and bloom dynamics of Dinophysis. In a first study, the organisms are still poorly studied. Thus, it was effects of temperature, irradiance and pH on physiological experimentally showed that Dinophysis and the aspects of T. amphioxeia were determined using a full pectenotoxin 2 (PTX2) impaired oyster oocytes, then factorial design before assessing the effect of contrasting affecting the fertilization rate, and causing damage to fish physiological states after acclimation to two light intensities gills and mortality of larvae. Overall, this study highlights of T. amphioxeia and the effect of quantity of the prey for the importance of a better understanding of the M. rubrum. The ecophysiological plasticity of T. amphioxeia physiology of the trophic chain of Dinophysis, and suggests a high tolerance to future ocean conditions as suggests negative effects of the dinoflagellate on aquatic well as the importance of prey abondance for growth ressources, which was unknown so far.