Diversity and genetics of Australasian dinoflagellates

Diversity and genetics of Australasian dinoflagellates, including Gambierdiscus spp. the causative agent of Ciguatera Fish Poisoning

by

Gurjeet Singh Kohli BSc, MTM

A thesis submitted in partial fulfilment of the requirements for the degree of Doctor of Philosophy (Ph.D.)

The School of Biotechnology and Biomolecular Sciences Faculty of Science The University of New South Wales Sydney, Australia

2013

ii

Supervisors

Prof. Brett A. Neilan School of biotechnology and Biomolecular Sciences The University of New South Wales

Assoc. Prof. Shauna A. Murray Plant functional Biology & Climate Change Cluster University of Technology, Sydney

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Acknowledgements This PhD thesis was only possible because of the help and support of so many people, but first and foremost I would like to thank my supervisors, Prof. Brett A. Neilan and Assoc. Prof. Shauna A. Murray for the opportunity to work with them. I learnt many valuable lessons from your insightful advice. This work was only possible because of the endless support, guidance and constructive critique both of you provided over all these years. I was honoured to visit the Cawthron Institute in Nelson, New Zealand several times during my PhD and would like to thank Dr. Lesley Rhodes for providing the cultures and much needed guidance during the fish feeding trials. I would like to thank Dr. Tim Harwood and Andy Selwood (Cawthron Institute) who performed the LC-MS analysis, for chapters three, four and five. I am grateful to Dr. Uwe John (AWI, Germany) for performing the transcriptomic sequencing for chapter five and guidance throughout the data analysis. I would like to thank Prof. Gustaaf Hallegraeff (UTAS) for doing the electron microscopy for chapter three and partially funding the trip to Merimbula. I also thank Dr. Rosa Figueroa (LU, Sweden) for performing the flow cytometry analysis for chapter five. I am grateful to Lauren Mayer and Dr. Angela Capper (JCU) for collecting some of the samples for chapter three and Prof. Manfred Lenzen (USyd) for providing some samples in chapter two. I thank Dr. Chris Bolch (UTAS) and Dr. Michelle Moffit (UWS) for providing much needed guidance during the annual reviews. I also thank Dr. Mark Brown (UNSW) and Dr. Mona Hoppenrath (Senkenberg Institute, Germany) for advice in chapter two, Dr. Hazel Farrell (UTS) for writing a part of chapter one, Dr. Nandan Deshpande and Dr. Federico Leuro (UNSW) for bioinformatics help during genome assembly in chapter five, and Prof. Tertulohi Matainaho (UPNG, Papua New Guinea) for organising the sampling trip in Papua New Guinea. The research underlying this thesis was funded directly by grants from the Australian Research Council (CIs Neilan, Moffit, Bolch, grant number DP0880264, genome sequencing in chapter five and support during first three years; CIs Murray, John, grant number DP120103199, support during last year), Australian Biological Resources Study (CIs Murray, Hoppenrath, Brown, Moore, grant number RFL210-34, sampling and analysis for chapter two and three) the Linnean Society of New South Wales (Kohli, partial pyrosequencing costs for Chapter three), Ministry of Science and Innovation, New Zealand (CIs, Rhodes, Olsen, fish feeding trials and mass spectrometric analysis in chapter four and five) and Alfred Wegener Institute (CI John, for transcriptomic sequencing in chapter five). I was awarded the following scholarships and thank the funding bodies: Australian Postgraduate Award, Post Graduate Research Support Scheme UNSW, School of BABS travel fund, ISSHA student travel scholarship and GRS student travel scholarship.

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I would also like to thank the support staff at the following institutes where this research was conducted: School of BABS, University of New South Wales; C3 Institute, University of Technology, Sydney; Sydney Institute of Marine Sciences and Ramaciotti centre for gene functional analysis, Sydney. I am grateful to my dear fiends Julia, Kristin, Alper, Jason, Adrian, Martina and all the past and present members of BGGM and Murray laboratory for encouraging and supporting me more than I could have ever hope for. Last but not least, I would like to thank my parents and my lovely wife Razveen, to whom I dedicate this thesis. I could have never done it without you; thank you so much for all the emotional support throughout my life and also the financial help whenever the going got tough or a holiday was needed. You’re simply the best!

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List of Publications and Presentations Kalaitzis, J. A., Chau, R., Kohli, G. S., Murray, S. A. & Neilan, B. A. 2010. Biosynthesis of toxic naturally-occurring seafood contaminants. Toxicon, 56, 244-258.

Kohli, G. S., Neilan, B. A., Brown, M. V., Hoppenrath, M. & Murray, S. A. 2013. Cob gene pyrosequencing enables characterization of benthic dinoflagellate diversity and biogeography. Environmental Microbiology. In press, available online DOI: 10.1111/1462-2920.12275

Kohli, G. S., Murray, S. A., Neilan, B. A. 2010. Diversity of Benthic Dinoflagellates from Papua New Guinea. Poster presentation. 14th International Conference on Harmful Algae, Hersonissos-Crete, Greece.

Kohli, G. S., Rhodes, L. L., Harwood, T., Selwood, A., Jerrett, A., Murray, S. A., Neilan, B. A. 2012. A feeding study to probe the uptake of maitotoxin by snapper (Pragus auratus). Oral presentation. 15th International Conference on Harmful Algae, Changwon-Gyeongnam, Korea.

Kohli, G. S., Murray, S. A., Rhodes, L. L., Harwood, T., Hallegraeff, G. M., Neilan, B. A. 2013. Occurrence of the Ciguatera Fish Poisoning causing dinoflagellate Gambierdiscus in temperate waters of New South Wales, Australia. Oral presentation. 9th International Conference on Molluscan Shellfish Safety, Sydney, Australia.

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Abstract Marine microbial protists from benthic habitats include species of the genera Gambierdiscus, which can produce polyketide toxins with ecosystem-wide impacts and impacts on human health. Knowledge of the presence, diversity, effects and distribution of benthic dinoflagellates and their toxins is vital both for basic ecological research and for managing potential impacts on human health. However, at present such information is limited. This study designed novel pyrosequencing-based tools that utilised 18S rRNA and mitochondrial cytochrome b barcoding marker genes. Using these tools, the largest set of dinoflagellate cytochrome b gene sequences to date was developed, and dinoflagellate species richness within marine benthic samples from five tropical marine benthic sites were determined. These tools were applied to samples from a temperate marine benthic site from southern NSW and the species Gambierdiscus carpenteri was found to be highly abundant throughout the region. This was unexpected, given that it is primarily considered a tropical species. The significance of finding this species is discussed in the context of long term monitoring data and its potential for toxin production. Species of Gambierdiscus produce two major toxin groups, ciguatoxins (CTXs) and maitotoxins (MTXs), that can accumulate via marine food chains in seafood and ultimately cause ciguatera fish poisoning (CFP) in humans. While the role of CTXs is well established in CFP, little is known about the role of MTXs. This work showed experimentally that MTXs can accumulate in liver and muscle tissues of carnivorous fish (Pagrus auratus). The levels of MTX detected in fish tissues showed for the first time that MTX may accumulate in fish at levels harmful for human consumption. In order to investigate the genetic basis of toxin biosynthesis in species of Gambierdiscus, two comprehensive transcriptomic libraries for the species Gambierdiscus australes and Gambierdiscus belizeanus, which produce MTXs were sequenced and assembled. Data analysis discovered three hundred and seven genes that formed eight clades within a phylogeny of dinoflagellate polyketide synthesis genes, and may be involved in MTX biosynthesis. This is the first step towards identifying prospective genes that may be involved in MTX biosynthesis, and provides a basis for the future development of molecular tools to assess the risk of CFP.

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Table of Contents Chapter 1

General Introduction ............................................................................................. 1

1.1

The genus Gambierdiscus ................................................................................................... 1

1.2

Morphology and phylogenetics ........................................................................................... 2

1.3

Geographic distribution and abundance .............................................................................. 7

1.3.1

The Pacific and Indian Ocean regions .............................................................................. 7

1.3.2

The Atlantic Ocean region ................................................................................................ 9

1.4

CTXs and MTXs ............................................................................................................... 11

1.5

Toxicity of different species of Gambierdiscus................................................................. 16

1.6

Detection of CTXs and MTXs in seafood ......................................................................... 17

1.7

Aims of the study............................................................................................................... 26

Chapter 2

Cob gene pyrosequencing enables characterisation of benthic dinoflagellate

diversity and biogeography ...................................................................................................... 27 2.1

Summary............................................................................................................................ 27

2.2

Introduction ....................................................................................................................... 27

2.3

Materials and methods ....................................................................................................... 29

2.3.1

Sample collection ............................................................................................................ 29

2.3.2

Culturing ......................................................................................................................... 30

2.3.3

DNA extraction ............................................................................................................... 31

2.3.4

PCR, construction and screening of dinoflagellate specific cob clone libraries ............. 32

2.3.5

Tag-Encoded FLX 454-Pyrosequencing......................................................................... 32

2.3.6

cTEFP sequence analysis ................................................................................................ 33

2.3.7

rTEFP sequence analysis ................................................................................................ 34

2.3.8

Phylogenetic analysis ...................................................................................................... 34

2.4

Results ............................................................................................................................... 34

2.4.1

Diversity of dinoflagellates via PCR, cob gene cloning and sequencing ....................... 34

2.4.2

Diversity of dinoflagellates via cTEFP ........................................................................... 36

2.4.3

Diversity of dinoflagellates via rTEFP ........................................................................... 44

2.5

Discussion.......................................................................................................................... 46

2.5.1

Comparison of cTEFP, rTEFP and cob cloning based sequencing methods .................. 46

2.5.2

Identification of novel dinoflagellate diversity ............................................................... 47

Chapter 3

Abundance of the potentially ciguatera fish poisoning causing dinoflagellate

Gambierdiscus carpenteri in temperate waters of New South Wales, Australia................... 50 3.1

Summary............................................................................................................................ 50 X

3.2

Introduction ....................................................................................................................... 50

3.3

Materials and methods ...................................................................................................... 52

3.3.1

Sample collection ........................................................................................................... 52

3.3.2

DNA extraction, Tag-Encoded FLX 454-Pyrosequencing and sequence analysis ........ 55

3.3.3

DNA extraction and PCR analysis for species identification ......................................... 55

3.3.4

Phylogenetic analysis ..................................................................................................... 56

3.3.5

Gambierdiscus population density and analysis............................................................. 56

3.3.6

Electron microscopy ....................................................................................................... 56

3.3.7

Toxin analysis via LC-MS and mouse bioassay............................................................. 57

3.4

Results ............................................................................................................................... 57

3.4.1

rTEFP analysis and LSU phylogenetic analysis ............................................................. 57

3.4.2

Microscopy and scanning electron microscope analysis ................................................ 61

3.4.3

Gambierdiscus distribution and cell density estimates .................................................. 64

3.4.4

Toxin analysis ................................................................................................................ 65

3.5

Discussion ......................................................................................................................... 66

3.5.1

Dinoflagellate diversity as determined by rTEFP analysis ............................................ 66

3.5.2

Gambierdiscus species in NSW ..................................................................................... 67

Chapter 4 auratus)

A feeding study to probe the uptake of Maitotoxin by snapper (Pagrus ........................................................................................................................... 70

4.1

Summary ........................................................................................................................... 70

4.2

Introduction ....................................................................................................................... 70

4.3

Materials and methods ...................................................................................................... 72

4.3.1

Culturing and growth conditions .................................................................................... 72

4.3.2

DNA extraction, PCR amplification and gene sequencing ............................................ 72

4.3.3

Fish feeding trials ........................................................................................................... 73

4.3.4

Sample extraction and LC-MS analysis ......................................................................... 75

4.4

Results and discussion ...................................................................................................... 77

4.4.1

Dinoflagellate culture and toxin production ................................................................... 77

4.4.2

Fish feeding trials and LC-MS analysis ......................................................................... 77

4.4.3

PCR analysis .................................................................................................................. 83

Chapter 5

Transcriptome and genome analysis of two species of Gambierdiscus ........... 85

5.1

Summary ........................................................................................................................... 85

5.2

Introduction ....................................................................................................................... 85

5.3

Materials and methods ...................................................................................................... 90

5.3.1

Gambierdiscus cell culture ............................................................................................. 90 XI

5.3.2

DNA and RNA extraction............................................................................................... 90

5.3.3

Toxin analysis via LC-MS and mouse bioassay ............................................................. 90

5.3.4

Genome size estimation via flow cytometry ................................................................... 91

5.3.5

Transcriptome analysis ................................................................................................... 91

5.3.6

Genome survey of Gambierdiscus australes .................................................................. 92

5.4

Results ............................................................................................................................... 92

5.4.1

Toxin profile of G. australes and G. belizeanus ............................................................. 92

5.4.2

Genome and transcriptome size ...................................................................................... 94

5.4.3

Transcriptome analysis ................................................................................................... 96

5.4.3.1

Sequences involved in polyketide synthesis in G. australes and G. belizeanus .......... 98

5.4.3.2

Other sequences putatively involved in polyether biosynthesis ................................ 103

5.4.3.3

Sequences of potential bacterial origin ...................................................................... 103

5.4.4 5.5

Genome survey of G. australes .................................................................................... 105 Discussion........................................................................................................................ 106

5.5.1

Genome size and gene content estimation .................................................................... 106

5.5.2

Comprehensiveness of the gene catalogues .................................................................. 107

5.5.3

Genes potentially involved in polyketide biosynthesis ................................................. 108

Chapter 6

Final discussion and future outlook ................................................................. 112

References

.......................................................................................................................... 118

Appendix A

Additional tables and figures ......................................................................... 135

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List of Figures Figure 1.1: Scanning electron micrograph of Gambierdiscus australes ....................................... 2 Figure 1.2: Comparative line drawings of the epitheca for 11 Gambierdiscus species ................ 6 Figure 1.3: Structure of Ciguatoxins (CTX) and Maitototoxin-1. .............................................. 13 Figure 2.1: Map showing location of different sampling sites.................................................... 30 Figure 2.2: Phylogenetic analysis using maximum likelihood of cob sequences obtained from PCR based cob gene cloning and sequencing ..................................................................... 35 Figure 2.3: Phylogenetic analysis using maximum likelihood of cob sequences identified as Gonyaulacales during cTEFP analysis ................................................................................ 41 Figure 2.4: Phylogenetic analysis using maximum likelihood of cob sequences identified as Gymnodiniales during cTEFP analysis ............................................................................... 42 Figure 2.5: Phylogenetic analysis using maximum likelihood of cob sequences identified as Suessiales and Prorocentrales during cTEFP analysis. ....................................................... 43 Figure 2.6: Phylogenetic analysis using maximum likelihood of SSU sequences obtained from rTEFP analysis. ................................................................................................................... 45 Figure 2.7: Venn diagram showing the distribution of unique OTUs obtained during cTEFP analysis................................................................................................................................ 48 Figure 3.1: Map of the Eastern coastline of Australia ................................................................ 54 Figure 3.2: Map of Merimbula Inlet ........................................................................................... 54 Figure 3.3: Phylogenetic analysis using maximum likelihood of 28S rRNA gene sequences obtained from LSU analysis ................................................................................................ 59 Figure 3.4: Phylogenetic analysis using maximum likelihood of 18S rRNA gene sequences obtained from rTEFP analysis............................................................................................. 61 Figure 3.5: Scanning electron microscopy micrographs of Gambierdiscus carpenteri from Merimbula Lake Inlet.......................................................................................................... 62 Figure 3.6: Scanning electron microscopy micrographs of Gambierdiscus carpenteri from Merimbula Lake Inlet.......................................................................................................... 63 Figure 3.7: Gambierdiscus cell densities (cells g-1 wet weight algae) from Merimbula in May 2013..................................................................................................................................... 64 Figure 3.8: Salinity (A) and temperature (B) records of Merimbula Inlet Lake from 2004 to 2013..................................................................................................................................... 65 Figure 4.1: Flow chart showing the mode of transmission of Ciguatoxins and Maitotoxins in Ciguatera fish poisoning ..................................................................................................... 71 Figure 4.2: Pictorial representations of mullet (A. forsteri) being injected with concentrated G.australes culture. ............................................................................................................. 74

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Figure 4.3: Structure of Maitotoxin-1 and its oxidative cleavage product obtained during oncolumn periodate oxidation and quantified via LC-MS/MS analysis, method 1................. 76 Figure 4.4: Bar graph representing the amount of Maitotoxin-1 detected in different tissue types during the first fish feeding trial. ......................................................................................... 78 Figure 4.5: Extracted ion chromatograms for Maitotoxin-1 from various extracts following SPE cleanup and on-column oxidation (Method 1) during the first fish feeding trial................. 79 Figure 4.6: Total ion chromatograms for intact Maitotoxin-1 molecule from various extracts following SPE cleanup (Method 2) during the second fish feeding trial............................. 80 Figure 4.7: Extracted ion chromatograms from the intact toxin analysis of various extracts obtained during second fish feeding trial............................................................................. 83 Figure 5.1: Proposed mechanism for Maitotoxin-1 production................................................... 89 Figure 5.2: Extracted ion chromatograms from the intact toxin analysis of various extracts obtained during Maitotoxin LC-MS. ................................................................................... 93 Figure 5.3: Correlation between cell dimensions (length of dorso-ventral axis of a cell) and the amount of nuclear DNA in 27 different species of dinoflagellates ..................................... 95 Figure 5.4: Number of protein coding and total genes (y-axis) in the genome of various dinoflagellates including that of G. australes and G. belizeanus (x-axis) ........................... 95 Figure 5.5: Number of annotated sequences (y-axis) involved in various metabolic processes present in the gene catalogues of G. australes and G. belizeanus (x-axis).......................... 97 Figure 5.6: Phylogenetic analysis of H2A histone proteins ........................................................ 98 Figure 5.7: Phylogenetic analysis of Type I and Type II ketoacyl synthase (KS) domains ...... 100 Figure 5.8: Phylogenetic analysis of Type I and Type II ketoreductase (KR) domains ............ 102 Figure 5.9: Number of sequences giving top blast hit to various unicellular eukaryotes, plants, fungi and prokaryotes. ....................................................................................................... 105

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List of Tables Table 1.1: Taxonomic and genetic identifications of different species of Gambierdiscus. .......... 3 Table 1.2: Geographic distribution and toxicity of different Gambierdiscus species. .................. 8 Table 1.3: Different congeners of CTXs and MTXs................................................................... 12 Table 1.4: Different congeners of Ciguatoxins detected by various assays in seafood and other animals. ............................................................................................................................... 19 Table 2.1: List of sampling sites, the macroalgae the samples were collected from, their water temperature at the time of sample collection and the type of analysis done with each sample. ................................................................................................................................ 31 Table 2.2: Details of diversity of dinoflagellates at each site based on PCR based cob gene cloning and sequencing. ...................................................................................................... 36 Table 2.4: Data obtained during cTEFP analysis. ....................................................................... 38 Table 2.4: Sequences identified during cTEFP analysis. ............................................................ 39 Table 2.6: Data obtained during rTEFP analysis. ....................................................................... 44 Table 3.1: Sampling sites, the macroalgae the samples were collected from, the water temperature at the time of sample collection and the type of analysis done with each sample. ................................................................................................................................ 53 Table 3.2: Data obtained from rTEFP analysis. .......................................................................... 58 Table 4.1: Data obtained during the first fish feeding trial. ........................................................ 78 Table 4.2: Data obtained during the second fish feeding trial. ................................................... 81 Table 5.1: Snapshot of the bioinformatics analysis conducted on the gene catalogue of G. australes and G. belizeanus. ............................................................................................... 97 Table 5.2: Sequences found in G. australes and G. belizeanus gene catalogue putatively involved in polyketide biosynthesis.. .................................................................................. 99 Table 5.3 A list of enzymes that are potentially involved in polyether biosynthesis. ............... 103 Table 5.4: A list of type I PKS domains, sulfotransferases and expoxide hydrolases considered as bacterial contaminants. ................................................................................................. 104 Table 5.5: Description of the type I PKS clusters found during genome analysis of G. australes. .......................................................................................................................................... 106

XV

List of Abbreviations ACP

acyl carrier protein

ACPS

acyl carrier protein synthase

AFsD

acute reference dose

AT

acyltransferase

AZA

azaspiracid

AZP

azaspiracid shellfish poisoning

BA

bioassay

BLAST

basic local alignment search tool

bp

base pairs

BSBA

brine shrimp bioassay

BTX

brevitoxin

cDNA

complementary deoxyribonucleic acid

CEGMA

core eukaryotic genes mapping approach

CFP

ciguatera fish poisoning

cob

cytochrome b gene

cox

cytochrome c oxidase gene

cTEFP

cob tag-encoded FLX 454-pyrosequencing

CTX

ciguatoxin

DA

domoic acid

DH

dehydratase

DNA

deoxyribonucleic acid

DLBA

diptera larvae bioassay

DSP

diarrheic shellfish poisoning

DTX

dinophysistoxin

ER

enoyl reductase

ELISA

enzyme-linked immunosorbent assay

EFSA

european food safety authority

HAB

harmful algal bloom

HELA

human erythrocytes lysis assay

HPLC

high performance liquid chromatography

KR

ketoreductase

KS

ketosynthase

LC

liquid chromatography

LG

Le and Gascuel

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LM

light microscopy

LSU

large ribosomal subunit gene

MA

membrane bioassay

m/z

mass-to-charge ratio

MBA

mouse bioassay

MGBA

mongoose bioassay

MQBA

mosquito bioassay

MS

mass spectrometry

MT

methyltransferase

MTX

maitotoxin

NCBI

National Centre for Biotechnology Information

NCMA

neuro-2a cell binding assay

NMR

nuclear magnetic resonance

NRPS

nonribosomal peptide synthetase

NSP

neurotoxic shellfish poisoning

OA

Okadaic acid

OTU

operational taxonomic unit

PCR

polymerase chain reaction

PKS

polyketide synthase

PST

paralytic shellfish toxins

PSP

paralytic shellfish poisoning

PTX

pectenotoxin

RBA

receptor binding assay

RIA

radioimmunoassay

RLB

radio ligand binding

RNA

ribonucleic acid

rRNA

ribosomal ribonucleic acid

rTEFP

ribosomal tag-encoded FLX 454-pyrosequencing

SEM

scanning electron microscopy

SEIA

stick enzyme immunoassay

SPIA

solid phase immunoassay

SSU

small ribosomal subunit gene

STX

saxitoxin

sp.

species (singular)

spp.

species (plural)

TCA

tricarboxylic acid

TE

thioesterase XVII

TLC

thin layer chromatography

UPLC

ultra performance liquid chromatography

YTX

yessotoxin

XVIII

Chapter 1

Chapter 1 General Introduction 1.1

The genus Gambierdiscus Recent advances in population and species genetics for phytoplankton have revealed

immense biodiversity at different taxonomic levels (Simon et al., 2009). Vast numbers of species remain to be documented, aided by rapidly developing molecular methods (Murray et al., 2012a, Murray et al., 2012b). To date, there are only ca. 160 described benthic (sand dwelling and epiphytic) dinoflagellates (Taylor et al., 2008). The first report (Yasumoto et al., 1977) of the involvement of a benthic dinoflagellate in ciguatera fish poisoning (CFP) brought increased attention to this group. This species was described as Gambierdiscus based on the type species Gambierdiscus toxicus, from samples collected in the Gambier Islands, French Polynesia (Adachi and Fukuyo, 1979). Species of the genus Gambierdiscus have now been recognised as the main producers of ciguatoxins (CTXs) and maitotoxins (MTXs) (Chinain et al., 1997, Holmes, 1998, Chinain et al., 1999a, Chinain et al., 2010, Rhodes et al., 2010, Fraga et al., 2011, Holland et al., 2013) (Figure 1.1). CFP is the most common nonbacterial illnesses associated with fish consumption (Friedman et al., 2008), affecting between 50,000 and 500,000 people per year (Fleming et al., 1998). The ingestion of herbivorous and carnivorous fish that have orally accumulated effective levels of CTXs, and possible MTXs, can cause CFP in humans (Bagnis et al., 1979, Gillespie, 1987, Sims, 1987). Recent reviews have illustrated the global increase in the frequency and intensity of harmful algal events (Glibert et al., 2005, Hallegraeff, 2010). Despite being significantly underreported, CFP occurrence worldwide is increasing, with reports of a 60% increase in CFP in the Pacific Islands over the last decade (Skinner et al., 2011). Figure 1 describes the major milestones achieved in furthering our understanding of CFP and Gambierdiscus. Once considered a monotypic taxon, new species of Gambieriscus are being discovered every year with evidence showing that each species might have its own characteristic toxin profile (Chinain et al., 2010, Fraga et al., 2011, Holland et al., 2013). As in the case of other dinoflagellate genera such as Alexandrium, Karenia, the production or not of certain toxin groups appears to generally vary at the species level, rather than being consistent within the genus. For this reason, species of harmful algal bloom (HAB) forming taxa are monitored, acting as early warning systems for shellfish and seafood safety. This review highlights the significant advances in the study of Gambierdiscus. This chapter provides a summary of the morphology and phylogenetics of species of Gambierdiscus, their toxicology, distribution, chemistry and methods for the detection of CTXs and MTXs in seafood. The review further

1

Chapter 1

outlines the major gaps in our current understanding of Gambierdiscus and outlines goals for future research in this field.

Figure 1.1: Scanning electron micrograph of Gambierdiscus australes (Faust, 2007).

1.2

Morphology and phylogenetics When originally described (Adachi and Fukuyo, 1979), Gambierdiscus was considered as

a monotypic taxon, however, variability in the morphology, differences in ribosomal RNA (rRNA) genes, toxicity and physiological characteristics (Adachi and Fukuyo, 1979, Bomber et al., 1988, Bomber et al., 1989a, Bomber et al., 1989b, Holmes et al., 1990, Holmes et al., 1991, Morton et al., 1993, Chinain et al., 1997, Richlen et al., 2008) led to the description of new species. Currently, 11 species of Gambierdiscus have been described, based on their distinct morphological and molecular genetic characteristics (Table 1.1).

2

Chapter 1

Table 1.1: Taxonomic and genetic identifications of different species of Gambierdiscus. Species

Cell size (depth-widthlength) (μm)

G. yasumotoi (Holmes)

(56.8 ± 5.6) – (51.7 ± 5.6) – (62.4 ± 4.3)

G. ruetzleri (Vandersea, Litaker, Faust, Kibler, Holland et Tester) G. belizeanus (Faust)

G. pacificus (Chinain et Faust)

G. excentricus (Fraga)

G. australes (Faust et Chinain)

G. caribaeus (Vandersea, Litaker, Faust, Kibler, Holland et Tester)

G. carpenteri (Vandersea, Litaker, Faust, Kibler, Holland et Tester) G. toxicus

Morphological Characteristics

Globular species Larger cell size- cell width larger than 42 μm

Genetics (accession numbers in Genbank)

SSU: EF202846-52 D1-D3 LSU: EF202965-68 D8-D10 LSU: EF498087-89 (45.5 ± 3.3) – Smaller size- cell SSU: EF202853-60 (37.5 ± 3.0) – width less than 42 D1-D3 LSU: (51.6 ± 4.9) μm EF202962-64 D8-D10 LSU: EF498081-85 Anterio-posteriorly compressed species (61.7 ± 3.1) – Narrow 1p plateSSU: EF202876-77 (60.0 ± 4.5) – heavily aerolated D1-D3 LSU: (48.1 ± 4.2) cell surfaceEF202940-43 different 2’ plate D8-D10 LSU: symmetry and size EF498028-34 (58.5 ± 3.9) – Narrow 1p plateSSU: EF202861-65 (53.6 ± 4.1) – smooth cell surface- D1-D3 LSU: (40.4 ± 3.6) 2’ hatch shaped EF202944-47 D8-D10 LSU: EF498012-13, EF498015-16 (97.8 ± 8) – Narrow1p plateD1-D3 LSU: (83 ± 10) – smooth cell surface- HQ877874, (37 ± 3) 2’ rectangular JF303063, shaped- cell size JF303065-71 bigger than G. D8-D10 LSU: australes (1.5 times JF303073-76 wider and deeper) (72.5 ± 3.8) – Narrow 1p plateSSU: EF202891-96 (63.4 ± 5.0) – smooth cell surface- D1-D3 LSU: (38.7 ± 3.8) 2’ rectangular EF202969-72 shaped- smaller than D8-D10 LSU: G. excentricus EF498072-74 (82.2 ± 7.6) – Broad 1p plate- 2’ SSU: EF202914-28 (81.9 ± 7.9) – Rectangular shaped- D1-D3 LSU: (60 ± 6.2) Symmetric 4’’ EF202929-37, EF202983, EF202985 D8-D10 LSU: EF498045-71 (81.7 ± 6.4) – Broad 1p plate- 2’ SSU: EF202908-13 (74.8 ± 5.9) – Rectangular shaped- D1-D3 LSU: (50.2 ± 6.1) Asymmetric 4’’ EF202938-39, EF202984 D8-D10 LSU: EF498038-44 (93 ± 5.5) – Broad 1p plate-2’ SSU: EF202878-90

References

(Holmes, 1998) (Litaker et al., 2009) (Litaker et al., 2009)

(Faust, 1995) (Litaker et al., 2009) (Chinain et al., 1999a) (Litaker et al., 2009)

(Fraga et al., 2011)

(Chinain et al., 1999a) (Litaker et al., 2009) (Litaker et al., 2009)

(Litaker et al., 2009)

(Adachi and 3

Chapter 1

Species

Cell size (depth-widthlength) (μm)

(Adachi et Fukuyo) Chinain, Faust, Holmes, Litaker et Tester)

(83 ± 2.3) – (54 ± 1.5)

G. polynesiensis (Chinain et Faust)

(66.3 ± 3) – (60.5 ± 5.9) – (44.3 ± 5.1)

Morphological Characteristics

Hatchet shapedDorsal end 1p pointed

Genetics (accession numbers in Genbank) D1-D3 LSU: EF202951-61 D8-D10 LSU: EF498017-27

References

Fukuyo, 1979) (Chinain et al., 1997) (Richlen et al., 2008) (Litaker et al., 2009) (Chinain et al., 1999a) (Litaker et al., 2009)

Broad 1p plate-2’ SSU: EF202902-07 Hatchet shapedD1-D3 LSU: Dorsal end 1p EF202976-82 oblique-smaller cell D8-D10 LSU: size than G. EF498076-80 carolinianus G. carolinianus (78.2 ± 4.8) – Broad 1p plate-2’ SSU: EF202897(Litaker et (Vandersea, (87.1 ± 7.1) – Hatchet shapedEF202901 al., 2009) Litaker, Faust, (51.4 ± 5.2) Dorsal end 1p D1-D3 LSU: Kibler, Holland et oblique-larger cell EF202973-75 Tester) size than G. D8-D10 LSU: polynesiensis EF498035-37 Genetically described phylotypes Gambierdiscus Not described D8-D10 LSU: (Litaker et ribotype 1 GU968512-20, al., 2010) GU968523 Gambierdiscus Not described D8-D10 LSU: (Litaker et ribotype 2 GU968499-500, al., 2010) GU968503, GU968505, GU968507-11 Gambierdiscus Not described SSU: AB64229-76, (Kuno et al., sp. type 1 AB605799-800, 2010) AB605811-12 (Nishimura LSU D8-D10: et al., 2013) AB765908-13 Gambierdiscus Not described SSU: AB764277(Kuno et al., sp. type 2 96 2010) LSU D8-D10: (Nishimura AB765913-18 et al., 2013) Gambierdiscus Not described SSU: AB764296(Nishimura sp. type 3 300 et al., 2013) LSU D8-D10: AB765923-24 The abbreviations are: SSU, small ribosomal subunit gene; LSU large ribosomal subunit gene. The following is an overview of the main morphological characteristics for each described species of Gambierdiscus. The original species descriptions consist of a comprehensive account of their characteristics Gambierdiscus cells are large (60-100 μm), armoured, have a distinct plate pattern and fishhook shaped apical pore. Species are either 4

Chapter 1

anterio-posteriorly compressed (lenticular) or slightly laterally compressed (globular) (Figure 1.2). The two globular species (G. yasumotoi and G. ruetzleri) can be distinguished from each other by cell size, size and shape of the 2’ apical and 2’’’’ antapical plate and depth to width ratio, described in detail in Litaker et al. (2009) . The remaining 9 species are anterio-posteriorly compressed and broadly classified by either a narrow (G. australes, G. belizeanus, G. pacificus and G. excenreicus) or broad (G. polynesiensis, G. carolinianus, G. toxicus, G. caribaeus, and G. carpenteri) 1p posterior intercalary plate. Among the species with a narrow 1p posterior intercalary plate can be further distinguished by either heavily areolated cell surface (G. belizeanus) or smooth cell surface species (G. australes, G. pacificus and G. excentricus). Species with a smooth cell surface can be distinguished by either having a hatchet-shaped 2’ apical plate (G. pacificus) or more conventional rectangular shaped 2’ apical plate (G. australes and G. excentricus). G. excentricus is at least 1.5 times wider and deeper than G.australes and further specifics distinguishing the two are described in detail in the original descriptions of the species (Chinain et al., 1999a, Fraga et al., 2011). Species that have a broad 1p posterior intercalary plate can be further differentiated as having a rectangular shaped 2’ apical plate (G. caribeaus and G. carpenteri) or a hatchet shaped 2’ apical plate (G. toxicus, G. polynesiensis and G. carolinianus). G. toxicus can be further discerned by a pointed dorsal end to the 1p posterior intercalary plate. Further differences between G. polynesiensis & G. carolinianus are detailed in Litaker et al. (2009). G. caribeaus and G. carpenteri, both possessing a rectangular shaped 2’ apical plate, are distinguished by the shape of 4’’ precingular plate, which is symmetric in G. caribaeus and asymmetric in G. carpenteri. The size and shape of the sulcal plates and various other specific morphological characteristics have also been described in the original descriptions of the species (Faust, 1995, Holmes, 1998, Chinain et al., 1999a, Litaker et al., 2009, Fraga et al., 2011). These features are straightforward to observe using light and scanning electron microscopy, however, within some species, a considerable amount of variability in features such as the size and shape of individual plates may be present.

5

Chapter 1

Figure 1.2: Comparative line drawings of the epitheca for 11 Gambierdiscus species. Scale bar equals 50 μm. Modified from Litaker et al., 2009. Another tool to identify different species of Gambierdiscus is by comparing sequences that are known to be characteristic at the species level, such as regions of rRNA genes. Based on phylogenetic analysis of regions of the SSU (small ribosomal subunit) rDNA, LSU (large ribosomal subunit) rDNA and ITS (internal transcribed spacer) rDNA, the genus Gambierdiscus is monophyletic (Chinain et al., 1999a, Richlen et al., 2008, Litaker et al., 2009, Kuno et al., 2010, Litaker et al., 2010, Fraga et al., 2011, Nishimura et al., 2013). Further, the lenticular and globular species form two distinct clades. Phylogenetic analysis has shown that the two globular species (G. ruetzleri and G. yasumotoi) diverged relatively early in the evolution of the genus 6

Chapter 1

Gambierdiscus (Litaker et al., 2009, Nishimura et al., 2013). Also, G. ruetzleri and G. yasumotoi are the two most closely related species in the genus. Based on LSU rDNA D08-D10 sequences, the mean p distance within species is 0.002 ± 0.002, and between species is 0.121 ± 0.036 (calculated based on sequences from 10 species/phylotypes) where minimum p distance between G. ruetzleri and G. yasumotoi is 0.007 (Nishimura et al., 2013). Using SSU rDNA sequences, the mean p distance within species is 0.003 ± 0.002 and between species is 0.139 ± 0.042 (calculated based on sequences from 10 species/phylotypes) where minimum p distance between G. ruetzleri and G. yasumotoi is 0.004 (Nishimura et al., 2013). These statistics are indicative of putative unknown species, and can be very useful in cases where morphological, physiological or other data is not yet available, or a strain is not present in culture. Based on D8-D-10 LSU rDNA phylogenetic analysis, 2 new putative phylotypes Gambierdiscus ribotype 1 and Gambieridsuc ribotype 2 were reported (Litaker et al., 2010), as the two clusters/clades separated from the others and there genetic distances equalled or exceeded those among the 10 described species (Litaker et al., 2010) (Table 1). Similarly, 3 new putative species/phylotypes of Gambieridiscus (Gambieridsucs sp. Type 1, type 2 and type 3) have been described from Japan based on differences in the regions D8-D10 of the LSU and SSU rDNA (Kuno et al., 2010, Nishimura et al., 2013) (Table, 1). In this case, the p distances between these two novel clades and known species of Gambierdiscus were larger than those separating G. yasumotoi from G. ruetzleri. Although the genetic data indicates that these phylotypes may be new species, their morphological circumscriptions are needed to support their status as new species. As sampling around the world becomes more intensive, it is likely that new species of Gambierdiscus will be described.

1.3

Geographic distribution and abundance Gambierdiscus is widely distributed in coastal zones at tropical and subtropical latitudes.

However, the distribution of species of Gambierdiscus is still poorly understood, as the discrimination of different species of Gambierdiscus has only occurred recently (Table 1.2).

1.3.1

The Pacific and Indian Ocean regions Gambierdiscus is named after the Gambier Islands in French Polynesia, where it was first

identified, (Adachi and Fukuyo, 1979) , and since then, G. toxicus, G. belizeanus, G. yasumotoi, G. australes, G. pacificus, G. polynesiensis, G. caribaeus and G. carpenteri have been reported from various Pacific islands, Hawaii, Australia, Southeast Asia and the Northern Indian Ocean (Table 2). Recently, three genetically distinct species from coastal and temperate waters of Japan were reported (Nishimura et al., 2013) (Table 1.2). In addition, Gambierdiscus has been reported from the Philippines (Gillespie, 1987), Hong Kong (Lu and Hodgkiss, 2004), Indonesia (Praseno and Wiadnyana, 1996) and Mauritius (Hurbungs et al., 2002), although species diversity in these areas is not known. Gambierdiscus has also been reported from the Mexican 7

Chapter 1

Pacific coast (Ceballos-Corona, 2006) and regions around Madagascar (Grzebyk et al., 1994), where cases of CFP have also been previously reported (Habermehl et al., 1994, HernandezBecerril et al., 2007).

Table 1.2: Geographic distribution and toxicity of different Gambierdiscus species. Species

Geographical Distribution

G. toxicus

Tahiti, French Polynesia1, 2, Mexican Caribbean3, New Caledonia2, Reunion Island2, Indian Ocean, Nha Trang –Vietnam4, 5 Belize7, Florida8, Mexican Caribbean3, Malaysia9, Pakistan10, Queensland, Australia, St. Barthelemy Island-Caribbean15 Singapore12, Japan13, Mexican Caribbean3, Queensland, Australia, Nha Trang-Vietnam5 French Polynesia2, Japan13, Cook Islands14, Hawaii USA8, Pakistan10

G. belizeanus

G. yasumotoi

G. australes

G. pacificus

G. polynesiensis

G. caribaeus

G. carolinianus

G. carpenteri

G. ruetzleri G. excentricus

8

French Polynesia2, Marshall Islands & Society Islands Micronesia15, Kota Kinabalu and Sipandan Island16, Nha Trang-Vietnam5 French Polynesia2, Canary Islands17, Pakistan10, Nha Trang-Vietnam5 Florida8, Belize-Caribbean8, Tahiti8, Palau, Hawaii8, Flower Gardens-Gulf Of Mexico, Osho Rios-Jamaica11, Bahamas15, Grand Caymam Island, Tol-truk Micronesia 15, Jeju Island Korea18 North Carolina, USA, Atlantic ocean8, Bermuda, Mexico15, Puerto Rico11, Flower Gardens-Gulf Of Mexico11, Osho Rios-Jamaica11, Crete-Greece11 Belize8, Guam8, Fiji8, Hawaii 15, Dry Tortugas-Florida11, Flower GardensGulf Of Mexico11, Osho RiosJamaica11 Merimbula & Exmouth Australia19 North Carolina, USA8, BelizeCaribbean8 Canary Islands17

Toxicity- Various Assays CTX MTX MBAMBAnegative 2 positive2 RBApositive6 RBAHELApositive6 positive11

Toxicity LC-MS

MBApositive12

N/K

N/K

MBApositive14, RBApositive6 MBApositive2

HELApositive11, MBApositive 14 MBApositive2

CTXN/D14 MTXYes N/K

MBApositive2, RBApositive6 N/K

MBApositive2

CTXYes6 MTXN/K N/K

N/K

HELApositive11

N/K

N/K

HELApositive11

N/K

N/K

HELApositive11 NCBApositive17

N/K

NCBApositive17

HELApositive11

N/K

N/K

N/K

Chapter 1

Species

Geographical Distribution

Toxicity- Various Assays CTX MTX N/K N/K

Toxicity LC-MS

Gambierdiscus Belize-Caribbean15 N/K ribotype 1 Gambierdiscus Belize-Caribbean15, MartiniqueN/K HELAN/K ribotype 2 Caribbean15, Puerto Rico11 positive11 Gambierdiscus Japan13 MBAMBAN/K sp. type 1 positive13 positive13 Gambierdiscus Japan13 MBAMBAN/K sp. type 2 negative13 negative13 Gambierdiscus Japan13 MBAMBAN/K 13 13 sp. type 3 negative positive The references are: 1. (Adachi and Fukuyo, 1979), 2. (Chinain et al., 1999a), 3. (HernándezBecerril and Almazán Becerril, 2004), 4. (Roeder et al., 2010), 5. (The, 2009), 6. (Chinain et al., 2010), 7. (Faust, 1995), 8. (Litaker et al., 2009), 9. (Leaw et al., 2011), 10. (Munir et al., 2011), 11. (Holland et al., 2013), 12. (Holmes, 1998), 13. (Nishimura et al., 2013), 14. (Rhodes et al., 2010), 15. (Litaker et al., 2010), 16. (Mohammad-Noor et al., 2007), 17. (Fraga et al., 2011), 18. (Jeong et al., 2012), 19. (Kohli et al., 2013). The abbreviations are: N/K, Not known; N/D not detected; MBA, Mouse bioassay; HELA, Human erythrocytes lysis assay; NCMA, Neuro-2a cell binding assay; RBA, Receptor binding assay;

1.3.2

The Atlantic Ocean region Early accounts of Gambierdiscus “look-alike” species date back to 1948 from Cape

Verde Islands (de Silva, 1956) and 1979 from Key Largo, Florida (Taylor, 1979b). So far, G. toxicus, G. belizeanus, G. yasumotoi, G. polynesiensis, G. caribaeus, G. carolinianus, G. carpenteri, G. ruetzleri, G. excentricus, Gambierdiscus ribotype 1 and Gambierdiscus ribotype 2 have been reported from the east coast of USA, Caribbean and the Mediterranean regions (Table 1.2). There are many other regions where Gambierdiscus has been reported, however, the exact species are yet to be determined. These include Cyprus, Rhodes, Saronikos Gulf (Aligizaki et al., 2009, K. Aligizaki et al., 2010), French West Indies (Lobel et al., 1988), Cuba (Delgado et al., 2006) and Veracruz (Okolodkov et al., 2007). Other confirmed reports of Gambierdiscus occurrence in Central and South America in the literature are from Costa Rica and Brazil (M. Montero pers. comm. in (Parsons et al., 2012) ). From Africa, there has been only one direct observation of Gambierdiscus, from the coast of Angola (Berdalet et al., 2012), however CFP cases from the west coast (Canary Islands and Cameroon) (Bienfang et al., 2008) of Africa have been reported, indicating the presence of Gambierdiscus in that region. Certain species of Gambierdiscus have been designated as being endemic to either the Pacific or the Atlantic regions (Litaker et al., 2010, Berdalet et al., 2012). So far, G. australes and G. pacificus have only been reported from the Pacific and G. ruetzleri, G. excentricus and Gambierdisucs ribotype 1 and 2 are only reported from the Atlantic region (Table 1.2). G. belizeanus, G. caribeaus, G. carpenteri and G. carolinianus are widely distributed in the 9

Chapter 1

Atlantic and Pacific Oceans (Litaker et al., 2009, Litaker et al., 2010, Berdalet et al., 2012). G. yasumotoi is widely distributed in the Pacific, however, there is only one report of its occurrence in Mexican-Caribbean (Hernández-Becerril and Almazán Becerril, 2004), which was reported before the discovery of the other globular species G. ruetzleri, which is widely distributed in the Atlantic region (Litaker et al., 2009). The distribution of G. toxicus needs to be refined, due to numerous misidentifications in the literature. G. polynesiensis is widespread in the Pacific (Table 1.2) with only one confirmed report from the Canary Islands in the Atlantic region (Fraga et al., 2011). Both, Litaker et al, 2010 and Berdalet et al, 2012 (Litaker et al., 2010, Berdalet et al., 2012) mention that none of the Pacific-specific species have been observed in hundreds of field samples analysed from Atlantic regions (Caribbean/Gulf of Mexico/ West indies/Southeast US coast from Florida to North Carolina). The absence of Atlantic-specific species in the Pacific region has not been confirmed, as the majority of the vast Pacific region remains unexplored. As under-sampling and under-reporting have occurred worldwide, but particularly in the Pacific region, much more work needs to be undertaken in order to determine whether endemism or restricted distributions exist in species of Gambierdiscus. While multiple species of Gambierdiscus can co-occur in one region, equally, there are regions from where only one species has been reported. For example, in Heron Island (Queensland, Australia) there are at least 3 species of Gambierdiscus that co-occur, however further south in Merimbula, New South Wales only G. carpenteri is known to occur (Murray unpubl. data). Localised benthic blooms of Gambierdiscus have been noted in the literature from both the Pacific and Atlantic regions (Nakajima et al., 1981, Withers, 1984, Chinain et al., 1999b, Darius et al., 2007). Cell densities in such blooms can range from anywhere between 10,000 to 100,000 cell g-1 wet weight algae (Litaker et al., 2010). There are no accurate estimates of cell densities at which a Gambierdiscus bloom leads to a CFP epidemic. The onset of CFP may also depend on other factors, such as the fact that different species of Gambierdiscus have varying toxicities. For example, in 2010, an unidentified species of Gambierdiscus was reported in Greece, however no CFP outbreaks have been reported there (Caillaud et al., 2010). In most habitats where species of Gambierdiscus occur, cell densities are below 1,000 cells g-1 wet weight algae (Litaker et al., 2010), however in some environments Gambierdiscus spp. are known to occur year-round at such cell densities (Chinain et al., 1999b). A constant exposure of low densities of cells could also lead to a build up of CFP- related toxins in fish. Much more research needs to be done in order to understand the relationship between Gambierdiscus abundance and CFP outbreaks. This is particularly challenging, as benthic dinoflagellates inhabit areas where quantitative sampling of microbial eukaryotes is not straightforward, for example, in sediments and on the surface of dead corals. Also, Gambierdiscus cell distribution can be very patchy, even over small distances, making it hard to 10

Chapter 1

estimate mean Gambierdiscus cell densities over a larger area (Ballantine et al., 1988, Lobel et al., 1988, Litaker et al., 2010).

1.4

CTXs and MTXs CTXs are sodium channel activators, particularly affecting the voltage sensitive channels

located along the nodes of Ranvier (peripheral nerve cells) (Lewis et al., 1992, Mattei et al., 1999, Lewis et al., 2000). When the sodium channels are activated there is a massive influx of Na+ ions, resulting in cell depolarisation (Lewis et al., 1992, Mattei et al., 1999, Lewis et al., 2000). This leads to the onset of spontaneous action potentials in effected cells, causing various symptoms in humans. Symptoms can include but are not limited to gastrointestinal, neurological and sometimes cardiovascular, in cases of severe intoxication (Sims, 1987) and can vary depending on geographical region (Lewis et al., 2000, Dickey, 2008). This can be due to the structural differences of CTXs in different regions, therefore it is very important to characterise CTXs from Pacific, Caribbean and the Indian Oceans. Local understanding of CTX accumulation patterns in different fish species can also help prevent CFP. However, the accurate identification of exact congeners of CTXs is necessary, in order to understand the toxicology and evaluate the local risks of CFP. Structurally, CTXs are thermostable, cyclic polyether ladders, which are liposoluble. They have been isolated from fish and different species of Gambierdiscus (Table 1.3). Based on their origin and differences in the structure of these toxins, they are divided into P-CTXs (Pacific Ocean), C-CTXs (Caribbean region) and I-CTXs (Indian Ocean). Due to their structural differences, P-CTXs are further divided into type I and type II, as suggested by Legrand et al. (Legrand et al., 1998). Type I P-CTXs have 13 rings and 60 carbon atoms (Murata et al., 1990, Lewis et al., 1991, Lewis and Holmes, 1993, Yasumoto et al., 2000). This category consists of the first CTX to be fully structurally described as CTX1B (Murata et al., 1990) (or CTX-1 as described by Lewis et al. 1991, (Lewis et al., 1991)) from moray eels, which is the principal toxin in the carnivorous fish from the Pacific (Murata et al., 1990, Lewis et al., 1991). Two other type I P-CTXs i.e. CTX-2 and CTX-3 were also described from the same extracts, which have slight variations in their structures leading to different toxicities in mice (Lewis et al., 1991) (Table 3). It has also been suggested that CTX-1, CTX-2 and CTX-3 may be derived from dinoflagellate precursors known as CTX-4A and CTX4B (also named as GTX-4B in (Murata et al., 1990)) (Lewis and Holmes, 1993, Yasumoto et al., 2000). Recently, CTX-4A and CTX-4B have been isolated from G. polynesiensis culture extracts (Chinain et al., 2010). CTX3C is a type II P-CTX with 13 rings, 57 carbon atoms and was first isolated from cultures of Gambierdiscus sp. (Satake et al., 1993) and later from G. polynesiensis (Chinain et al., 2010). Two more congeners of CTX3C called as 49-epi-CTX-3C (also called as CTX-3B in (Chinain et al., 2010)) and M-seco-CTX-3C have also been isolated from Gambierdiscus sp. (Satake et 11

Chapter 1

al., 1993) and G. polynesiensis (Chinain et al., 2010). Later, 2 new type II P-CTXs i.e. 2,3 dihydroxyCTX3C (also called as CTX2-A1) and 51-hydroxyCTX3C were isolated from Moray eel (Satake et al., 1998) that might be oxygenated metabolites of CTX3C (Yasumoto et al., 2000).

Table 1.3: Different congeners of CTXs and MTXs. Origin

Toxin Name

Molecular Ion [M +H]+

Pacific (type I)

CTX1B1, CTX-12

1111.61, 2

Source

Ciguatoxins Moray eel (Gymnothorax javanicus) 1 Moray eel (Lycodontis javanicus, Muraenidae) 2

Toxicity (LD 50 doses calculated via i.p. injection in mice) CTX1B- 0.35 μg/kg1 CTX-1- 0.25 μg/kg2

Moray eel (Lycodontis 2.3 μg/kg2 2 javanicus, Muraenidae) CTX-3 1095.52 Moray eel (Lycodontis 0.9 μg/kg2 2 javanicus, Muraenidae) 3 CTX4A 1061.6 Gambierdiscus sp. 3 12 μg/kg4 G. polynesiensis4 3 CTX4B 1061.6 Gambierdiscus sp. 3 20 μg/kg4 4 G. polynesiensis 5 Pacific CTX3C 1023.6 Gambierdiscus sp. 5 2.5 μg/kg4 4 (Type II) G. polynesiensis 49-epi1023.64 Gambierdiscus sp.5 8 μg/kg4 4 CTX-3C G. polynesiensis M-seco1041.64 Gambierdiscus sp. 5 10 μg/kg4 4 CTX-3C G. polynesiensis Caribbean C-CTX-1 1141.66, 7 Horse-eye jack (Caranx 3.6 μg/kg6 6, 7 (Type-3) latus) C-CTX-2 1141.66, 7 Horse-eye jack (Caranx Toxic6 6, 7 latus) Indian I-CTX-1 1141.68 Red bass (Lutjanus bohar) Toxic8 and red emperor (Lutjanus sebae) 8 Maitotoxins 9, 10 Pacific MTX-1 3422 Gambierdiscus sp.9 0.05 μg/kg10 MTX-2 32989 Gambierdiscus sp. 9 0.08 μg/kg9 9 9 MTX-3 1060 Gambierdiscus sp. Toxic9 The references are: 1. (Murata et al., 1990), 2. (Lewis et al., 1991), 3. (Yasumoto et al., 2000), 4. (Chinain et al., 2010), 5. (Satake et al., 1993) 6. (Vernoux and Lewis, 1997), 7. (Pottier et al., 2002b) 8. (Hamilton et al., 2002b) 9. (Holmes and Lewis, 1994), 10. (Murata et al., 1993). CTX-2

12

1095.52

Chapter 1

Figure 1.3: Structure of Ciguatoxins (CTX) and Maitototoxin-1. P-CTX-1, P-CTX-2 and CCTX-1 were derived from fish and P-CTX-3C, P-CTX-4B and Maitotoxin-1 were derived from Gambierdiscus spp. Caribbean CTXs (Type-3) are slightly bigger than P-CTXs and have 14 rings and 62 carbon atoms (Vernoux and Lewis, 1997, Lewis et al., 1998, Pottier et al., 2002b, Pottier et al., 2003). Many congeners of C-CTXs have been isolated from carnivorous fish, which includes C13

Chapter 1

CTX1, C-CTX-2, C-CTX-1141, C-CTX-1127, C-CTX-1143, C-CTX-1157, C-CTX-1159 (Vernoux and Lewis, 1997, Lewis et al., 1998, Pottier et al., 2002b, Pottier et al., 2003). Unlike P-CTXs there have been no reports of C-CTXs being originating from Gambierdiscus sp. However, recently G. excentricus has been identified as a major CTX producer in the Caribbean (Fraga et al., 2011) and CTXs from this strain are being characterised. Recently 4 CTXs (I-CTX-1, I-CTX-2, I-CTX-3, I-CTX-4) have been isolated from carnivorous fish from Indian Ocean and have higher molecular ion masses than P-CTXs and CCTXs (Hamilton et al., 2002a, Hamilton et al., 2002b, Caillaud et al., 2010). However, there structures need to be elucidated (Hamilton et al., 2002a, Hamilton et al., 2002b). I-CTX-1 is toxic to mice via intraperitoneal injection (Hamilton et al., 2002b). Based on Mouse bioassays (MBA), different congeners of CTXs can have variable toxicities (Table 1.3), however this needs to be further validated as well. Maitotoxins (MTXs) are one of the largest non-proteinous and highly toxic natural products known (Yokoyama et al., 1988, Murata et al., 1993). This, polyether ladder type compound was first discovered as a water-soluble toxin in the guts of herbivorous fish Acanthurids (surgeonfish) in 1976 (Yasumoto et al., 1976). In the 1990s, stereoscopic studies and partial synthesis were used to determine the structural elucidation and stereochemistry of the extraordinary complex and large MTX (Murata et al., 1993, Murata et al., 1994, Murata and Yasumoto, 1995, Satake et al., 1995, Nonomura et al., 1996, Zheng et al., 1996). Simultaneously, Holmes & Lewis described 2 large (MTX-1 & MTX-2) and one small MTX (MTX-3) from different strains of Gambierdiscus sp. isolated from Queensland, Australia (Holmes and Lewis, 1994) (Table 1.3). MTX-1 from this study may have been the MTX originally described from guts of Acanthurids, however it is not clearly proven. When compared to other natural toxins, MTX is a highly potent calcium channel activator (LD50 0.05 μg/kg, i.p., mice), only exceeded by a handful of bacterial proteinaceous toxins (Yokoyama et al., 1988, Murata et al., 1993). Despite its high level of potency, the complete mode of action and the primary target of MTX in mammalian cells have not yet been fully elucidated. In fact, the activation of voltage dependent calcium channels induced via MTXs is a secondary effect of membrane depolarisation (for review and more details see ref. (Van Dolah, 2000) ). Recently, it has been reported that the biophysical mechanisms of pacific MTXs are different to Caribbean MTXs (Lu et al., 2013). Whether this is due to a structural difference is not known, as the Caribbean MTXs have not been fully characterised. Although MTX appears to have a low tendency of accumulating in fish flesh, as compared to stomach or intestines (Yasumoto et al., 1976), its possible role in CFP cannot be disregarded, as eating uneviscerated fish is a common practice in many Pacific Island nations. The sulphate esters in the structures of MTXs make it amenable to detect and quantify MTX by LC_ESI_MS (Liquid chromatography-electronspray ionisation-Mass spectrometry) (T. Harwood pers. comm.) and Solvolysis (desulphonation) 14

Chapter 1

reduces the toxicity of MTXs significantly, at least by 100 fold (Murata et al., 1991). However, more research is essential to understand the exact role of MTXs in CFP including its mode of action and target in mammalian cells. Cyclic polyether ladders are almost exclusively known to be produced by dinoflagellates. Other than CTXs and MTXs, this class of secondary metabolites also includes Brevetoxins (BTXs), produced by Karenia spp. (Shimizu et al., 1986) and Yessotoxins (YTXs), produced by wide array of dinoflagellates including Lingulodinium polydrum (Draisci et al., 1999), Gonyaulax spinifera (Rhodes et al., 2006) and Protoceratium reticulatum (Eiki et al., 2005). Based on their high structural similarities, the synthesis of these compounds likely involves common biosynthetic mechanisms (Lin et al., 1981, Golik et al., 1982, Chou and Shimizu, 1987). Stable-isotope labelling of precursors, to elucidate the biosynthesis pathway of CTXs and MTXs has never been performed. However, precursor studies to reveal the biosynthesis pathways of BTXs and YTXs have indicated the polyketide origin of these cyclic polyether ladders (Shimizu 1986, Lee et al., 1989). Several schematic pathways involving different enzymes have been suggested and are detailed in Kalaitzis et al (Kalaitzis et al., 2010) and Kellmann et al (Kellmann et al., 2010). It is speculated that the biosynthesis involves the normal polyketide synthase (PKS) enzyme complex with a few additional enzymes i.e. expoxidases and thioesterases (Shimizu, 2003). Essential domains present in the PKS are: acyltransferase domain (AT), β-ketosynthase domain (KS) and acyl carrier protein (ACP) (Khosla et al., 1999). In addition to that PKS can include β-ketoacyl reductase (KR), enoyl reductase (ER) and dehydrogenase (DH) domains (Khosla et al., 1999). In the past 10 years, a few genes that encode the essential domains of the PKSs, particularly KS domains in dinoflagellates have been identified for the first time. However, with the availability of next generation sequencing tools, a few candidate genes encoding KS and KR domains in Karenia brevis have been associated with biosynthesis of BTXs (Monroe and Van Dolah, 2008). A recent study published a comprehensive transcriptome library of Lingulodinium polydrum for which genes encoding KS domains were reported, however no link between these genes and YTX production has been established (Beauchemin et al., 2012). In the past, a few studies have identified genes encoding KS domains in Amphidinium sp. (Murray et al., 2012a), which produces numerous macrolides (cyclised linear polyethers) such as amphidinolides. No studies have been done to identify genes involved in CTX and MTX biosynthesis. However, an extensive marine microbial eukaryote transcriptome project, undertaken by the Moore Foundation, is in the process of sequencing 652 trancriptomes, which includes 2 strains of Gambierdiscus species. Analysis of data obtained for such diverse arrays of dinoflagellate species may shed light on the genes involved in secondary metabolite synthesis in dinoflagellates.

15

Chapter 1

1.5

Toxicity of different species of Gambierdiscus There is clear evidence that Gambierdiscus species produce CTXs and/or MTXs (Murata

et al., 1990, Holmes et al., 1991, Satake et al., 1993, Holmes and Lewis, 1994, Satake et al., 1996). However, many wild and cultured strains of Gambierdiscus have not been found to produce detectable amounts of CTXs (Gillespie et al., 1985, Holmes et al., 1990). Unfortunately, most of these studies describe the identity of the cultures as Gambierdiscus toxicus, since it was the only known species of Gambieriscus at that time. It is imperative to study the toxin profile of all of the species and genotypes now known. Table 1.2 provides the data available on the toxicity of each species of Gambierdiscus detected via various assays and LC-MS (Liquid chromatography-mass spectrometry) based detection. In 2010, Chinain et al. (Chinain et al., 2010) described the toxin profile of G. polynesiensis based on LC-MS analysis and receptor binding assay (RBA). This species produces both Type 1 (CTX-4A, CTX-4B) and Type 2 P-CTXs (CTX-3C, M-seco-CTX-3C, 49-epiCTX-3C), however P-CTX-3C was the major toxin produced by this species. Two different strains of G. polynesiensis were tested and found to produce same suite of toxins, in different proportions (Chinain et al., 2010). Haemolytic activity was found in a study of 56 strains of 6 different species (G. belizeanus, G. caribaeus, G. carolinianus, G. carpenteri, Gambierdiscus ribotype 2) over a period of 2 years, using the human erythrocyte lysis assay (HELA) (Holland et al., 2013). The intraspecific toxicity varied slightly among different strains of same species, however the level of toxicity of each strain remained unchanged over the period of the study (Holland et al., 2013). The HELA assay measures the haemolytic activity of Gambierdiscus cell extracts, which is directly proportional to the amount of MTX present. For this assay, it is widely assumed that all haemolytic activity is due to the MTXs in Gambierdiscus cell extracts, however, there is no direct evidence in the literature, which proves that the haemolytic activity in the extracts, is due to MTX specifically. The water-soluble fraction of the extracts of G. polynesiensis has been found to be toxic via MBA (Chinain et al., 1999a), indicating the presence of MTXs. However, the toxins that produced this effect have not been characterised from this strain. Another species from the Caribbean, G. excentricus, may produce CTXs and MTXs (as determined via Neuro-2a cell based assay) (Fraga et al., 2011), however the exact toxin profile needs to be verified via LC-MS analysis. The toxicities of the liposoluble and water-soluble fractions of G. australes extracts, isolated from the Cook Islands, were found to be toxic via MBA, indicating the presence of CTXs and MTXs (Rhodes et al., 2010). However no CTXs were detected via LCMS analysis (Rhodes et al., 2010). Another strain of G. australes from French Polynesia tested positive for CTXs via the RBA, however the level of toxicity was low when compared to G. polynesiensis (Chinain et al., 2010). These results are intriguing and require further analysis. While bioassays are important to determine toxicity, only LC-MS based analysis techniques can determine the exact toxin profile of different species of Gambierdiscus. As we only know the 16

Chapter 1

partial toxin profiles of two species of Gambierdiscus via LC-MS based techniques, this area of research needs urgent attention.

1.6

Detection of CTXs and MTXs in seafood Originally, CFP was derived from the word “cigua”, used by native Cubans to describe a

turban shelled snail and implicated in an outbreak of the sickness in Spanish explorers to Cuba in the 1500s (Gudger, 1930). The occurrence of CTX in the turban snail Turbo argyrostoma has been confirmed (Yasumoto and Kanno, 1976). However, to date, the majority of cases reporting occurrences of CFP have followed consumption of large reef fish (e.g. (Hokama and Yoshikawa-Ebesu, 2001, Lewis, 2001, Dechraoui et al., 2005, Laurent et al., 2005)). This circumstance has been a critical factor in the diagnosis of the disease, as in many cases there has been no fish sample retained for chemical verification or the appropriate test facilities are unavailable. Although hundreds of cases of CFP have been documented worldwide, it is estimated that less than 20% of actual cases have been reported (Dickey and Plakas, 2010). There is a high likelihood of misdiagnosis for CFP. The number of documented symptoms, which are in excess of 175 (Sims, 1987), may vary depending on portion size (Wong et al., 2008), individual susceptibility or accumulation of toxin with age (Bagnis et al., 1979, Glaziou and Martin, 1993) and could also be associated with other illnesses (e.g. decompression sickness (Adams, 1993), chronic fatigue sydnrome, multiple sclerosis (Lindsay, 1997, Ting and Brown, 2001) and brain tumors (Lindsay, 1997)). The number of fish species implicated in ciguatera outbreaks is suggested to be of the order of several hundred (Halstad, 1978, FAO, 2004). However, with the above limitations and the absence of a reliable, commercially available, test kit it is difficult to express an exact figure. While carnivorous fish are the main culprits, herbivorous fish (e.g. surgeonfish and parrotfish), a key component of the toxic food chain (Randall, 1958, Cruz-Rivera and Villareal, 2006), have also been linked to CFP outbreaks. Table 4 provides a summary of over 90 fish species and other marine fauna that have tested positive for CTXs, from ciguatera prone regions and following reported outbreaks. The CTX-positive cases in Table 4 are predominantly concerned with the mid-latitude tropical and sub-tropical zones. This is fitting with the distribution of Gambierdiscus as described in Table 2. However, CFP has also been reported in non-endemic areas because of an increase in seafood imports (Glaziou and Legrand, 1994, Ting and Brown, 2001). While the majority of studies have focused on reef fish, toxin accumulation has been observed in eels, sea cucumbers, starfish, seals and jelly fish (see table 4 and references therein). Sharks have also been suspected of causing CFP following outbreaks of human illness, remnant samples for testing were unavailable (Boisier et al., 1995, Lehane and Lewis, 2000). Further studies are required to address the deficit in information for species other than fish and to identify potential toxin vectors in coastal systems. For the most part, CFP studies have focused on CTX rather 17

Chapter 1

than MTX. The MBA has been used previously to test for MTX, with positive results in Ctenochaetus striatus (striped bristletooth) (Bagnis et al., 1986). A gap in our existing knowledge is whether the presence of MTX in small (herbivorous) fish species is transferred up the food chain to larger carnivorous species. Often, in small island nations, native fishermen are aware of ciguatera prone zones and avoid certain fish species. Such knowledge certainly has its merits, however, a study by (Darius et al., 2007) in French Polynesia demonstrated the presence of CTXs in fish species that were considered safe to eat by locals. Experimentally, CTX toxin profiles and structures have been determined by chromatographic techniques (HPLC, UPLC and LC-MS), accompanied by nuclear magnetic resonance (NMR) (Legrand et al., 1989, Murata et al., 1990, Lewis et al., 1991, Satake et al., 1996) and radio ligand binding (RLB) (Hamilton et al., 2002a, Hamilton et al., 2002b). However, these methods are not commonplace or practical for routine testing, as they are costly and require special expertise. Confirmation of toxin by UPLC/HPLC followed by LC-MS involves the isolation and fractionation of the various CTX compounds, and their known molecular weights (see table 3). Although a rapid method for sample analysis has been proposed (Lewis et al., 2009), acquiring purified CTX standards is problematic due to the limited supply of natural CTX compounds (Berdalet et al., 2012) and though artificial synthesis of CTX is possible (Hirama et al., 2001), it is highly complex. Without a consistent source of reference material, absolute quantification of CTXs and their congeners is hard to achieve. In addition, technical issues such as co-eluting peaks of similar compounds and inhibiting/promoting matrix effects remain unresolved. Several biological assays have been developed for the detection of ciguateric fish. These have included the use of chickens (Pottier et al., 2000), cats (Larson and Rothman, 1967), mongooses (Hokama et al., 1977), diptera larva (Labrousse and Matile, 1996), brine shrimp (Granade et al., 1976) and mosquitoes (Bagnis et al., 1987). However, each assay has its own constraints and limitations, largely relating to toxin specificity and quantification but also due to inefficiencies and ethical considerations (summarised in de Fouw, 2001, (Dickey and Plakas, 2010). While the MBA by intraperitoneal injection does not provide a linear dose-response relationship with CTX toxicity (Hoffman et al., 1983), it remains the most widely used biological assay (see Table 4). Numerous biochemical assays have been proposed as alternatives to biological assays for testing seafood. The development of a radioimmunoassay (Hokama et al., 1977) progressed to a cheaper alternative enzyme-linked immunosorbent assay (ELISA) with higher throughput (Hokama et al., 1983). The ELISA test has recently shown promising correlations with biological assays (Campora et al., 2008, Campora et al., 2010). Stick enzyme immunoassay (SEIA) (Hokama, 1985) and solid phase immunoassay (SPIA) (Hokama, 1990) tests have led to the development of commercial kits (i.e. Cigua-check ® and Ciguatect ®). However, these products have yielded a large number of false positive and false negative results (Wong et al., 2005) and the 18

Chapter 1

Cigua-check ® test is no longer being manufactured. Other assays utilised for screening CTXs in fish are the sodium channel binding assay (N2A) (Dickey, 2008) and RBA (Poli et al., 1997, Darius et al., 2007). Both of these assays have shown promising results and have been recommended by the European Food Standard Association (EFSA, 2010). These assays cannot quantify specific congeners of CTXs and MTXs. This can only be achieved via further development and validation via LC-MS analysis, and there is an urgent need to do so. The progress has been disadvantaged by the lack of available purified standards (Guzman-Perez and Park, 2000). Other challenges are the presence of more than one type of CTXs (see Table 3) and any other unidentified metabolites produced by Gambierdiscus being present in fish specimens (Endean et al., 1993, Vernoux and Lewis, 1997).

Table 1.4: Different congeners of Ciguatoxins detected by various assays in seafood and other animals. Latin name (Common name) Seriola dumerili (Greater amberjackKahala)

Seriola fasciata (Lesser amberjack)

Seriola rivoliana (Almaco jack-Kahala)

Sphyraena barracuda (Great barracuda)

Sphyraena jello (Pickhandle barracuda) Sphyraena sp. (Barracuda) Sphyraena spp. (Barracuda fish eggs)

Source

CTX (if detected) AmberjackC Canary Islands1, C-CTX-17, CTXSelvagens Islands 1B2, CTX-3C and (Madeira Arquipelago) CTX analogues 2 , Hawaii3, 4,5,6, Haiti7, from Carribean or St. Barthelemy, Indic waters2 8 Caribbean Sea , St Thomas, Carribean Sea9 Selvagens Islands C-CTX-110, CTX(Madeira Arquipelago) 1B2, CTX-3C and 2 , West Africa10 CTX analogues from Carribean or Indic waters2 11 Canary Islands , C-CTX-111 3, 12 Hawaii , St Thomas, Carribean Sea9 BarracudaC 13 The Bahamas , West C-CTX-115, 16 14 Africa , Florida Keys, USA15, French West Indies16, St. Barthelemy, Caribbean Sea8, 17, Guadeloupe17, French Polynesia18 Hervey Bay, CTX – positive19 Queensland, Australia19 California20 CTX – positive20 CTX – positive21

South Taiwan21

Method of detection UPLC/MS2, HPLC/MS7, TLC8, BSBA9, MGBA4, 9, MBA5, 6,8, S-EIA6, SPIA2, RIA4, 5, ELISA3, 5 , N2A1, 3, RBA7 LCMS/MS10, UPLC/MS2

LCMS/MS11, BSBA9, MGBA9, ELISA3, 12, N2A3, 11,12 Cat BA18, Chick BA17, MQBA18, MBA18, N2A13

TLC & MBA19 S-EIA20, SPIA20, N2A20 MBA21, N2A21

EelC 19

Chapter 1

Latin name (Common name) Gymnothorax funebris (Green Moray) Gymnothorax javanicus (Moray eel)

Lethrinus olivaceus (Longface emperor) Lethrinus miniatus (Trumpet emperor) Monotaxis grandoculis (Big eye bream) Mulloidichthys auriflamma (Goldstriped goatfish) Mulloidichthys martinicus (Yellow goatfish) Parupeneus insularis (Twosaddle goatfish) Conus spp. (Cone snails) Giant ClamH Tridacna sp. (Giant Clam) Hippopus hippopus (Giant Clam) Cephalopholis argus (Blue-spotted grouper, Roi) Cephalopholis miniata (Coral cod/Coral grouper) Epinephelus coioides (Orange-spotted grouper) Epinephelus lanceolatus (Giant grouper) Epinephelus microdon 20

Source St. Barthelemy, Caribbean Sea8 Tuamotu Archipelago and Tahiti (French Polynesia) 22, 23, 29, Tarawa, Republic of Kiribati, central Pacific Ocean26, Hawaii27

CTX (if detected) CTX – positive8

CTX-123, 25, CTX4B23, 25, CTX-225, CTX-325, P-CTX126, P-CTX-226, PCTX-326 and analogues of CTX 3C: 2,3dihydroxyCTX3C and 51hydroxyCTX3C28 Emperor breamC CTX - positive30

Nuku Hiva (Marquesas)30 French Polynesia18

CTX - positive18

French Polynesia 18

CTX - positive18

Hawaii

20

GoatfishC CTX - positive20

Method of detection TLC8, MBA8 HPLC/MS26, 28, HPLC/HNMR 23, 25, 29, TLC27, DLBA22, MBA 26, 27, 28

RBA30 Cat BA18, MQBA18, MBA18 Cat BA18 , MQBA18, MBA18 S-EIA20, SPIA20

St. Barthelemy, Caribbean Sea8

CTX – positive8

TLC8, MBA8

Nuku Hiva (Marquesas) 30

CTX - positive30

RBA30

Hawaii

31

GastropodC CTX - positive31

New Caledonia, French Polynesia32 Rep. of Vanuatu32

CTX - positive32

MBA32, N2A32, RBA32

CTX - positive32

N2A32, RBA32

GrouperC CTX – positive3,

Nuku Hiva (Marquesas)30, Hawaii3, French Polynesia18 Fiji33, 34, Arafura Sea, Australia35

Ciguatect ®31

18, 30

Cat BA18, MQBA18, MBA18, ELISA3, N2A3, RBA30

P-CTX-133, 34,35

HPLC/MS35, MBA35, N2A33, 34

Hong Kong36

CTX - positive36

MBA36

Hong Kong37

CTX - positive37

MBA37

French Polynesia18

CTX - positive18

Cat BA18, MQBA18,

Chapter 1

Latin name (Common name) (Marble grouper) Epinephelus mystacinus (Misty grouper) Epinephelus morio (Red grouper) Epinephelus sp. Mycteroperca bonaci (Black grouper) Mycteroperca prionura (Sawtail grouper) Mycteroperca sp. Mycteroperca venenosa (Yellowfin grouper) Plectropomus areolatus (Squaretail coral grouper) Plectropomus laevis (Blacksaddled coral grouper) Plectropomus leopardus (Coral trout/leopard coral grouper) Plectropomus spp. (Coral trout) Serranidae

Variola albimarginata (Lyretail) Pomadasys maculatus (Blotched javelin)

Bodianus bilunulatus (Tarry hogfish (a'awa)) Bodianus rufus (Spanish hogfish) Bodianus sp. Caranx ignobilis (Giant trevally (ulua))

Source

CTX (if detected)

Method of detection 9

MBA18 BSBA9, MGBA9

St Thomas, Carribean Sea9

CTX - positive

St. Barthelemy, Caribbean Sea8 Baja California, Mexico38 Key Largo, Florida, USA34 Baja California, Mexico39

CTX - positive8

TLC8, MBA8

CTX - positive38

MBA38

C-CTX-134, CCTX-234 CTX-139

LCMS/MS34, N2A34

Baja California, Mexico38 Guadeloupe and St. Barthelemy, Caribbean Sea17 Hong Kong36

CTX - positive38

MBA38

CTX - positive17

Chick BA17

CTX – positive36

MBA36

Hong Kong40

CTX - positive40

MBA40

French Polynesia, Tubuai (Australes)30, Hong Kong36, Tahiti41, French Polynesia18 Great Barrier Reef, Australia42 French West Indies43

CTX – positive18,

Cat BA18, MQBA18, MBA18, 36, 41, RBA30

Hong Kong40

Platypus Bay, Queensland, Australia42 Hawaii6

30, 36, 41

CTX-142, CTX242, CTX-342 C-CTX-143, CCTX-2 and isomers43, CTX congeners, other compounds43 CTX - positive40

GruntC CTX-142, CTX242, CTX-342 HogfishC CTX – positive6

St. Barthelemy, CTX - positive8 8 Caribbean Sea Hawaii20 CTX - positive20 Jacks and ScadsC French Polynesia, CTX – positive6,30 30 Tubuai (Australes) ,

HPLC/MS39, MBA39

HPLC/MS42, MBA42 MBA43

MBA40 HPLC/MS42, MBA42

MBA6, S-EIA6 TLC8, MBA8 SPIA20 MBA6, S-EIA6, RBA30

21

Chapter 1

Latin name (Common name)

Caranx latus (Horse-eye jack)

Caranx lugubris (Black jack) Caranx melampygus (Bluefin trevally) Caranx papuensis (Brassy trevally) Caranx sp. (Trevally (ulua, papio)) Cnidaria sp. Scomberomorus cavalla (King mackerel “Coronado” (Kingfish)) Scomberomorus commerson (Spanish mackerel)

Crenimugil crenilabis (Fringelip mullet) Liza vaigiensis (Thinlip grey mullet) Oplegnathus punctatus (Spotted knifejaw) Chlorurus frontalis (Pacific slopehead parrotfish) Chlorurus microrhinos (Steephead parrotfish) Scarus altipinnis (Filament-finned parrotfish) 22

Source St. Barthelemy, Caribbean Sea8 French West Indies43, St. Barthelemy, Caribbean Sea44, 45, The Bahamas46, St Thomas, Carribean Sea9 French West Indies43

Nuku Hiva (Marquesas)30, French Polynesia18 French Polynesia, Tubuai (Australes) 30 Hawaii6,20

CTX (if detected)

Method of detection

12 CTXs (inc CCTX-1, C-CTX1a, C-CTX-2)43, C-CTX-144, 45 and C-CTX-244, 45

HPLC/MS43,44,45, BSBA9, Cat BA46, MGBA9, MBA43,44

C-CTX-1 and isomers, CTX congeners43 CTX – positive18,30

MBA43

CTX - positive30

RBA30

CTX - positive6,20

MBA6, S-EIA6,20 SPIA6,20

JellyfishO American Samoa47 CTX - positive47 MackeralO 34 Florida, USA , St. C-CTX-134, CBarthelemy, Caribbean CTX-234 Sea8, 17, Guadeloupe17 Hervey Bay, CTX-142, CTXQueensland, 242, CTX-342 Australia19, Hervey Bay, Queensland, Australia48 MulletO Nuku Hiva CTX - positive30 (Marquesas)30, French Polynesia18 Nuku Hiva CTX - positive30 30 (Marquesas) KnifejawO 49 Miyazaki, Japan CTX-3C49

Cat BA18, MQBA18, MBA18, RBA30

SPIA47 LCMS/MS34, TLC8, Chick BA17, MBA8, N2A34 HPLC/MS42, TLC48, MBA19,42,48

MQBA18, MBA18, RBA30 RBA30 HPLC/MS49

ParrotfishH French Polynesia, CTX - positive30 30 Tubuai (Australes)

RBA30

French Polynesia, Tubuai (Australes) 30

CTX - positive30

RBA30

French Polynesia, Tubuai (Australes) 30

CTX - positive30

RBA30

Chapter 1

Latin name (Common name) Scarus ghobban (Blue-barred parrotfish) Scarus gibbus (Heavy beak parrotfish)

CTX (if detected) CTX - positive30

Method of detection

CTX-4A50

HPLC/HNMR50, MQBA, MBA 18, 41, 50

Scarus jonesi

French Polynesia50 (Satake et al., 1996), Tahiti41, French Polynesia18 French Polynesia18

CTX - positive18

Scarus rubroviolaceus (Ember parrotfish)

Nuku Hiva (Marquesas) 30

CTX - positive30

Cat BA18, MQBA18, MBA18 RBA30

Siganus rivulatus (Marbled spinefoot) Farmed salmon Kyphosus cinerascens (Blue sea chub)

Holothuria spp. Monachus schauinslandi (Hawaiian monk seal) Aphareus furca (Black forktail snapper (wahanui)) Aprion virescens (Bluge green snapper) Lutjanus argentimaculatus (Mangrove red snapper) Lutjanus bohar (Two spot red snapper (red bass))

Lutjanus buccanella (Blackfin snapper) Lutjanus gibbus (Humpback red snapper)

Source French Polynesia, Tubuai (Australes) 30

Eastern Mediterranean51

RabbitfishH CTX - positive51

SalmonO Chile CTX - positive52 Sea chubO French Polynesia, CTX – positive30 Tubuai (Australes) 30, Nuku Hiva (Marquesas) 30 Sea cucumberH Hawaii53 CTX - positive53 C Seal Hawaii54 P-CTX-3C54 52

Hawaii

20

SnapperC CTX - positive20

RBA30

Cigua-check ®51 SPIA52 RBA30

Ciguatect ®53 LCMS/MS54, N2A54

S-EIA20, SPIA20

French Polynesia18

CTX - positive18

Hong Kong40

CTX - positive40

Republic of Mauritius55, Minamitorishima (Marcus) Island, Japan49, French Polynesia, Tubuai (Australes)30 ,Nuku Hiva (Marquesas)30, Hawaii20, French Polynesia18 St Croix, US Virgin Islands56 Nuku Hiva (Marquesas) 30, French Polynesia18

I-CTX-155, CTX1B49

HPLC/MS55, Cat BA18, MGBA55, MQBA18, MBA18,55, RBA30

CTX - positive56

TLC56, MBA56

CTX – positive 18,

MQBA18, MBA18, RBA30

30

Cat BA18, MQBA18, MBA18 MBA40

23

Chapter 1

Latin name (Common name) Lutjanus griseus (Grey snapper)

Source

Lutjanus kasmira (Bluestripe snapper (taape)) Lutjanus monostigma (One-spot Snapper) Lutjanus sebae (Red emperor)

Hawaii6

Lutjanus spp. (Snapper)

Lutjanus stellatus (Star snapper) Myripristis kuntee (Epaulette Soldierfish (squirrelfish)) Sargocentron spiniferum (Sabre squirrelfish) Ophiocoma spp. (Ophiuroids (brittle stars)) Acanthurus dussumieri (Dussumier's surgeonfish (palani)) Acanthurus nigroris (Bluelined surgeonfish (maiko)) Acanthurus olivaceus (Orangeband surgeonfish (naenae)) Acanthurus sp. Acanthurus xanthopterus (Yellowfin surgeonfish) Ctenochaetus striatus (Striped Bristletooth) Malacanthus plumieri (Sand tilefish) Gymnosarda unicolor 24

French West Indies43

Nuku Hiva (Marquesas) 30 Republic of Mauritius (Nazareth, Saya de Malha, Soudan) 55 Antigua20, Okinawa, Japan49, West Africa14, Baja California, Mexico58, St Thomas, Carribean Sea9 Hong Kong40

CTX (if detected) C-CTX-1 and isomers43, CTX congeners43 CTX - positive6

Method of detection

CTX - positive30

RBA30

I-CTX55, I-CTX255, I-CTX-355, ICTX-455 CTX-1B49

HPLC/MS55, HPLC/MS/RLB55, MGBA55, MBA55 HPLC/MS49, BSBA9, MGBA9, MBA58, SEIA20, SPIA20, N2A14

CTX - positive40

MBA40

Squirrelfish and SoldeirfishC Hawaii6 CTX – positive6

Nuku Hiva (Marquesas) 30

CTX - positive30 StarfishO CTX - positive53

MBA43 MBA6, S-EIA6, SPIA6

MBA6, S-EIA6, SPIA6 RBA30

Hawaii

53

Hawaii

6

SurgeonfishH CTX - positive6

MBA6, S-EIA6

Hawaii6

CTX - positive6

MBA6, S-EIA6

Hawaii6

CTX - positive6

MBA6, S-EIA6

Hawaii20 Nuku Hiva (Marquesas) 30

CTX - positive20 CTX - positive30

S-EIA20, SPIA20 RBA30

Nuku Hiva CTX – positive30, 30 57 57 (Marquesas) , Tahiti TilefishC St. Barthelemy, CTX - positive8 8 Caribbean Sea TunaC Nuku Hiva CTX - positive30

Ciguatect ®53

RBA30 TLC8, MBA8 Cat BA18, MQBA18,

Chapter 1

Latin name (Common name) (Dogtooth tuna)

Naso brachycentron (Humpback unicornfish) Naso brevirostris (Spotted unicornfish) Naso hexacanthus (Sleek unicornfish) Naso lituratus (Orangespine unicornfish) Naso unicornis (Bluespine unicornfish)

Source

CTX (if detected)

(Marquesas) 30, French Polynesia18 UnicornfishC Nuku Hiva CTX - positive30 30 (Marquesas)

Method of detection MBA18, RBA30 RBA30

Nuku Hiva (Marquesas) 30 Nuku Hiva (Marquesas) 30 Nuku Hiva 18, 30 (Marquesas)

CTX - positive30

RBA30

CTX - positive30

RBA30

CTX – positive 18, 30

Cat BA18, MQBA18, MBA18, RBA30

Nuku Hiva (Marquesas) 30

CTX - positive30

RBA30

WrasseC Cheilinus undulatus French Polynesia , CTX – positive 18, Cat BA18, MQBA18, 36, 37 (Humphead Wrasse) Hong Kong36, 37 MBA18, 36, 37 30 Coris aygula French Polynesia, CTX - positive RBA30 30 (Clown coris) Tubuai (Australes) Semicossyphus sp. Baja California, CTX - positive38 MBA38 38 Mexico The abbreviations are: Typical feeding behaviour C = Carnivore, H = herbivore, O = omnivore; LC-MS/MS, Liquid chromatography tandem mass spectrometry; UPLC/MS, Ultra performance liquid chromatography/mass spectrometry; HPLC/MS, High performance liquid chromatography/mass spectrometry; HPLC/HNMR, High performance liquid chromatography/H Nuclear Magnetic Resonance; HPLC/MS/RLB, High performance liquid chromatography/mass spectrometry/radio ligand binding; TLC , Thin layer chromatography; BSBA, Brine shrimp bioassay; DLBA, Diptera larvae bioassay; MGBA, Mongoose bioassay; MQBA, Mosquito bioassay; MBA, Mouse bioassay, SEIA, Stick enzyme immunoassay; SPIA, Solid phase immunoassay; RIA, Radioimmunoassay; ELISA, enzyme-linked immunosorbent assay; N2A, Neuroblastoma cytotoxicity assays; RBA, receptor-binding assay; MA, Membrane assay; BA, bioassay; The references are: 1. (Caillaud et al., 2012) 2. (Otero et al., 2010) 3. (Campora et al., 2008) 4. (Hokama et al., 1977) 5. (Hokama et al., 1983) 6. (Hokama, 1985) 7. (Poli et al., 1997) 8. (Vernoux and Elandaloussi, 1986) 9. (Granade et al., 1976) 10. (Boada et al., 2010) 11. (PérezArellano et al., 2005) 12. (Campora et al., 2010) 13. (O'Toole et al., 2012) 14. (Bienfang et al., 2008) 15. (Dechraoui et al., 2005) 16. (Pottier et al., 2003) 17. (Pottier et al., 2001) 18. (Bagnis et al., 1987) 19. (Lewis and Endean, 1984) 20. (Hokama, 1990) 21. (Hung et al., 2005) 22. (Labrousse and Matile, 1996) 23. (Murata et al., 1990) 24. (Legrand et al., 1989) 25. (Lewis et al., 1991) 26. (Lewis and Jones, 1997) 27. (Scheuer et al., 1967) 28. (Satake et al., 1998) 29. (Legrand et al., 1989) 30. (Darius et al., 2007) 31. (Park et al., 2001) 32. (Laurent et al., 2012) 33. (Arnett and Lim, 2007) 34.(Dickey, 2008) 35. (Lucas et al., 1997) 36. (Wong et al., 2005) 37. (Wong et al., 2009) 38. (Lechuga‐Devéze and Sierra‐Beltrán, 1995) 39. (Sierra-Beltran et al., 1997) 40. (Wong et al., 2008) 41. (Pompon and Bagnis, 1984) 42. (Lewis and Sellin, 1992) 43. (Pottier et al., 2002a, Pottier et al., 2002b) 44. (Vernoux and Lewis, 1997) 45. (Lewis et al., 1998) 46. (Larson and Rothman, 1967) 47. (Zlotnick et al., 1995) 48. (Endean et al., 1993) 49. (Yogi et al., 2011) 50. (Satake et al., 1996) 51. (Bentur and Spanier, 2007) 52. (Ebesu et al., 1994, Dinubile and Hokama, 1995) 53. (Park et al., 2001) 54. (Bottein et al., 2011) 55. (Hamilton et al., 2002a, Hamilton et al., 2002b) 56. (Hoffman et al., 1983) 57. (Bagnis et al., 1985) 58. (Parrilla-Cerrillo et al., 1993) 18

25

Chapter 1

1.7

Aims of the study With the recognition of Gambierdiscus as the main cause of CFP, major advances have

been made in the study of species in this genus. Concurrently, new questions and challenges have also been raised. The aims of this work were to:

1) Design and test new methods based on next generation sequencing to accurately identify dinoflagellate species present in benthic habitats, 2) Determine the relationship between species presence and the presence of toxins (CTXs and MTXs) associated with Gambierdiscus spp. in an environmental sample set. 3) Investigate experimentally the accumulation potential of toxins associated with a species of Gambierdiscus in carnivorous fish. 4) Investigate the genetic basis of CTXs and MTXs in species of Gambierdiscus through extensive transcriptomic and genomic sequencing.

26

Chapter 2

Chapter 2 Cob gene pyrosequencing enables characterisation of benthic dinoflagellate diversity and biogeography 2.1

Summary Dinoflagellates in marine benthic habitats living epiphytically on macroalgae are an

important but highly understudied group of protists. Many produce toxins that can have severe economic impacts on marine-based economies, and improved monitoring tools are required to enhance the management of toxin related hazards. This study analysed the distribution and diversity of epibenthic dinoflagellates inhabiting eight sites in Cocos (Keeling) Islands (CKI), Papua New Guinea (PNG), Broome and Exmouth, Western Australia. We used pyrosequencing approaches based on two DNA barcoding marker genes; 18S rRNA and mitochondrial cytochrome b (cob), and compared these to an approach based on clone libraries (197 sequences) using the cob gene. Dinoflagellate sequences accounted for 133 (64 unique OTU) out of 10529 18S rRNA gene sequences obtained from all samples. However, using the dinoflagellate specific assay targeting the cob gene marker, the analysis obtained 9748 (1217 unique OTU) dinoflagellate sequences from the same environmental samples, providing the largest, to date, set of dinoflagellate cob gene sequences and reliable estimates of total dinoflagellate richness within the samples and biogeographic comparisons between samples. This study also reports the presence of potentially toxic species of the genera Gambierdiscus, Ostreopsis, Coolia, Prorocentrum and Amphidinium from the above-mentioned geographical regions. This chapter is published as “Kohli, G. S., Neilan, B. A., Brown, M. V., Hoppenrath, M. & Murray, S. A. 2013. Cob gene pyrosequencing enables characterisation of benthic dinoflagellate diversity and biogeography. Environmental Microbiology. In press, available online DOI: 10.1111/1462-2920.12275”

2.2

Introduction More than 2000 extant species of dinoflagellates, a group of microbial eukaryotes, are

known (Taylor et al., 2008). However, our knowledge of their diversity and global distribution in specific environments is far from comprehensive. Dinoflagellates in marine benthic habitats are an important but understudied group of protists. They are widely distributed in both tropical and temperate environments, living epibenthically on dead corals, sediments and numerous macroalgae (Murray, 2010). Some benthic taxa, such as species of Gambierdiscus, Prorocentrum and Ostreopsis produce secondary metabolites (toxins) that are linked to seafoodrelated toxin diseases worldwide (Murakami et al., 1982, Usami et al., 1995, Chinain et al., 2010, Kalaitzis et al., 2010). For example, ciguatera fish poisoning affects between 50,000 and 27

Chapter 2

500,000 people per year (Fleming et al., 1998) and is caused by the ingestion of fish that have accumulated toxins produced by Gambierdiscus spp. Similarly, Ostreopsis spp. produces the secondary metabolite palytoxin, which is highly toxic, found throughout certain reef-associated food webs and causes human illness and death (Usami et al., 1995, Deeds and Schwartz, 2010). Prorocentrum lima and related species of Prorocentrum are producers of okadaic acid (Murakami et al., 1982, MacKenzie et al., 2005), a toxin which bioaccumulates in shellfish and causes diarrhetic shellfish poisoning following consumption (Quilliam and Wright, 1995). Recently, a thecate peridinioid dinoflagellate was identified as a potent producer of pinnatoxin, a toxin which accumulates in oysters (Smith et al., 2011). With reports of a 60% increase in ciguatera fish poisoning in the Pacific islands over the last decade, despite being highly underreported (Skinner et al., 2011) , it is important to understand the biogeography and distribution of interstitial (sand-dwelling) and epiphytic benthic dinoflagellates worldwide. An increased knowledge of their diversity, ecology and distribution would enable improved monitoring and management of toxin related hazards, and would allow the development of a baseline in order to determine the effect of future changes to ocean temperature, reef condition, and currents. Most of the studies which describe benthic dinoflagellate diversity have investigated it using microscopic observation of live or fixed field samples or have characterised single-cells and cultures using molecular genetic techniques, and more recently, a combination of genetics and microscopy has been used (Taylor, 1979a, Fukuyo, 1981, Carlson and Trindall, 1985, Ballantine et al., 1988, Faust and Balech, 1993, Morton and Faust, 1997, Pearse et al., 2001, Okolodkov et al., 2007, Rhodes et al., 2010). While these methods are accurate, they do not generally provide information about the total dinoflagellate diversity in a particular area. For example, one of the most comprehensive taxonomic studies of epibenthic dinoflagellates to date revealed 12 genera and 39 species, including unidentified taxa (Turquet et al., 1998), however, there are indications that many cryptic species of dinoflagellates are present in field samples (Stern et al., 2010). Also, microscopic observations are time consuming and require taxonomic expertise. In the last decade, numerous attempts have been made to describe the diversity of microbial eukaryotes directly from environmental samples based on sequencing of ribosomal RNA (rRNA) genes via cloning (Moon-van der Staay et al., 2001, López-García et al., 2003), metagenomics (Cuvelier et al., 2010) and pyrosequencing (Brown et al., 2009, Edgcomb et al., 2011, Steven et al., 2012). However, these general approaches are unlikely to retrieve the full range of benthic dinoflagellate species diversity due to: 1) the use of primers which amplify all eukaryotes in environmental samples and are biased towards certain taxonomic groups (Potvin and Lovejoy, 2009) or highly abundant organisms, can “drown out” the signal of low abundance organisms such as dinoflagellates if they constitute a very small proportion of the community 28

Chapter 2

and 2) use of generic statistical analysis methods which have been primarily developed for estimating prokaryotic diversity and don’t take into account the huge range of rRNA operons in dinoflagellate genomes. These deficiencies highlight the need to develop molecular tools for the specific purpose of describing dinoflagellate diversity, such as PCR primer sets that can selectively amplify only dinoflagellate sequences directly from environmental samples. In 2006, a dinoflagellate-specific 18S rRNA primer set was developed for describing dinoflagellate community structure from environmental samples (Lin et al., 2006) . This method was used to amplify most of the dinoflagellate genera as tested in that study, however its product size (1.6kb) is not suitable for developing pyrosequencing assays. Recently, an internal transcribed spacer (ITS) region based on a barcoding technique has revealed hidden diversity in cultured dinoflagellates (Stern et al., 2012). However, the potential of a method based on a region of ITS rRNA to be used as an environmental barcoding marker is limited, due to the presence of many gene paralogues, the potential for unidentifiable chimaeras and priming across taxa (Stern et al., 2012). Other studies used the mitochondrial genes cob (Lin et al., 2009) and cytochrome c oxidase (cox) (Stern et al., 2010) to determine diversity of dinoflagellates directly from environmental DNA samples. Though these studies revealed a reasonable amount of dinoflagellate diversity, they used conventional PCR-based cloning and Sanger sequencing with results limited to < 200-800 clone sequences (Lin et al., 2009, Stern et al., 2010). However, with the availability of next generation sequencing, sequencing depth has been increased exponentially and has yet to be used to reveal dinoflagellate diversity in benthic systems. This study describes the diversity of epibenthic dinoflagellates in several different geographical locations in the Australasian region. Conventional barcoding molecular techniques targeting the cob and 18S rRNA genes were compared with more advanced Tag-Encoded FLX 454-Pyrosequencing. The new pyrosequencing assay used a cob-based dinoflagellate-specific primer set (Lin et al., 2009), which produces short PCR products that provide good taxon resolution to identify species. Using this assay, the study isolated the largest, to date, set of epibenthic dinoflagellate cob gene sequences that provide reliable estimates of total dinoflagellate richness within our samples.

2.3 2.3.1

Materials and methods Sample collection Samples were collected from 8 different sites from 4 different tropical regions around

Australasia (Figure 2.1, Table 2.1). Except Broome, all sampling sites were coral reefs. As dinoflagellates are known to live epibenthically on numerous macroalgae, specimens of Padina sp. and Sargassum sp. were collected (Bomber et al., 1989a, Holmes, 1998). These genera of macroalgae were chosen, as they were present at every site. The identity of the species of 29

Chapter 2

Padina and Sargassum were obtained following the next generation pyrosequencing. About 750 g of macroalgae was collected from approximately 1 m deep water at low tide and briefly placed in plastic bags containing 200-500 ml of ambient seawater. They were shaken vigorously for 5 mins to detach the epiphytic dinoflagellates from the macroalgae. The seawater was collected in a separate container immediately. This seawater was passed through >100 µm mesh filter to remove any remaining larger fauna and debris. From this, approximately 50 ml of each sample was filtered using a 3 µm filter (Merck Millipore®, Billerica, MA). Cells were washed from the filters using about 5 ml RNAlater (Ambion®, Austin, TX) for preservation and stored at 4 °C.

Figure 2.1: Map showing location of different sampling sites: (B) Broome, Western Australia. (E) Exmouth, Western Australia. (C) Cocos (Keeling) Islands. (P1) Tawali, Milne Bay, Papua New Guinea. (P2) Lion Island, Port Moresby, Papua New Guinea. Shaded areas in the map indicate sampling sites.

2.3.2

Culturing To enhance the cob reference database the following dinoflagellate species were obtained

from the Cawthron Institute Culture Collection (Nelson, New Zealand): Gambierdiscus australes CAWD149, isolated from the Cook Islands (21°13’S 159°46’W) (Rhodes et al., 2010), Ostreopsis siamensis CAWD173, isolated from New Zealand (34°58’S 173°16’E) , 30

Chapter 2

Ostreopsis ovata CAWD174 isolated from Cook Islands (21°14’S 159°47’W), Coolia monotis CAWD98 isolated from New Zealand (41°11’S, 173°20’E) and Prorocentrum compressum CAWD30 isolated from New Zealand (38°10’S, 174°41’E) . All cultures were grown at 25 °C, in 12:12 h light:dark cycle with 60 μmol m−2 s−1 light intensity in f/2 medium. Five hundred ml cultures were harvested once the cells reached early stationary phase and pellets were preserved in 2 ml RNAlater at 4 °C for DNA extraction. Table 2.1: List of sampling sites, the macroalgae the samples were collected from, their water temperature at the time of sample collection and the type of analysis done with each sample. Sample ID

Location (Latitude, Longitude), Water Temperature (°C)

Macroalgae

B1

Cable Beach, Broome, WA, Australia (17°55'34.34"S, 122°12'31.00"E), 25 Gantheaume Point, Broome, WA, Australia (17°58'24.02"S , 122°10'43.46"E ), 25 Town Beach, Broome, WA, Australia (17°58'23.31"S, 122°14'9.68"E), 25 Gantheaume Point, Broome, WA, Australia (17°58'24.02"S , 122°10'43.46"E ), 25 South Lagoon, Cocos (Keeling) Islands (12°9'43.76"S, 96°50'59.15"E) South Island, Cocos (Keeling) Islands (12°9'45.86"S, 96°53'38.36"E) Town Beach, Ningaloo reef, Exmouth, WA, Australia (21°56'21.37"S, 114° 8'26.00"E), 26 Lion Island, Port Moresby, Papua New Guinea (9°32'9.11"S, 147°16'34.32"E) 28 Tawali, Milne Bay, Papua New Guinea (10°19'54.57"S, 150°30'42.83"E), 28

Padina sp.

Diversity analysis PCRcTEFP Cloning yes yes

Padina sp.

yes

yes

yes

Padina sp.

yes

yes

yes

Sargassum sp.

no

yes

yes

Padina sp.

yes

yes

yes

Sargassum sp.

no

yes

yes

Padina sp.

yes

yes

yes

Padina sp.

yes

no

no

Padina sp.

yes

no

no

B2

B3

B4

C1

C2

E1

P1

P2

2.3.3

rTEFP yes

DNA extraction A 50 mg pellet of the preserved samples/culture cell pellet was collected via

centrifugation (12000 × g, 5 min) and used to extract total genomic DNA via FastDNA® Spin kit for Soil (MP Biomedicals, Solon, OH). The manufacturer’s protocol was followed and samples were stored at -20°C until PCR amplification.

31

Chapter 2

2.3.4

PCR, construction and screening of dinoflagellate specific cob clone libraries All PCR reactions were performed in 25 μl reaction volumes containing 5 μl Taq

polymerase buffer, 2.5 μl GC Melt (Clontech, Mountain View, CA), 0.2 mM deoxynucleotide triphosphates, 10 pmol each of the forward and reverse primers, between 10 and 100 ng genomic DNA, and 0.5 U of Advantage®-GC 2 Taq Polymerase (Clontech). PCR for cob was performed using primers dinocob4f and dinocob6r (Lin et al., 2009) and the following cycle conditions: initial denaturation at 96°C for 3 min followed by 32 cycles of denaturation at 94°C for 20 s, primer annealing at 55°C for 20 s and an extension at 68°C for 1 min followed by a final extension step at 68°C for 3 min. The PCR amplicons were cloned using TOPO TA cloning kit (Invitrogen, Carlsbad, CA) according to the manufacturer’s protocol. More than 25 clones were sequenced for each sample. Sequencing was performed using the PRISM BigDye cycle sequencing system and a model 373 sequencer (Applied Biosystems, Foster City, CA) at the Ramaciotti Centre for Gene Function Analysis (Sydney, Australia). Sequence data were trimmed and analysed using Geneious® software (Kearse et al., 2012) while identity/similarity values to other sequences were determined using BLAST in conjugation with the National Center for Biotechnology Information (NIH, Bethesda, MD). Finally, phylogenetic analysis combining reference sequences and sequences obtained during this study was carried out to determine the diversity of dinoflagellates in each sample.

2.3.5

Tag-Encoded FLX 454-Pyrosequencing Cob Tag-Encoded FLX 454-Pyrosequencing (cTEFP) and 18S rRNA gene ribosomal

Tag-Encoded FLX 454-Pyrosequencing (rTEFP) was performed using primers designed in previous studies. For cTEFP assay, the primer pair dinocob4f (AGCATTTATGGGTTA TGTNTTACCTTT) and dinocob6r (ATTGGCATAGGAAATACCATTCAGG) (Lin et al., 2009), which amplifies a 441 bp barcoded region of the cob gene, was tailored for pyrosequencing. This was done by adding a biotin and fusion linker sequence at the 5’ end of the forward primer and a fusion linker and a proprietary 12 bp barcode sequence at the 5’ end of the reverse primer. Sequencing was performed at the 3’ end of the product rather than the 5’ end using the reverse primer. For rTEFP assay the primer pair euk-A7F (AACCTGGTTGAT CCTGCCAGT) (Medlin et al., 1988) and euk-570R (GCTATTGGAGCTGGAATTAC) (Weekers et al., 1994), which amplifies a 563bp region of the 18S rRNA gene, was tailored for pyrosequencing. This was done by adding a fusion linker and the 12 bp barcode sequence at the 5’ end of the euk-A7F forward primer and a biotin and fusion linker sequence at the 5’ end of the reverse primer. Sequencing was performed from the 5’ end of the product using the forward primer. PCRs and FLX-Titanium sequencing (Roche, Nutley, NJ) were carried out at the

32

Chapter 2

Research and Testing Laboratory (Lubbock, TX) according to their established protocols (Dowd et al., 2008).

2.3.6

cTEFP sequence analysis Trimming and selection of high quality reads was done using the MOTHUR software

package (Schloss et al., 2009). Sequences with any ambiguous base calls and/or shorter than 380bp in length were discarded. A local BLAST using the nr/nt database was performed and any sequence with the closest BLAST match (based on sequence percentage similarity) to a non-dinoflagellate cob sequences were discarded. Unique sequences were obtained using the unique.seqs and cluster.fragments command in MOTHUR. Chimeras were identified using chimera.uchime in MOTHUR and eliminated from the unique sequences list. After removing the chimeras all the sequences were combined with 46 reference sequences (Table 1, Appendix A) and phylogenetic analysis was carried out to identify and segregate sequences based on their taxonomic orders, which were Gonyaulacales, Peridiniales, Gymnodiniales, Prorocentrales, Suessiales and Dinophysiales. Sequences that could not be assigned to any dinoflagellate order based on phylogenetic analysis were marked as unknown sequences and not analysed any further. This divided the dataset from each sample into 6-subsample datasets based on the 6 orders. Sequences from each subsample were aligned and trimmed using MAFFT v6.814b (Katoh et al., 2002) in Geneious®. Coverage and number of unique sequences, based on percentage similarity, was calculated using the following commands in MOTHUR: unique.seqs, dist.seqs, cluster, summary.single (calc=nseqs-coverage-sobs). In MOTHUR, coverage calculator is based on Goods equation (Good, 1953). To determine the number of species/genera within each sample, an accurate identification of sequences based on percentage similarity could not be performed due to the limited availability of sequences to construct a comprehensive cob database. No single universal distance threshold could be established to define a distinct species or genus across the whole Dinophyceae class, as the diversity in this gene was strikingly different for some genera. To identify and estimate diversity at the genus level accurately, the minimum distance between different genera within each order was calculated, based on the percentage similarity of reference sequences. These were calculated as: Gonyaulacales (99%), Peridiniales (99%), Gymnodiniales (97%), Prorocentrales (distance between different species 99%) and Suessiales (96%). Due to sequencing error, inflation of diversity estimates at a 99% cut-off has been associated with pyrosequencing in prokaryotes (Kunin et al., 2010). To minimise this, sequences from all the seven samples belonging to a particular order were pooled together and sequences that occurred only once in this whole dataset, across seven samples at specific percentage similarity, were removed. For example, after pooling all the gonyaulacalean sequences together from all the samples, if a sequence was present once in multiple samples 33

Chapter 2

such as B1 and B2 it was not deleted. However, if the sequence was present in sample B1 once and not present in any other sample, it was deleted. A final phylogenetic analysis was performed using unique sequences from each order and 49 reference sequences (Table 1, Appendix A) to identify the genera present in each sample. Very limited information is available on the copy number of the cob gene in the mitochondrial genomes of dinoflagellates (Waller and Jackson, 2009, Jackson et al., 2012). Recent studies have shown that only very slight variations exist between different copies of cytochrome c oxidase 1 genes within the same species (Scrippsiella and Prorocentrum- 0.2% pairwise distances, Symbiodinium- 1.3% pairwise distances) (Stern et al., 2010). The cut off values of different OTUs used in the current study were 96-99% of sequence similarity, as described above. This range of percentage sequence similarity is higher than the diversity observed intra-genomically in the cytochrome c oxidase 1 gene.

2.3.7

rTEFP sequence analysis Sequences obtained via rTEFP were trimmed and unique sequences were selected using

the following commands in MOTHUR: trim.seqs, unique.seqs and cluster.fragments. On the selected sequences a local BLAST using the nr/nt database was performed and sequences with closest BLAST match to dinoflagellate SSU sequences were selected (based on sequence percentage similarity). These sequences were combined with 61 reference sequences (Table 2, Appendix A) and phylogenetic analysis was carried out for identification purposes.

2.3.8

Phylogenetic analysis All steps were performed in Geneious® software (Kearse et al., 2012). Sequences from

different datasets were aligned using MAFFT v6.814b (Katoh et al., 2002). Alignments were trimmed manually to ensure they spanned the same cob/18s rRNA region. After aligning the sequences, the best substitution model was determined using ModelTest (Posada and Crandall, 1998) and a maximum likelihood phylogenetic analysis was carried out using the program PhyML (Guindon et al., 2010) with 500 bootstraps.

2.4 2.4.1

Results Diversity of dinoflagellates via PCR, cob gene cloning and sequencing Three samples from Broome, two from PNG, and one each from CKI and Exmouth were

analysed via PCR and generating cob gene clone libraries. Sequences from all 197 clones were identified as cob genes representing various genera belonging to Dinophyceae, based on a high level of similarity in a BLAST search (Table 2.2). Phylogenetic analysis was performed to facilitate accurate identification of sequences (Figure 2.2).

34

Chapter 2

Figure 2.2: Phylogenetic analysis using maximum likelihood of cob sequences obtained from PCR based cob gene cloning and sequencing. Major orders of Dinophyceae were shown on the right. Bootstrap values were based on 500 replicates; the thickest branches denote bootstrap value of >90%, medium-thick branches values of 70-90% and thin branches values of 90%, medium-thick branches, 7090% and thin branches, 90%, medium-thick branches, 7090% and thin branches, 90%, mediumthick branches, 70-90% and thin branches, 90%, medium-thick branches values of 70-90% and thin branches values of 100-fold variation in toxicity, but also suggests a much broader range of environmental tolerances for potentially CFP-causing dinoflagellates. In Australia, Gambierdiscus is known to be present in the tropical waters of Queensland (QLD), with more than 1,400 cases of CFP reported in the past 15 years, including two fatalities (Gillespie et al., 1986, Stewart et al., 2010). In the past, Gambierdiscus has been reported from the tropical Great Barrier Reef (Flinders Reef, Arlington Reef, Hastings Reef, Heron Island) down to Platypus Bay, just north of Brisbane (25oS) (Gillespie et al., 1985, Hallegraeff, 1993, Holmes and Lewis, 1994). However, previous Australian toxicological studies (Holmes et al., 1990, Holmes and Lewis, 1994) do not allow for an unambiguous identification of the precise Gambierdiscus species involved, since no archived cultures or samples are available. Along the coast of New South Wales (NSW) there exist 31 estuaries, which are monitored fortnightly for occurrences of harmful planktonic microalgae as part of the state’s shellfish safety program. Ajani et al. (2013) reported the incidence of Gambierdiscus from five sites along the NSW coast in the period 2005-2009 (Camden Haven River, Wallis Lake, Tuross Lake, Wapengo Lagoon, Merimbula Lake and Womboyn River) (Ajani et al., 2013). Murray (2010) and Hallegraeff (2010) first drew attention to this unusual warmtemperate Gambierdiscus population, stretching as far as 37o south, with Gambierdiscus previously considered to be mostly a tropical species. The major aim of this study was to identify the Gambierdiscus species present in southern New South Wales estuaries. Light microscopy and 18S rRNA gene ribosomal Tag-Encoded FLX 454-Pyrosequencing (rTEFP) methods were applied to study the dinoflagellate community 51

Chapter 3

structure at three sites. Results revealed the presence of a single species of Gambierdiscus at all three sites. Detailed morphological studies via electron microscopy were carried out to confirm the identity of the Gambierdiscus species in the samples. In May 2013, a bloom of G. carpenteri was reported, and samples were taken to estimate cell densities. Bloom samples were also tested for the presence of CTXs and MTXs using Liquid chromatography- Mass spectrometry (LC-MS) analysis. The toxicity of the bloom sample extracts was also determined via mouse-bioassay.

3.3 3.3.1

Materials and methods Sample collection In May 2012 and 2013, samples were collected from various sites for rTEFP analysis, cell

density estimation and toxin analysis. Sample location, identification and analysis are listed in Table 3.1. Figure 3.1and Figure 3.2 detail the locations of the sampling sites. Several dinoflagellate species are known to live epibenthically on a variety of macroalgae (Bomber et al., 1989a, Holmes, 1998). Hence, during sampling the most abundant types of macroalgae at each of the sites were collected (Table 3.1). Various amounts of macroalgae (750 g for rTEFP analysis, 5-10 g for cell density estimates, 1 kg for toxin analysis) were collected from approximately 1 m deep water and briefly placed in a plastic bag containing ambient seawater (300-500 ml for rTEFP analysis, 5-30 ml for cell density estimates, 2 L for toxin analysis). The bag was shaken vigorously for 5 min. The liquid contents were then passed through a 104 µm mesh filter to remove larger fauna and debris. For rTEFP analysis, 50 ml of each sample was filtered using a 3 µm filter (Merck Millipore®, Billerica, MA) and cells were washed from the filters using 5 ml RNAlater (Ambion®, Austin, TX) for preservation and stored at 4°C for DNA extraction. For cell density estimates, the filtrate obtained after mesh filtration was preserved in Lugol’s solution and stored at 4°C for further analysis. For toxin analysis, a 1 ml sub-sample was taken for estimating the cell density and the filtrate was centrifuged at 1000 rpm for 10 min. All the samples were stored at 4°C until further analysis.

52

Chapter 3

Table 3.1: Sampling sites, the macroalgae the samples were collected from, the water temperature at the time of sample collection and the type of analysis done with each sample. Sampling site (latitude-longitude)

Wagonga Inlet

No. of samples collected (sample name) 1 (WG1)

Date Macroalgae collected (water temperature)

Analysis conducted

May, 2012 (16.5 °C) May, 2012 (17 °C) May, 2012 (17.5 °C)

Padina sp.

rTEFP analysis rTEFP analysis rTEFP analysis

Wapengo Lagoon

1 (WP1)

Merimbula Inlet (36°53'29.80"S 149°54'39.18"E) Merimbula Inlet (36°53'29.80"S 149°54'39.18"E) S1-mid (Merimbula Inlet, 36°53'29.80"S 149°54'39.18"E) S1-low (Merimbula Inlet, 36°53'29.80"S149°54'39.18"E) S2 (Merimbula Inlet, 36°53'50.47"S 149°55'0.45"E) S3 (Merimbula Inlet, 36°53'56.33"S 149°55'37.81"E) S4 (Merimbula Inlet, 36°53'46.28"S 149°54'36.87"E)

1 (MB1)

1 (MB2)

May, 2012 (17.5 °C)

Phyllospora sp.

rTEFP analysis

4 (S1-mid-N1 to N4)

May, 2013 (17 °C)

Phyllospora sp.

Abundance counts

4 (S1-low-N1 to N4)

May, 2013 (17 °C)

Phyllospora sp.

Abundance counts

4 (S2-N1 to N4)

May, 2013 (17 °C)

Zosteraa sp.

Abundance counts

4 (S3-N1 to N4)

May, 2013 (17 °C)

Phyllospora sp.

Abundance counts

5 (S4-N1 to N5)

May, 2013 (17 °C)

S4-N1 and N2 Phyllospora sp., S4-N3 and N4 – Ecklonia sp. S4-N5Phyllospora sp. and Ecklonia sp.

Abundance counts

Phyllospora sp. Phyllospora sp.

53

Chapter 3

Figure 3.1: Map of the Eastern coastline of Australia. It shows locations where sampling was carried out, i.e. Wagonga Inlet, Wapengo Lagoon and Merimbula Lake Inlet.

S1

Merimbula

S4

Merimbula Lake

S2

S3 Tasman Sea

Merimbula 50 m

Figure 3.2: Map of Merimbula Inlet. It shows different sampling sites (S1-S4) during the May 2013 bloom event. Dotted lines (-----) show the approximate position of one of the shellfish farming sites.

54

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3.3.2

DNA extraction, Tag-Encoded FLX 454-Pyrosequencing and sequence analysis Two samples from Merimbula Inlet and one sample each from Wagonga Inlet and

Wapengo Lagoon (Table 3.1, Figure 3.1) were collected as described above. A 50 mg cell pellet of the preserved samples was collected via centrifugation (12000 × g, 5 min) and used to extract total genomic DNA using FastDNA® Spin kit for Soil (MP Biomedicals, Solon, OH), according to the manufacturer’s protocol. The samples were stored at -20°C until rTEFP analysis. 18S rRNA gene ribosomal Tag-Encoded FLX 454-Pyrosequencing was performed according to the protocols described in section 2.3.5. Trimming and selection of high quality reads obtained by rTEFP analysis was carried out using the MOTHUR software package (Schloss et al., 2009). Sequences with any ambiguous base calls and/or shorter than 400 bp in length were discarded. BLAST using the nr/nt database was performed on the selected sequences and sequences with closest BLAST match to dinoflagellate SSU sequences were selected (based on sequence percentage similarity). BLAST was carried out via BLAST2GO pro software. These selected sequences were aligned and trimmed using MAFFT v6.814b (Katoh et al., 2002) in Geneious® (Biomatters Ltd., Auckland, New Zealand). Coverage and number of unique sequences, based on percentage similarity, was calculated using the following commands in MOTHUR: unique.seqs, dist.seqs, cluster, summary.single (calc=nseqs-coverage-sobs). In MOTHUR, the coverage calculator is based on the Goods equation (Good, 1953). Due to sequencing error, inflation of diversity estimates at a 99% sequence similarity cut-off has been associated with pyrosequencing in prokaryotes (Kunin et al., 2010). In this study a cut off value of 98% based on sequence similarity was used to determine the number of unique operational taxonomic units (OTUs). These unique sequences were combined with 108 reference sequences and phylogenetic analysis was carried out for identification purposes.

3.3.3

DNA extraction and PCR analysis for species identification From the 2012 field trip, a culture of Gambierdiscus sp. [maintained in f/2 media

(Guillard and Ryther, 1962) at 25°C, 100 μmol m-2 s-1 photon flux, 12:12h L:D] was centrifuged (30 ml; 542 g, 15 min, RT) and genomic DNA was extracted from the resulting pellet using the UltraCleanTM Soil DNA isolation kit (MoBio, Carlsbad, CA). For the 2013 field trip, about 400500 individual Gambierdiscus cells were selected under microscopy. Cells were centrifuged and DNA was extracted using the FastDNA® Spin kit for Soil (MP Biomedicals, Solon, OH), as per the manufacturer’s protocol. For both samples, the D1–D3 fragment of the large subunit ribosomal DNA (LSU rDNA) gene was PCR amplified using the primers D1R-F (Scholin et al., 1996) and D3B-R (Nunn et al., 1996). PCR amplifications were carried out in 50 μl reaction 55

Chapter 3

volumes containing i-Taq 2 × PCR master mix (Intron, Gyeonggi-do, Korea) or ImmoMixTM (Bioline, Australia), 0.4 mM of forward and reverse primers and 50-150 ng of template DNA. Thermocycling conditions consisted of an initial denaturing step of 95°C for 5 min, followed by 32 cycles of 94°C for 30 s, 60°C for 2 min, and a final extension of 72°C for 6 min. PCR products were purified with AxyPrepTM PCR clean-up kit (Axygen Biosciences, Tewksbury, MA) and sequenced in both directions using BigDye Terminator v 3.1 Cycle Sequencing Kit (Applied Biosystems, Carlsbad, CA) by external contractors (Genetic Analysis Services, University of Otago, Dunedin, New Zealand and Macrogen Inc., Seoul, Korea). Sequence chromatograms were examined visually and any base-calling errors corrected manually using Geneious® v6.0.4 (Kearse et al., 2012). Both forward and reverse sequences were aligned and any ambiguities resolved by manual inspection.

3.3.4

Phylogenetic analysis All steps were performed in Geneious® v6.0.4 (Kearse et al., 2012). Sequences from

different datasets were aligned using MAFFT v6.814b (Katoh et al., 2002). Alignments were trimmed manually to ensure they spanned the same 18S rRNA gene region. After aligning the sequences, the best substitution model was determined using ModelTest (Posada and Crandall, 1998) and a maximum likelihood phylogenetic analysis was carried out using the program PhyML (Guindon et al., 2010) with 500 bootstrap resampling events.

3.3.5

Gambierdiscus population density and analysis Macroalgal samples from four locations around Merimbula inlet were collected (Table

3.1, Figure 3.2) in May 2013. At location S1, the samples were collected at two different time points, at mid tide and low tide. At each location, 4-5 macroalgal samples were collected (Figure 3.2) as described in section 3.3.1. For Gambierdiscus cell density estimates, each sample was counted using a 1 ml Sedgewick Rafter counting chamber under a bright field microscope (Motic, Shanghai, China). Samples were counted in triplicate and an average cell count/gram of wet weight macroalgae were reported. As the macroalgae was freeze-dried for preservation purposes, a conversion factor of 0.19 was used to calculate the approximate wet weight of macroalgae from the recorded dry weight (Kraemer and Alberte, 1993, Fleming, 1995). A logarithmic graph to represent the estimated cell densities was generated using Prism 6 software package (GraphPad software, La Jolla, CA).

3.3.6

Electron microscopy For scanning electron microscopy (SEM), live cells were gently filtered onto 1 µm

Nuclepore filters (Whatman, Maidstone, Kent, United Kingdom), rinsed in distilled water, and gently air-dried. Filters were mounted on stubs and sputter coated with gold-palladium. Cells were observed using a FEI Quanta 600 scanning electron microscope (FEI Company, Hillsboro, 56

Chapter 3

OR) at the University of Tasmania (Electron Microscopy and X-Ray Microanalysis, Central Science Laboratory) at 5-15 kV.

3.3.7

Toxin analysis via LC-MS and mouse bioassay In May 2013, a cell pellet of the Gambierdiscus bloom sample was collected in

Merimbula as described in section 2.1. The pellet was freeze-dried and later extracted using a previously standardised method for CTX (Rhodes et al., 2010) and MTX analysis (section 4.1.4). The LC-MS analysis was performed at the Cawthron Institute, Nelson, New Zealand. Samples were analysed on a Waters Acquity uPLC system coupled to a Quattro Premier triple quadrupole mass spectrometer equipped with a Z-Spray ion source (Waters, Milford, MA). LCMS/MS was performed with multiple-reaction monitoring for CTX-3b, CTX-3C, CTX-4A, CTX-4B, MTX and MTX-3. Mouse bioassays were conducted at Agri Research, Hamilton, New Zealand, using extracts from the same pellet that was sampled during the bloom event. The pellet was extracted using methanol and freeze-dried overnight. This extract was then taken up in 1% Tween 60 in saline and injected both intraperitoneally and orally in Swiss albino mice (body weight 18–20 g) at various dose levels. LD50 values were determined by the up and down method (OECD, 2006). All mouse experiments were approved by the Ruakura Animal Ethics Committee.

3.4 3.4.1

Results rTEFP analysis and LSU phylogenetic analysis Two samples from Merimbula and one each from Wagonga and Wapengo Lagoons

collected in 2012 were analysed by rTEFP. After screening 121,935 sequences, a total of 11,609 were more than 400 bp long and had known dinoflagellate sequences as their closest BLAST match in GenBank (nr/nt database). A total of 45 unique OTUs at 98% sequence similarity were identified using phylogenetic analysis, which represented 23 distinct genera (Figure 3.4). A list of all dinoflagellate genera found at each of the sampling sites is shown in Table 3.2. From a total of 11,609 dinoflagellate sequences, Gambierdiscus carpenteri sequences were the most abundant OTU in each dataset and comprised 88% (10221 OTUs) of the combined dataset for all four samples. To confirm the identity of Gambierdiscus species in the 2012 and 2013 samples, the D1D3 region of the LSU rDNA was obtained via PCR amplification. BLASTn and phylogenetic analysis revealed that the sequences were 99% similar to Gambierdiscus carpenteri (Figure 3.3).

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Table 3.2: Data obtained from rTEFP analysis. Sample ID Location

WG1 Wagonga, NSW

WP1 Wapengo, NSW

Total number of trimmed sequences Total number of dinoflagellate sequences Total number of dinoflagellate sequences above 400bp Coverage at 98% sequence similarity Total number of unique OTUs at 98% sequence similarity Closest identifiable sister group in the phylogeny (number of different unique sequences identified phylogenetically)

35572

58

25688

MB1 Merimbula, NSW 15599

MB2 Merimbula, NSW 45076

783

2176

1932

7736

675

1840

1683

7411

94.93

97.96

98.57

98.22

31

30

16

17

Gambierdiscus carpenteri (1), Alexandrium sp. (2), Coolia monotis (1), Coolia canariensis (1), Ostreopsis sp. (1), Amphidinium sp. (7), Amphidiniella sedentaria (1), Bysmatrum sp. (1), Prorocentrum concavum (1), Prorocentrum lima (1), Prorocentrum sp. (2), Warnowia sp. (1), Gymnodinium dorsalisulcum (1), Lepidodinium sp. (1), Symbiodinium sp. (3), Scrippsiella sp. (2), Durinskia sp. (1), Kryptoperidinium sp. (1), Galeidinium sp. (1), Azadinium sp. (1)

Gambierdiscus carpenteri (7), Alexandrium sp. (2), Coolia monotis (1), Coolia canariensis (1), Ostreopsis sp. (1), Ceratium sp. (1), Ceratocorys horrida (1), Amphidinium sp. (5), Prorocentrum concavum (1), Prorocentrum lima (1), Prorocentrum sp. (1), Warnowia sp. (1), Gymnodinium dorsalisulcum (1), Symbiodinium sp. (1), Scrippsiella sp. (1), Kryptoperidinium sp. (1), Galeidinium sp. (1), Takayama sp. (1), Dinophysis sp. (1)

Gambierdiscus carpenteri (7), Coolia monotis (1), Amphidinium sp. (3), Prorocentrum concavum (1), Prorocentrum lima (1), Prorocentrum sp. (1), Symbiodinium sp. (1), Scrippsiella sp. (1)

Gambierdiscus carpenteri (7), Coolia monotis (1), Ostreopsis sp. (1), Amphidinium sp. (2), Prorocentrum concavum (1), Prorocentrum lima (1), Prorocentrum sp. (1), Warnowia sp. (1), Scrippsiella sp. (1), Galeidinium sp. (1)

Chapter 3

Figure 3.3: Phylogenetic analysis using maximum likelihood of 28S rRNA gene sequences obtained from LSU analysis. Support values are bootstrap values based on 500 replicates. Reference sequence names are followed by the strain number, if available, and then accession

59

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60

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Figure 3.4: Phylogenetic analysis using maximum likelihood of 18S rRNA gene sequences obtained from rTEFP analysis. Support values are bootstrap values based on 500 replicates. Each unique OTU is represented by one representative sequence (highlighted in red colour). For example “MB2_32715 (WP1-1143, MB1-1398, MB2-6498)” indicates that the representative sequence “MB2_32715” was found 1143 times in sample WP1, 1398 times in sample MB1 and 6498 times in sample MB2. MB1 and MB2 are samples collected from Merimbula, WP1 from Wagonga and WG1 from Wagonga. Reference sequence names are followed by the strain number, if available, and then accession numbers.

3.4.2

Microscopy and scanning electron microscope analysis Figure 3.5 and Figure 3.6 illustrate morphological variation of epithecae and hypothecae

of combined field and cultured Merimbula Gambierdiscus cells. The shape of the 2’ plate was variable, ranging from hatchet-shaped (as in G.toxicus) to nearly rectangular as described for G. carpenteri (Figure 42 in Litaker et al. 2009). Plate overlap often prevented adequate resolution of the shape of the 2’ plate using SEM, while LM allowed for more consistent plate morphology assessment. Sometimes the same plate was overlapping or underlapping. Critically, none of our cells ever exhibited the thecal groove shown by G.carpenteri; nor the dorsal rostrum (Figure 41 in Litaker et al 2009). The 4” plate was asymmetrical (consistent with G.carpenteri), while the 1p plate was very broad, but sometimes with a pointed dorsal end (resembling G.toxicus). The lenticular cells most commonly showed transdiameters of 67-86 m, and comparable cell heights of 68-86 m, but in cultured material some cells as small as 62 or as large as 110 m were observed.

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B

C D A G

F

H E J

K I

M

N

L

Figure 3.5: Scanning electron microscopy micrographs of Gambierdiscus carpenteri from Merimbula Lake Inlet. (A and I) Apical view. Scale bar= 10μm; (E and J) Apico lateral view. Scale bar equals 10μm; (B, C, D, F, G and H) Apical view showing 2’ plate where the plate shape ranges from hatchet-shaped (as in G.toxicus) to nearly rectangular as described for G. carpenteri (Figure 42 in Litaker et al. 2009). Scale bar equals 10μm; (K-N) Apical pore plate. Scale bar equals 2μm.

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B A

D

C

E

F

G

Figure 3.6: Scanning electron microscopy micrographs of Gambierdiscus carpenteri from Merimbula Lake Inlet. (A, C, D and E) Antapical view, arrows in figure C show overlapping of plate boundry. Scale bar equals 5μm; (B) Antapico lateral view. Scale bar equals 5μm (F-G) Antapical view showing 1p plate. Scale bar equals 10μm.

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3.4.3

Gambierdiscus distribution and cell density estimates rTEFP analysis revealed that G. carpenteri was the most dominant microbial eukaryotic

species present in the samples collected in May 2012. This indicated the presence of a Gambierdiscus bloom, which was also evident from direct visual examination of live samples via microscopy at the time of sampling. In May 2013, another bloom of G. carpenteri was recorded in Merimbula and macroalgal samples were taken for abundance estimates. Samples taken from sites S1 and S4 (Figure 3.2) revealed maximum densities of 6,565 cells g-1wet weight algae and 8,255 cells g-1 wet weight algae (Figure 3.7), respectively. Samples taken outside the Merimbula inlet (sites S2 and S3, Figure 3.2) had lower cell densities, ranging between 0.4 and 56 cells g-1 wet weight algae (Figure 3.7). This indicated that the Gambierdiscus bloom was localised within the Merimbula inlet, including at sites less than 50 m away from aquaculture farms. Routine monitoring of plankton samples from New South Wales shellfish growing areas during 2005 to 2008 occasionally reported low concentrations of Gambierdiscus cells (50-700 cells/L, Brett, pers. comm.) at Camden Haven, Wallis Lake, Tuross Lake, Wagonga Inlet, Bermagui, Wapengo Lake, Merimbula, and Wonboyn Lake, exclusively in the December-June period (Ajani et al., 2013). This seasonality was confirmed by monthly monitoring of plankton net tows from Merimbula during a full annual cycle in 2012-3. The organism was abundant in May 2012, subsequently nearly absent throughout July to November, 2012, but then again became dominant throughout April to May 2013. Seasonal water temperatures in Merimbula ranged from 8-28oC (Figure 3.8), with Gambierdiscus thriving at temperatures above 16.5-17oC. Salinities in Merimbula most commonly ranged from 30-35 PSU, with occasional lows down to 20 and highs up to 40 PSU (Figure 3.8).

Figure 3.7: Gambierdiscus cell densities (cells g-1 wet weight macroalgae) from Merimbula in May 2013. S1-S4 refers to four different sites from where the samples were taken in Merimbula lake inlet. Each bar represents cell densities of individual sample collected at the particular site. Samples were counted in triplicate.

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Figure 3.8: Salinity (A) and temperature (B) records of Merimbula Inlet Lake from 2004 to 2013. The data was collected fortnightly during the phytoplankton monitoring programme run by New South Wales Food Authority.

3.4.4

Toxin analysis The Gambierdiscus bloom sample taken in May 2013 was estimated to contain a total of

3.7x107 Gambierdiscus cells. LC-MS analysis revealed the absence of CTX-3b, CTX-3C, CTX4A, CTX-4B, MTX and MTX-3 in the extract. However, the extract was toxic to mice via intraperitoneal injection with an LD50 dose of 2.4 mg/kg causing abdominal breathing, decreased respiration rates and death through respiratory paralysis. The necropsy analysis revealed fluid accumulation in the duodena of mice, however, no lesions in the glandular stomach were observed. Mice given a dose of 500 mg/kg via oral administration showed transient signs of toxicity, but survived. These symptoms are similar to those in assays conducted for the characterisation of MTX (Holmes et al., 1990, Holmes and Lewis, 1994) and for cell extracts obtained from cultures of Gambierdiscus belizeanus CCMP401, which produce MTXs (section 5.4.1).

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3.5 3.5.1

Discussion Dinoflagellate diversity as determined by rTEFP analysis The 18S rRNA gene phylogeny (Figure 3.4) of 108 reference dinoflagellate sequences

and 45 unique OTUs obtained from the rTEFP analysis, inferred the monophyletic origin of dinoflagellates, which is concurrent with previous 18S rRNA gene phylogenies (Murray et al., 2005, Orr et al., 2012). Among the major lineages of dinoflagellates, Gonyaucales, Suessiales, Peridiniales, and Dinophysiales appeared to be monophyletic while Gymnodiniales and Prorocentrales appeared to be polyphyletic. Recent, concatenated phylogenies inferred from 18S+5.8S+28S rRNA gene sequences also found Gymnodiniales to be polyphyletic, however, Prorocentrales appeared to be monophyletic (Orr et al., 2012). Results from the rTEFP analysis revealed the presence of a rich and diverse benthic community of dinoflagellates from all the three sites, Merimbula, Wapengo and and Wagonga. Surprisingly, sequences similar to Azadinium sp. were reported from Wagonga (Table 3.2). This potentially toxic genus responsible for causing Azaspiracid shellfish poisoning in humans (Tillmann et al., 2009) has never been reported in Australia. The analysis also revealed the presence of three potentially toxic, benthic, Gonyaucalean genera, Gambierdiscus, Ostreopsis and Coolia in samples collected from all the three sites. Seven unique OTUs were basal to the clade, consisting of the NOAA (National Oceanic and Atmospheric Administration) strain identified as G. carpenteri, which in turn was a sister clade to the NOAA strain identified as G. caribaeus (Figure 3.4). This indicated the presence of a single species of Gambierdiscus, G. carpenteri, at all the three sites on the coast of NSW. This is unusual as most previous studies have reported the presence of more than one species of Gambierdiscus co-occurring at one geographical region (Chinain et al., 1999b, Litaker et al., 2010). A recent study has demonstrated that different environmental factors such as temperature and salinity regimes may affect species distribution and abundance (Kibler et al., 2012). Also, at 98% sequence similarity, the presence of seven unique OTUs might indicate the presence of intraspecific diversity within G. carpenteri, a phenomenon known to occur in other Gonyaucalean taxa, such as Ostreopsis ovata (Sato et al., 2011). Two species of Coolia, C. monotis and C. canariensis, were present in the samples. While C. monotis is fairly common around the coast of NSW (Murray, 2010), C. canariensis has never been reported before from Australia. Prorocentrum lima, Gymnodinium dorsalisulcum and various Amphidinium species were among the other toxic species present in the samples analysed. The analysis also revealed the presence of a distinct “Dinotoms” clade (Imanian et al., 2012) within the Peridiniales and three out of the five know dinotom genera, Galeidinium, Kryptoperidinium and Durinskia were present in the samples analysed.

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3.5.2

Gambierdiscus species in NSW Identification of Gambieridicus species found in NSW was carried out via light and

scanning electron microscopy, as well as molecular genetics. Morphologically the Merimbula cells resembled what has been designated as the Tahiti epitype of G. toxicus GTT-91 (Figure 1 in Litaker et al. 2009) with a diagnostic hatchet-shaped 2’ plate. Plate overlap often prevented adequate resolution of the shape of the 2’ plate using SEM, while LM reflected more consistent plate morphology of 2’ being near rectangular but commonly with a small protrusion pointing towards the APC. Critically, none of these cells exhibited the thecal groove shown for G. carpenteri, or the dorsal rostrum (Figure 41 in Litaker et al 2009). Transdiameters of our Australian material exhibited a much broader range than described for the Belize-type G. carpenteri, with the larger sized cells (86-110 m) more commonly associated with G. toxicus (Tester et al. 2008, Litaker et al. 2009). Such large size variation for G. toxicus has been previously attributed to represent multiple species (Tester et al., 2008), but morphological variation of this taxon in nature remains poorly defined. Based on the phylogeny of two rRNA genes, 28S (Figure 3.3) and 18S (Figure 3.4), the species was conclusively identified as G. carpenteri with no evidence of other Gambierdiscus species present. G. carpenteri was first described by Litaker et al. in 2009 from Belize as a large anterio-posteriorly compressed species. Since then it has been reported from Guam, Fiji (Litaker et al., 2009), Hawaii (Litaker et al., 2010), Dry Tortugas (Florida), Flower Gardens (Gulf Of Mexico) and Ocho Rios (Jamaica) (Holland et al., 2013). Recently in Australia, G. carpenteri has been reported from Exmouth, Western Australia (WA) (Kohli et al, 2013) and QLD (Capper, pers. comm.). SEM micrographs of Australian Flinders Reef (27oS) Gambierdiscus cells collected in September 1983 (Gillespie et al., 1986; Figure 6D in Hallegraeff 1993) closely resemble the present Merimbula material, even though without molecular sequencing no conclusive identification is possible. The optimum salinity range at which this species can grow has been claimed to be 19.6-39.1 ppt with maximum growth observed at 27.3 ppt (Kibler et al., 2012). This indicates that G. carpenteri has a high tolerance towards salinity change and that it could survive in estuarine waters, where the salinity can change periodically. As shown in section 3.4.3, that cell counts at sites S2 and S3 were substantially lower than at S1 and S4. This may be due to the fact that sites S2 and S3 are open ocean sites and lie outside the Merimbula Inlet. These sites experience greater wave action when compared to sites S1 and S4, which lie within the Merimbula inlet where waters are shallower and currents are weaker. Therefore it forms a sheltered habitat for benthic dinoflagellates, such as Gambierdiscus, and in turn explains the high cell counts recorded at sites S1 and S4. The consistent seasonal presence of Gambierdiscus in Merimbula Lake from 2005 to 2013 (Ajani et al., 2013) suggested that the organism is now well-established in the region, rather than it being reintroduced every summer by the the East Australian Current as suggested by Hallegraeff, (2010) for other gonyaucalean 67

Chapter 3

taxa. Recent studies have also reported the presence of Gambierdiscus in other temperate regions such as G. reutzerli in North Carolina (Litaker et al., 2009), G. carolinianus in Crete (Holland et al., 2013) and Gambierdiscus sp. in the Mediterranean (Aligizaki and Nikolaidis, 2008). However, to our knowledge there is no evidence in the literature of Gambierdiscus forming cysts and there is no information regarding the different stages of the Gambierdiscus cell cycle. Therefore, the ability of sparse populations to overwinter in Merimbula Lake at temperatures as low as 8oC, as well as whether tropical and warm-temperate Australian populations of G. carpenteri are identical or represent different ecotypes, requires further investigation. Instances of a species of Gambierdiscus being the most numerically abundant and dominant species in a planktonic or benthic sample have been recorded in the literature since the early 1980s (Carlson et al., 1984, Litaker et al., 2010). However, very few studies have reported the identity of the causative species forming the bloom. It is often difficult to do so, as usually more than one species of Gambierdiscus are present at a given site, and identification and enumeration of different species of Gambierdiscus using light microscopy is challenging. This study is the first report of the single species G. carpenteri being the most numerically abundant microbial eukaryote at a benthic or planktonic site. Only 10% of reported blooms of a species of Gambierdiscus reach a cell density as high as 1000-10,000 cells g-1 wet weight algae (Litaker et al., 2010). In Merimbula, the highest density recorded in May 2013 was 8256 cells g-1 wet weight algae. While it is established that the cell density of Gambierdiscus populations is directly proportional to the risk of CFP in a particular area (Litaker et al., 2010), the risk also strongly depends on the species of Gambierdiscus present in that area. This is due to the fact that each species of Gambierdiscus has a unique toxin profile (section 1.5). Hence, characterisation of the toxin profile of each species is necessary in order to estimate the risk of CFP in a given area. The toxin profile of G. carpenteri has not yet been determined using LC-MS. Toxicity of G. carpenteri has only been studied using the in vitro human erythrocyte lysis assay, which showed cell extracts to be toxic (Holland et al., 2013). Due to the previous reports of Gambierdisucs populations in Flinders reef containing MTX but not CTXs (Gillespie et al. 1985), this toxicity was attributed to MTXs (Holland et al., 2013). MTX is a highly toxic compound, with a LD50 in mice via intraperitoneal injection of 0.05 μg/kg (Murata et al., 1993). Little is known about whether MTX plays a significant role in causing human illness. A recent feeding study carried out to probe the uptake of MTX has shown that it can accumulate in fish mussel tissue, digestive organs and liver (Chapter 4). In order to determine the risk of CFP in NSW due to G. carpenteri blooms, we tested the bloom sample for certain congeners of CTXs and MTXs via LC-MS. Neither CTXs nor MTXs were detected, however, only masses for the MTX and MTX-3 congeners were monitored. On the other hand, the extracts of the G. 68

Chapter 3

carpenteri bloom samples were found to be toxic in mouse bioassays. The symptoms of intoxication in mice after intraperitoneal injection (abdominal breathing, decreased respiration rates and death through respiratory paralysis) were similar to those in assays conducted for characterisation of MTX (Holmes et al., 1990, Holmes and Lewis, 1994) and of cell extracts obtained from cultures of Gambierdiscus belizeanus CCMP401, which produces MTXs (section 5.4.1). Therefore this toxicity was attributed to MTXs and not due to the classic polyunsaturated fatty acid effect observed with extracts from dinoflagellates. Neither CTXs nor MTXs were detected, however, only masses for the MTX and MTX-3 congeners were monitored. Hence, it is possible that G. carpenteri produces a MTX congener that was not detected by LC-MS. The results reported here clearly necessitate further investigations into the toxin profile of the G. carpenteri bloom sample.

69

Chapter 4

Chapter 4 A feeding study to probe the uptake of Maitotoxin by snapper (Pagrus auratus) 4.1

Summary The role of CTX in CFP has been investigated previously, but little is known about MTX

and whether it plays a significant role in causing human illness. The MTXs are known to have slightly higher potency than CTXs when administered intraperitoneally in mice, but are less potent when administered orally. It is not known whether MTXs accumulate in snapper (Pagrus auratus) muscle and to investigate this further, fish feeding trials with P. auratus were undertaken. Replicate P. auratus were fed with juvenile mullet (Aldrichetta forsteri) injected with a pellet of a known quantity of Gambierdiscus australes, which is a known producer of MTX. The levels of MTX in different fish tissues were determined using two newly developed sensitive LC-MS/MS assays for MTX that monitors either a specific cleavage fragment (generated from micro-scale oxidation of the intact toxin) or the intact toxin itself. The investigations revealed the presence of MTX in P. auratus viscera, liver and muscle samples. The presence of Gambierdiscus-related genes in P. auratus digestive organs was confirmed using DNA amplification technology.

4.2

Introduction Ciguatera Fish Poisoning (CFP) is a disease caused by ingestion of tropical fish that have

orally accumulated effective levels of toxins produced by a range of species of the genus Gambierdiscus (Holmes and Lewis, 1994, Chinain et al., 2010). Ciguatoxins (CTXs) and Maitotoxins (MTXs) are the two major toxin groups produced by Gambierdiscus (Holmes et al., 1990, Holmes et al., 1991, Holmes and Lewis, 1994, Holmes, 1998, Chinain et al., 1999a, Chinain et al., 2010, Rhodes et al., 2010, Fraga et al., 2011, Holland et al., 2013). The role of CTXs in CFP is well established. These toxins accumulate in the food web when toxic Gambierdiscus cells attached to surfaces, for example macroalgae, are consumed by herbivorous fish, which are then predated by carnivorous fish (Figure 4.1). The toxins undergo several oxidation steps during this passage through fish digestive systems, which increase their potency (Murata et al., 1990, Holmes et al., 1991, Lewis and Holmes, 1993, Yasumoto et al., 2000). Several CTX congeners have been isolated from the viscera-digestive organs, liver and muscle of various carnivorous fish (Murata et al., 1990, Lewis et al., 1991, Vernoux and Lewis, 1997, Lewis et al., 1998, Hamilton et al., 2002b, Pottier et al., 2002b, Pottier et al., 2003).

70

Chapter 4

Figure 4.1: Flow chart showing the mode of transmission of Ciguatoxins and Maitotoxins in Ciguatera fish poisoning. (1. (Legrand et al., 1989) 2. (Lewis and Sellin, 1992)) MTX was first described as a water-soluble cyclic polyether ladder toxin in 1976 (Yasumoto et al., 1976). Since this time a total of three MTX congeners have been described (Murata et al., 1993, Holmes and Lewis, 1994). The structural elucidation, stereochemistry and other chemical properties described for MTX and related analogues have been derived from materials extracted from cultures of Gambierdiscus. The principle MTX described in 1993 (Murata et al., 1993), also known as MTX-1 (Holmes and Lewis, 1994), represents the largest natural non-biopolymer toxin known (Yokoyama et al., 1988, Murata et al., 1993) with a protonated [M+H]+ mass of m/z 3422 (Murata et al., 1993, Holmes and Lewis, 1994) and LD50 dose of 0.05 μg/kg via intraperitonial injection in mice (Murata et al., 1993, Holmes and Lewis, 1994), making it one of the most toxic substances known to date (Murata et al., 1994). MTX-2 is also a large polyether ladder toxin with a protonated [M+H]+ mass of m/z 3298 and LD50 dose of 0.08 μg/kg via intraperitonial injection in mice (Holmes and Lewis, 1994). MTX-3 represents a small MTX analogue that has a reported protonated [M+H]+ mass of m/z 1060 (Holmes and Lewis, 1994). MTX-3 was found to be toxic to mice via intraperitonial injection(Holmes and Lewis, 1994), although the exact LD50 dose remains unknown. The only evidence of MTXs being accumulated in fish liver and viscera dates back to two studies published in the 70’s (Yasumoto et al., 1971, Yasumoto et al., 1976). Here, the toxic potency of a partially purified water soluble tissue extract (liver and viscera) was reported as 80 71

Chapter 4

mg/kg (Yasumoto et al., 1971) and 140 mg/kg (Yasumoto et al., 1976), and the toxicity of the purified MTX was reported as 15-20 mg/kg intraperitoneally (Yasumoto et al., 1976). Since this time 3 MTX analogues have been described and found to be far more potent via the intraperitonial route than earlier estimated. The major aims of this study were: 1. Investigate whether MTX-1, a large water-soluble marine toxin, can accumulate in various tissues of carnivorous fish fed with Gambierdiscus australes. 2. Detect the presence of Gambierdiscus australes cells in carnivorous fish viscera using genetic markers. In order to achieve this, two fish trials were conducted. In each experiment, carnivorous snapper (Pagrus auratus) were fed with juvenile herbivorous mullet (Aldrichetta forsteri) that had been injected with known quantities of G. australes. The levels of MTX-1 in the liver, viscera and muscle of Pagrus auratus specimens were determined using two recently developed sensitive LC-MS/MS assays. A PCR-based approach using a dinoflagellate specific genetic marker was employed to detect the presence of G. australes cells in P. auratus viscera.

4.3 4.3.1

Materials and methods Culturing and growth conditions A strain of Gambierdiscus isolated from Rarotonga, Cook Islands was used in the present

study (Rhodes et al., 2010). This strain was identified as Gambierdiscus australes (CAWD 149) and is maintained at the Cawthron Institute Culture Collection of Micro-algae (CICCM). Cultures were grown at 25°C, in 14:10 hour light:dark cycle with 60 μmol m−2 s−1 light intensity and f/2 medium (Guillard and Ryther, 1962) in 2 litre conical flasks for 20-24 days. For harvesting, cultures were shaken gently taking care not to disrupt the mucous layer at the bottom, poured into beakers and left for 3-4 hours for cells to settle down. Once the cells settled at the bottom of the beaker, 2/3rds of the media was skimmed from the top. Sub samples (100 μL) were taken for counting using an Utermohl chamber. Known numbers of cells from the thick culture concentrate was used for feeding purposes. Centrifugation of cultures was not used to harvest cells in order to minimise cell disruption.

4.3.2

DNA extraction, PCR amplification and gene sequencing A 50 mg pellet of homogenised gut content of P. auratus was used to extract total

genomic DNA via FastDNA® Spin kit for Soil (MP Biomedicals, Solon, OH). The manufacturer’s protocol was followed, and samples were stored at -20°C until PCR amplification.

72

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All PCR reactions were performed in 50 μL reaction volumes containing 10 μL Taq polymerase buffer, 5 μL GC Melt (Clontech, Mountain View, CA), 0.2 mM deoxynucleotide triphosphates, 10 pmol each of the forward and reverse primers, between 10 and 100 ng genomic DNA, and 0.5 U of Advantage®-GC 2 Taq Polymerase (Clontech, Mountain View, CA). A nested PCR approach for applied to amplify the large ribosomal subunit of rRNA. The first round of PCR was performed using primers D1F (ACCCGCTGAATTTAAGCATA) (Scholin et al., 1994) and DS-D1R (ACACCTCGGAAGACAAGT) and the following cycle: initial denaturation at 96°C for 3 min followed by 22 cycles of denaturation at 94°C for 20 s, primer annealing at 55°C for 20 s and an extension at 68°C for 2 min followed by a final extension step at 68°C for 3 min. In the second round of PCR, the 2 μL of the PCR product from the first round was used as a template. The PCR was performed using primers D1F and D3B and the following cycle: initial denaturation at 96°C for 3 min followed by 25 cycles of denaturation at 94°C for 20 s, primer annealing at 55°C for 20 s and an extension at 68°C for 1 min followed by a final extension step at 68°C for 3 min. The PCR products were visualised via agarose gel electrophoresis. The correct size bands were excised from the gel and purified using the ZymocleanTM gel DNA recovery kit (Zymo Research, Irvine, CA) and sequenced. Sequencing was performed using the PRISM BigDye cycle sequencing system and a model 373 sequencer (Applied Biosystems, Foster City, CA) at the Ramaciotti Center for Gene Function Analysis (University of NSW). Sequence data were trimmed and analysed using Geneious® software (Kearse et al., 2012) while identity/similarity values to other sequences were determined using BLAST in conjugation with the National Center for Biotechnology Information (NIH, Bethesda, MD).

4.3.3

Fish feeding trials Two fish trials were conducted using snapper (Pagrus auratus) obtained from Plant &

Food Research Limited (New Zealand). The average snapper weight used in the two trials was 60 g and 246.6 g respectively. The large average mass difference was due to the availability of the fish at that time. In the first trial there were four experimental fish holding tanks (capacity 50 L) with four fish in each tank. The control tank contained three snapper. After arrival from the breeding facility, the snapper were acclimatised for three days without feeding. On day four they were fed with frozen juvenile mullet (Aldrichetta forsteri) injected with known quantities of Gambierdiscus australes. To prepare the mullet for feeding, 400-600 μL of culture concentrate (section 4.3.1) was carefully injected between the mullet skin and muscle to prevent the culture from spilling out in the tank during feeding (Figure 4.2).

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A

B

Figure 4.2: Pictorial representations of mullet (A. forsteri) being injected with concentrated G.australes culture; A- before injecting the culture, B- after injecting the culture in between the mullet skin and muscle. The mullet were prepared fresh each time before feeding the snapper and one mullet per snapper were given in each feeding session. During the course of seventeen days the snapper were fed fifteen times. The control tank was fed with mullet without any Gambierdiscus. During the trials and before feeding, the fish excrement was siphoned out of the tanks using a serological pipette and kept for further analysis. Simultaneously, after each feeding session the tanks were cleaned and half the water changed to maintain hygienic conditions. At the end of the trials the snapper were anaesthetised by giving them an overdose of Aqui-S10 (Aqui-S New Zealand Ltd, Lower Hutt, New Zealand) which contains 5% Isoeugenol. An appropriate amount of Aqui-S10 was poured in each tank mixed to reach a final concentration of 500 ppm. After 30 min the snapper were removed from the tanks and sacrificed by an incision in the brain. The viscera, liver and muscle tissues were dissected and kept at -20°C for further analysis. Muscle tissue refers to the whole fish carcass including the bones and scales, which were homogenised and stored at -20°C for further analysis. In the second fish trial there were seven experimental fish holding tanks with sixteen snapper (five tanks with two snapper in each tank and two large tanks with three snapper each). After arrival from the breeding facility, the snapper were acclimatised for four days. On day five, the snapper were fed with frozen juvenile mullet (Aldrichetta forsteri) injected with 300 μL of Gambierdiscus australes culture concentrate. During the course of eight days (day five to day twelve) the snapper were fed twelve times. Three snapper died at different time points during the eight days feeding period. The viscera, liver and muscle samples of these dead snapper were dissected and preserved at -20°C for LC-MS analysis. Ten of the thirteen snapper were sacrificed after eight days of feeding (at day twelve) and three were transferred into separate tanks and kept alive for studying the toxin depuration time. Toxin depuration was carried out for ten days (day thirteen to day twenty-two). One snapper was sacrificed at day sixteen and the other two were sacrificed at day twenty-two. Throughout the depuration time, the snapper were fed twice daily with frozen mullet not infused with Gambierdiscus cells. 74

Chapter 4

Viscera, liver and muscle samples of the all the sacrificed fishes were dissected and stored at 20°C for LC-MS analysis. Unlike the first fish feeding trial, this time only the fish muscle tissue was dissected from the rest of the carcass and used for LC-MS analysis.

4.3.4

Sample extraction and LC-MS analysis Novel, sensitive LC-MS/MS methods were developed for quantitative analysis of MTX-1

from algal cultures and fish matrices. These methods rely on quantitation of either a specific oxidative cleavage product (method 1) or the intact molecule (method 2). For the intact MTX method, it was also possible to simultaneously monitor a putative MTX analogue previously described as MTX-3 (Holmes and Lewis, 1994) and produced by various Gamberdiscus species, including G. australes CAWD149 used here. The principle of the oxidative cleavage method is similar to that recently described for palytoxin monitoring a specific vicinal diol cleavage fragment (Figure 4.3) generated by periodate oxidation of the intact molecule (Selwood et al., 2012). Method 1 was used to quantify samples from the first fish trials while method 2 was used for quantification of samples obtained during the second trial. For comparison, 3 muscle samples from the first trial were also quantified using method 2. For sample preparation, P. auratus muscle, viscera or liver samples (2.0 g) were homogenised with 18 mL methanol–water (1:1 v/v) using an ultra-Turrax (IKA, Guangzhou, China) at 19,000 rpm for 1 min followed by centrifugation at 3000 g for 10 min. Samples were left on ice for at least 2 hours to aid protein precipitation and then re-centrifuged to pellet insoluble debris. The supernatant was decanted and subject to either an on-column oxidation procedure (method 1) or SPE clean up (method 2). In method 1, to monitor the oxidative cleavage product, MTX standard (20-80 ng in 6 mL methanol–water (1:1 v/v) or tissue extract (up to 12 mL) was loaded onto a 60 mg Strata-X SPE cartridge (Phenomenex, CA), pre-conditioned with 3 mL methanol followed by 3 mL of methanol-water (1:1 v/v) containing 0.1% formic acid. The column was washed with 2 mL methanol–water (1:1 v/v) containing 0.1% formic acid followed by 2 mL water. The sample was then oxidised on the column by passing through 2 mL of 50 mM periodic acid at approximately 1 mL min-1. The column was then eluted, without washing, using 1.8 mL of acetonitrile–water (1:4 v/v) containing 20mM ammonium acetate. The eluent was transferred into a glass vial and used for further LC-MS/MS analysis. Figure 4.3 shows the oxidative cleavage reaction of MTX-1 and its product generated during the on-column oxidation procedure. In method 2, to monitor the intact toxin, MTX standard (15-75 ng in 6 mL methanol–water 1:1 v/v) or tissue extract (6 mL) was loaded onto a 60 mg Strata-X SPE cartridge (Phenomenex, CA), pre-conditioned with 3 mL methanol followed by 3 mL Milli-Q water. The column was washed with 3 mL methanol–water (3:2 v/v) containing 0.1% formic acid, and eluted with 3 mL acetonitrile-water (1:1 v/v) containing 0.2% ammonium hydroxide. 75

Chapter 4

The eluent was mixed and transferred to a glass vial for analysis. Both the methods used an Acquity ultra-performance liquid chromatography system (Waters, Milford, MA), coupled to a Xevo TQ-S triple quadrupole mass spectrometer (Waters, Manchester, UK).

Figure 4.3: Structure of Maitotoxin-1 and its oxidative cleavage product obtained during oncolumn periodate oxidation and quantified via LC-MS/MS analysis, method 1. Separation of oxidation products using method 1 was achieved on a Thermo Hypersil Gold column 50 x 1 mm (Waters, Milford, MA, USA). For quantitative analysis, the following multiple reaction monitoring (MRM) transition was acquired with 50 ms dwell time: m/z 971.4/96.8 (CE 80 eV). The fragment ion monitored represents the bisulphite anion of the MTX oxidative cleavage product (m/z 971.4). In method 2, chromatographic separation of intact MTX was achieved using an Acquity UPLC BEH Phenyl 1.7 µm column 100 x 2.1 mm (Waters, Milford, MA, USA). For quantitative analysis, a pseudo multiple-reaction-monitoring (MRM) transition was acquired with a 100 ms dwell times: m/z 1689.6/1689.6 (CE 50 eV) for intact MTX. The parent ion monitored (m/z 1689.6) represents the doubly charged anion [M-2H]2- of intact MTX; m/z 1037.6/96.8 (CE 70 eV) for the putative MTX-3 analogue. The parent ion monitored (m/z 1037.6) in this case represents [M-H]-. For both methods, peak areas were generated and sample concentrations calculated from linear calibration curves generated from MTX standards. TargetLynx software (Waters, Milford, MA) was used for the analysis. For method 1, the extraction efficiency (SPE recovery) for each type of tissue was calculated as follows: muscle -118%, viscera 114%, liver 83%. The limit of detection (LOD) for this method was determined to be 10 μg/kg for muscle tissue and 20 μg/kg for viscera and liver tissue samples. In regards to the yield during oxidative cleavage reaction, a molar excess of 76

Chapter 4

oxidant was used and this would be expected to result in 100% of reaction product being generated. This is demonstrated by the good recoveries observed with the MTX-1 oxidative cleavage method, and with palytoxin oxidative cleavage fragments (Selwood et al., 2012). For method 2, the extraction efficiency of intact MTX-1 from the SPE cartridge was reasonably poor with less than 50% of the MTX recovered. This occurred even in the absence of matrix. To account for these losses the MTX standards were subjected to the SPE procedure at the same time as the fish samples (muscle, viscera or liver) were prepared. When this was done, recoveries for all three matrices were greater than 95%. The LOD for this method was determined to be 1 μg/kg. This analysis was performed on a Waters Xevo TQ-S LC-MS (Waters, Milford, MA), which is more sensitive than the instrument that was used for the oxidative cleavage analysis that is, the Waters Premier XE LC-MS (Waters, Milford, MA).

4.4 4.4.1

Results and discussion Dinoflagellate culture and toxin production The toxin profile of G. australes used in this study is well defined. It does not produce

any known CTX congeners (Rhodes et al., 2010) but does produce MTX. The amount of MTX1 produced by each cell was estimated to be 8.6 pg/cell via the method 1 LC-MS/MS analysis. As the main focus of this study was to test the accumulation potential of MTXs in fish, this strain was used in both the fish feeding trials due to its unique toxin profile.

4.4.2

Fish feeding trials and LC-MS analysis In the first fish trial, after feeding with mullet injected with Gambierdiscus, the levels of

MTX-1 were quantified in the P.auratus viscera, liver and muscle samples. All three tissues were positive for MTX-1 accumulation and the amount of MTX-1 in each sample was quantified by LC-MS analysis using method 1 (Table 4.1). Figure 4.5 shows chromatograms obtained when using the oxidative cleavage method for samples that were positive for MTX-1. The level of MTX-1 observed in P. auratus viscera and liver tissue was much higher than observed in the muscle (Figure 4.4). According to the mass balance calculations, out of the total amount of MTX-1 offered to snapper during the first fish trial, (samples taken at the end of the trial) about 4.9 % was present in the viscera, 1.6 % was present in liver and 32 % accumulated in the muscle samples. As accumulation and depuration are continuous processes, these values represent a snapshot of one time point (day seventeen of fish trial). To reconfirm the observed results, the amount of MTX-1 in fish muscle was also quantified using the intact toxin LC-MS/MS analysis (method 2). This method of toxin analysis directly measures the intact toxin, which ensures only the analyte of interest is being monitored. Both detection methods showed MTX-1 was present in the muscle samples from the first fish feeding trial. However, the amount of MTX-1 measured using the intact MTX method was 77

Chapter 4

almost 3 times lower than that obtained when using the oxidative cleavage method. This difference might be due to the quantification of other structurally related chemical compounds (ie metabolites of MTX) that give the same oxidation product when using method 1. Degradation of sample during storage might also be another reason for different values as quantification of MTX-1 via method 2 was carried out 8 months after the method 1. Regardless, these results show for the first time that despite being a water-soluble toxin, MTX-1 can accumulate in fish muscle.

Figure 4.4: Bar graph representing the amount of Maitotoxin-1 detected in different tissue types during the first fish feeding trial. Table 4.1: Data obtained during the first fish feeding trial. Tank Number Number of samples of P. auratus

1 4 (pooled) 13-18 60

2 4 (pooled) 13-18 60

3 4 (pooled) 13-18 60

4 4 (pooled) 13-18 60

Control 3 (pooled) -60

Days of Feeding- No. of Feeds Average P. auratus Weight (grams) 116 116 199 199 -Total MTX-1 offered (μg)1 MTX-1 in Viscera (μg/kg) via 109 298 582 631 96.8 (Monitoring bisulphate anion) 5.70e4

0.40

0.50

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0.70

0.80

0.90

1.00

1.10

1.20

1.30

1.40

1.50

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1.70

1.80 1.90 1: MRM of 3 Channels ES971.4 > 96.8 (Monitoring bisulphate anion) 8.83e5

0.40

0.50

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0.90

1.00

1.10

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%

B

1.70

1 0.00 0.10 PLTX120615_32

0.20

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%

C

0 0.00 0.10 PLTX120615_21

0.20

0.30

%

D

0 0.00

Time 0.10

0.20

0.30

1.80

1.90

Figure 4.5: Extracted ion chromatograms for Maitotoxin-1 from various extracts following SPE cleanup and on-column oxidation (Method 1) during the first fish feeding trial. A. Blank snapper muscle sample; B. Maitotoxin-1 positive muscle sample from first fish feeding trials; C. Maitotoxin-1 positive cell extract from G. australes CAWD149; D. Maitotoxin-1 calibration standard (25 ng/mL). The second set of feeding trials were conducted to confirm the findings from the first trial, quantify levels of MTX-1 in the P. auratus viscera, liver and muscle samples and also to estimate the depuration time of MTX-1. The levels of MTX-1 in all the samples were quantified via the second LC-MS method, which is specific for the intact MTX toxin. After eight days of feeding P. auratus with mullet injected with G. australes cells, MTX-1 was detected in all P. 79

Chapter 4

auratus viscera samples and one P. auratus liver sample (Table 4.2). However, MTX-1 levels were below the detection limit in P. auratus muscle samples. The P. auratus used in this trial were four times larger than the first experiment (247g versus 60g), and were fed for fewer days than the first trial (Table 4.2). This might explain the absence of MTX-1 in the muscle samples, as the toxin amount offered (per gram of fish muscle) in the first feeding trial was much greater than the second fish trial. Therefore it is possible that MTX-1 might be present in the muscle samples in much lower amounts and below the detection limit of the assay. Figure 4.6 shows chromatograms obtained via the second method of LC-MS analysis where viscera samples were positive for MTX-1. MTX130515_31

MRM of 2 Channels ES1689.6 > 1689.6 5.37e3

%

A

15 2.20 MTX130515_26

2.40

2.60

2.80

3.00

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4.80 MRM of 2 Channels ES1689.6 > 1689.6 3.46e4

2.80

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4.80 MRM of 2 Channels ES1689.6 > 1689.6 3.25e4

2.80

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4.00

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4.80 MRM of 2 Channels ES1689.6 > 1689.6 6.98e4

3.00

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4.00

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%

B

2 2.20 MTX130515_39

2.40

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%

C

2 2.20 MTX130515_24

2.40

2.60

%

D

1

Time 2.20

2.40

2.60

2.80

4.80

Figure 4.6: Total ion chromatograms for intact Maitotoxin-1 molecule from various extracts following SPE cleanup (Method 2) during the second fish feeding trial. A. Blank snapper viscera sample; B. Maitotoxin-1 positive viscera sample from second fish feeding trial; C. Maitotoxin-1 positive cell extract from G. australes CAWD149; D. Maitotoxin-1 calibrant (25 ng/mL). 80

Chapter 4

To estimate the depuration time of MTX-1, 3 P. auratus were kept alive after eight days of initial feeding (day five to day twelve) and then sacrificed at day sixteen and day twenty-two. MTX-1 was observed in lower amounts in the P. auratus viscera sample from day sixteen (after two days of depuration) and no MTX-1 was observed in the P. auratus viscera from day twenty-two onwards (ten days of depuration). During the intact toxin LC-MS analysis (method 2), apart from the MTX-1, it was also possible to monitor another putative MTX analogue that has been described as MTX-3 (Holmes and Lewis, 1994). The putative MTX was also observed in the muscle samples obtained during the first fish trial, when samples were re-analysed using the intact toxin LC-MS/MS analysis method. Figure 4.7 shows chromatograms obtained via the second method of LC-MS analysis where the samples showed the presence of the putative MTX-3, during the second fish feeding trials.

Table 4.2: Data obtained during the second fish feeding trial.

Fish Tank

A B C D

E F G

Fish Number 1 2 1 2 1 2 1 2 3 1 2 3 1 1 2

Days of FeedsNo. of Feeds

Depuration Time (days)

Average Snapper Weight (grams)

5-9 7-11 No Depuration 246.6 8-12

2 10

MTX-1 in Viscera (μg/kg) via method 2 8.9 2.3 1.4 5.1