Biosensors for the Marine Environment: Present and Future Challenges 303132000X, 9783031320002

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The Handbook of Environmental Chemistry 122 Series Editors: Damià Barceló · Andrey G. Kostianoy

Fiona Regan Peter-Diedrich Hansen Damià Barceló   Editors

Biosensors for the Marine Environment Present and Future Challenges

The Handbook of Environmental Chemistry Volume 122 Founding Editor: Otto Hutzinger Series Editors: Damia Barcelo´ • Andrey G. Kostianoy

Editorial Board Members: Jacob de Boer, Philippe Garrigues, Ji-Dong Gu, Kevin C. Jones, Abdelazim M. Negm, Alice Newton, Duc Long Nghiem, Sergi Garcia-Segura, Paola Verlicchi, Stephan Wagner, Teresa Rocha-Santos, Yolanda Pico´

In over four decades, The Handbook of Environmental Chemistry has established itself as the premier reference source, providing sound and solid knowledge about environmental topics from a chemical perspective. Written by leading experts with practical experience in the field, the series continues to be essential reading for environmental scientists as well as for environmental managers and decisionmakers in industry, government, agencies and public-interest groups. Two distinguished Series Editors, internationally renowned volume editors as well as a prestigious Editorial Board safeguard publication of volumes according to high scientific standards. Presenting a wide spectrum of viewpoints and approaches in topical volumes, the scope of the series covers topics such as • • • • • • • •

local and global changes of natural environment and climate anthropogenic impact on the environment water, air and soil pollution remediation and waste characterization environmental contaminants biogeochemistry and geoecology chemical reactions and processes chemical and biological transformations as well as physical transport of chemicals in the environment • environmental modeling A particular focus of the series lies on methodological advances in environmental analytical chemistry. The Handbook of Environmental Chemistry is available both in print and online via https://link.springer.com/bookseries/698. Articles are published online as soon as they have been reviewed and approved for publication. Meeting the needs of the scientific community, publication of volumes in subseries has been discontinued to achieve a broader scope for the series as a whole.

Biosensors for the Marine Environment Present and Future Challenges Volume Editors: Fiona Regan
Peter-Diedrich Hansen
Damia Barceló

With contributions by F. S. A. Bracken
C. Briciu-Burghina
M. Campas
M. Castro-Freitas
E. Costa-Rama
A. Dahlhaus
H. Dalhlhaus
C. Delerue-Matos
J. Dioge`ne
G. Gaiani
P.-D. Hansen
J. Hauslage
R. Hemmersbach
O. Idelegbagbon
S. Kurbanoglu
C. Moldaenke
C. Murphy
H. P. A. Nouws
Y. Pan
A. Parle-McDermott
F. Regan
J. P. Rocha
F. W. Scheller
B. Schierwater
R. Torre
E. Unruh
M. A. Williams
Z. Yang
A. Yarman
A. Zaake
X. Zhang

Editors Fiona Regan School of Chemical Sciences Dublin City University Dublin, Ireland

Peter-Diedrich Hansen International Consulting Ecotoxicology Technische Universita¨t Berlin, REM Berlin, Germany

Damia Barcelo´ Catalan Institute for Water Research (ICRA-CERCA) Girona, Spain

ISSN 1867-979X ISSN 1616-864X (electronic) The Handbook of Environmental Chemistry ISBN 978-3-031-32000-2 ISBN 978-3-031-32001-9 (eBook) https://doi.org/10.1007/978-3-031-32001-9 © The Editor(s) (if applicable) and The Author(s), under exclusive license to Springer Nature Switzerland AG 2023 This work is subject to copyright. All rights are solely and exclusively licensed by the Publisher, whether the whole or part of the material is concerned, specifically the rights of translation, reprinting, reuse of illustrations, recitation, broadcasting, reproduction on microfilms or in any other physical way, and transmission or information storage and retrieval, electronic adaptation, computer software, or by similar or dissimilar methodology now known or hereafter developed. The use of general descriptive names, registered names, trademarks, service marks, etc. in this publication does not imply, even in the absence of a specific statement, that such names are exempt from the relevant protective laws and regulations and therefore free for general use. The publisher, the authors, and the editors are safe to assume that the advice and information in this book are believed to be true and accurate at the date of publication. Neither the publisher nor the authors or the editors give a warranty, expressed or implied, with respect to the material contained herein or for any errors or omissions that may have been made. The publisher remains neutral with regard to jurisdictional claims in published maps and institutional affiliations. This Springer imprint is published by the registered company Springer Nature Switzerland AG The registered company address is: Gewerbestrasse 11, 6330 Cham, Switzerland

Series Editors Prof. Dr. Damia Barcelo´

Prof. Dr. Andrey G. Kostianoy

Department of Environmental Chemistry IDAEA-CSIC Barcelona, Spain and Catalan Institute for Water Research (ICRA) Scientific and Technological Park of the University of Girona Girona, Spain [email protected]

Shirshov Institute of Oceanology Russian Academy of Sciences Moscow, Russia and S.Yu. Witte Moscow University Moscow, Russia [email protected]

Editorial Board Members Prof. Dr. Jacob de Boer VU University Amsterdam, Amsterdam, The Netherlands

Prof. Dr. Philippe Garrigues Universite´ de Bordeaux, Talence Cedex, France

Prof. Dr. Ji-Dong Gu Guangdong Technion-Israel Institute of Technology, Shantou, Guangdong, China

Prof. Dr. Kevin C. Jones Lancaster University, Lancaster, UK

Prof. Dr. Abdelazim M. Negm Zagazig University, Zagazig, Egypt

Prof. Dr. Alice Newton University of Algarve, Faro, Portugal

Prof. Dr. Duc Long Nghiem University of Technology Sydney, Broadway, NSW, Australia

Prof. Dr. Sergi Garcia-Segura Arizona State University, Tempe, AZ, USA

Prof. Dr. Paola Verlicchi University of Ferrara, Ferrara, Italy

Prof. Dr. Stephan Wagner Fresenius University of Applied Sciences, Idstein, Germany

Prof. Dr. Teresa Rocha-Santos University of Aveiro, Aveiro, Portugal

Prof. Dr. Yolanda Picó Desertification Research Centre - CIDE, Moncada, Spain

Series Preface

With remarkable vision, Prof. Otto Hutzinger initiated The Handbook of Environmental Chemistry in 1980 and became the founding Editor-in-Chief. At that time, environmental chemistry was an emerging field, aiming at a complete description of the Earth’s environment, encompassing the physical, chemical, biological, and geological transformations of chemical substances occurring on a local as well as a global scale. Environmental chemistry was intended to provide an account of the impact of man’s activities on the natural environment by describing observed changes. While a considerable amount of knowledge has been accumulated over the last four decades, as reflected in the more than 150 volumes of The Handbook of Environmental Chemistry, there are still many scientific and policy challenges ahead due to the complexity and interdisciplinary nature of the field. The series will therefore continue to provide compilations of current knowledge. Contributions are written by leading experts with practical experience in their fields. The Handbook of Environmental Chemistry grows with the increases in our scientific understanding, and provides a valuable source not only for scientists but also for environmental managers and decision-makers. Today, the series covers a broad range of environmental topics from a chemical perspective, including methodological advances in environmental analytical chemistry. In recent years, there has been a growing tendency to include subject matter of societal relevance in the broad view of environmental chemistry. Topics include life cycle analysis, environmental management, sustainable development, and socio-economic, legal and even political problems, among others. While these topics are of great importance for the development and acceptance of The Handbook of Environmental Chemistry, the publisher and Editors-in-Chief have decided to keep the handbook essentially a source of information on “hard sciences” with a particular emphasis on chemistry, but also covering biology, geology, hydrology and engineering as applied to environmental sciences. The volumes of the series are written at an advanced level, addressing the needs of both researchers and graduate students, as well as of people outside the field of vii

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Series Preface

“pure” chemistry, including those in industry, business, government, research establishments, and public interest groups. It would be very satisfying to see these volumes used as a basis for graduate courses in environmental chemistry. With its high standards of scientific quality and clarity, The Handbook of Environmental Chemistry provides a solid basis from which scientists can share their knowledge on the different aspects of environmental problems, presenting a wide spectrum of viewpoints and approaches. The Handbook of Environmental Chemistry is available both in print and online via https://link.springer.com/bookseries/698. Articles are published online as soon as they have been approved for publication. Authors, Volume Editors and Editors-in-Chief are rewarded by the broad acceptance of The Handbook of Environmental Chemistry by the scientific community, from whom suggestions for new topics to the Editors-in-Chief are always very welcome. Damia Barcelo´ Andrey G. Kostianoy Series Editors

Preface

The marine environment represents our planet’s largest ecosystem. It is critical for the stability of our climate, carbon storage, maintaining biodiversity, and supporting human life through the provision of resources and food. We depend on the seas and oceans for our well-being. However, this important resource is not valued or adequately studied. In order to support sustainable, ecosystem-based policies and measures for marine ecosystems, significantly more research and observations are needed. Under the threat of climate change, the world’s oceans, coasts, and marine ecosystems are undergoing great changes caused by increasing greenhouse gases, coastal pollution, overfishing, coastal development, and increasing population pressures. Ocean science has progressed dramatically over the last century by exploring, monitoring, and understanding the impact of changes in the ocean system. The UN Decade of Ocean Science for Sustainable Development calls on scientists to study the ocean’s responses to pressures. Ocean observations and research are essential to predict the consequences of change, to help design mitigation, and to guide adaptation measures. Sensors are traditionally used in marine studies to determine physical parameters, but there is increasing demand for real-time information about chemical and effects-related biological parameters. These parameters are currently measured in samples collected at sea and subsequently analysed in the laboratory. With the growing scientific capacity in merging biological recognition with platform engineering, we are now able to achieve data gathering on all aspects of marine health. By integrating a hierarchy of measurement systems from satellite to in situ, we have the potential to observe the ocean in real time to make important decisions and protect this vital resource. That being said, this book is timely published and extremely helpful as well. This book draws together leading research in the area of marine biosensing by reporting key developments and applications. This book contains ten chapters written by experts in this field. In short, some topics described here are at the fundamental stage with the potential for ground-breaking advances for monitoring, while others ix

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are already being tested in the marine environment. The range of applications and target parameters reported in this book is very comprehensive with examples from single species detection, marine toxins, and global environmental parameters as well. In this sense, biosensors are implemented in the so-called in situ effects monitoring programmes in line with the recommendations of ICES (International Council for the Exploration of the Sea) for monitoring effects of offshore and onshore aquaculture effluents, toxic algae blooms, offshore explorations, coastal management, etc. Another practical example on the use of biosensors in the marine environment is the monitoring of the UV-Treatment of Ballast Water by a wholecell sensor as a controlling unit of a “Ballast Water Management System (BWMS)”. Whole-cell biosensors play an important role in the documentation of the changes in the immune system of organisms caused by a changing marine environment. Other types of recently developed sensors well covered in different chapters of this book are electrochemical molecular imprinted polymers (MIPS) as well as paper-based biosensors, recently popular as well for SARS-CoV-2 control in wastewater. We believe that the developments and applications reported here are relevant to the objectives of the UN Ocean Decade and in the context of a changing climate and consequently changes in marine ecosystem services. A take-home message is the recommendation to continue to invest in biosensor research and development. This book is a good example of the outcomes of such investments. This book will help to better understand the changes in the marine environment as well as to monitor in short- and long-term ranges, the impacts on site and in reference areas simultaneously using validated Decision Support Systems (DSS). Lastly, we would like to thank all recognized authors in this field for preparing their timely, practical, and easily readable chapters. This volume is highly recommended for marine scientists, analytical chemists, ecotoxicologists, and (bio)sensor experts as well as for newcomers and graduate students who want to learn more about this challenging topic of sensors for the marine environment. We are sure that all of them will enjoy this insightful and enjoyable read! Dublin, Ireland Berlin, Germany Girona, Spain

Fiona Regan Peter-Diedrich Hansen Damia Barcelo

Contents

Biosensors for the Marine Environment: Introduction . . . . . . . . . . . . . . F. Regan and Peter-Diedrich Hansen Microgravity Changes Membrane Properties and Triggers Bioluminescence in Pyrocystis noctiluca as an Approach for New Biosensor Concepts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Jens Hauslage, Ruth Hemmersbach, and Bernd Schierwater

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Addressing Ciguatera Risk Using Biosensors for the Detection of Gambierdiscus and Ciguatoxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Greta Gaiani, Jorge Dioge`ne, and Mo`nica Campas

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Antibody, Aptamer and Affimer-Based Affinity Tools for Marine Toxin Biosensing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Caroline Murphy

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Environmental DNA as a Tool for Single Species Detection . . . . . . . . . . Molly Ann Williams, Fiona S. A. Bracken, Osatohanmwen Idelegbagbon, and Anne Parle-McDermott

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Paper-Based Devices for Virus Detection in Water . . . . . . . . . . . . . . . . Yuwei Pan and Zhugen Yang

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Electrochemical MIP Sensors for Environmental Analysis . . . . . . . . . . 139 Sevinc Kurbanoglu, Aysu Yarman, Xiaorong Zhang, and Frieder W. Scheller Whole-Cell Biosensors and Phagocytosis with Cryo-Conserved Cells in Coastal Areas and in Orbit (ISS) Under Microgravity . . . . . . . . . . . . 165 Peter-Diedrich Hansen and Eckehardt Unruh Marine Whole-Cell Biosensing for “Real-Time” Determination of the Ballast Water Treatment Efficiency . . . . . . . . . . . . . . . . . . . . . . . 183 C. Moldaenke, A. Zaake, A. Dahlhaus, and H. Dalhlhaus xi

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Sensors for Monitoring Faecal Indicator Bacteria in Bathing Waters . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 201 Ciprian Briciu-Burghina and Fiona Regan Electrochemical (Bio)sensors for Toxins Control in the Marine Environment . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 229 Jose´ Pedro Rocha, Ricarda Torre, Maria Castro-Freitas, Estefanı´a Costa-Rama, Henri P. A. Nouws, and Cristina Delerue-Matos

Biosensors for the Marine Environment: Introduction F. Regan and Peter-Diedrich Hansen

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Challenges . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Components of a Biosensor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 The Future of Marine Sensing . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

2 3 4 7 7

Abstract The marine environment is impacted by the increasing population and conflicts by industrial activities in the coastal areas. Activities such as dumping of waste, port construction, dredging and extraction processes all contribute to the water quality in the coastal zone. These and the climate related impacts are increasing the pressures on the marine environment and the ecosystem that it supports. The UN Decade of Ocean Science for Sustainable Development calls on scientists to study the ocean’s responses to pressures. Ocean observations and research are essential to predict the consequences of change, design mitigation and guide adaptation. Sensors are traditionally used in marine studies to determine physical parameters, but there is increasing demand for real-time information about chemical and biological parameters. These parameters are currently measured in samples collected at sea and subsequently analysed in the laboratory. With the growing scientific capacity in merging biological recognition with platform engineering we are now able to achieve data gathering on all aspects of marine health. By integrating a hierarchy of measurement systems from satellite to in situ, we have the potential to F. Regan (✉) School of Chemical Sciences, Dublin City University, Dublin, Ireland e-mail: [email protected] P.-D. Hansen (✉) International Consulting Ecotoxicology (ICEco), Technische Universität Berlin, Faculty VI, Institute for Ecology, FG Ecotoxicology, Berlin, Germany e-mail: [email protected] Fiona Regan, Peter-Diedrich Hansen, and Damià Barceló (eds.), Biosensors for the Marine Environment: Present and Future Challenges, Hdb Env Chem (2023) 122: 1–10, DOI 10.1007/698_2022_952, © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023, Published online: 29 January 2023

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observe the ocean in real time to make important decisions and protect this vital resource. Graphical Abstract

Keywords Ocean observing, Marine toxins, Real-time monitoring, Innovative biosensor technologies

1 Introduction The oceans cover over 70% of our planet. Over 30% of marine habitats are overused or have been destroyed and ocean acidity is up 26% with oxygen depletion occurring in key areas. In order to assess the impacts of activities and implement remediation, it is necessary to monitor a range of marine parameters [1, 2]. This book “Biosensors for the Marine Environment: Present and Future Challenges” is a follow-up of the volume on Biosensors for the Environmental Monitoring of Aquatic Systems published in this Handbook Series in 2009. Several biosensors and biochemical responses were developed and coordinated within EU projects to guide technological developments in Biosensors for Environmental Technology (BIOSET). Within the EU projects PRENDISENSOR and SANDRINE project the biosensor VITELLO was developed to measure endocrine effects related indicators like the so-called Vitellogenin (VTG) level in male fish. The EU project ALGAETOX contributed to the phagocytosis effects related biosensors by algae toxins and units with biosensors with unique applications in harsh environments in orbit. In the book chapter whole-cell biosensors, examples are given to demonstrate the special matrices in marine waters and sensors. European funding provided the stimulus for research and development on marine sensors. Out of the Oceans of Tomorrow,1 a network of projects was established.

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https://webgate.ec.europa.eu/maritimeforum/en/node/4073.

Biosensors for the Marine Environment: Introduction

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One such project dealt with the real-time monitoring of Sea contaminants by an autonomous Lab-on-a-Chip biosensor. This project, SEA-on-a-Chip, demonstrated with the applications and progressed the directions and needs for the future. The MariaBox project targeted eight different pollutants in seawater: four biological (azaspiracid, domoic acid, microcystin (structurally related variants), saxitoxin and derivatives) and four chemical (camphechlor, naphthalene, perfluorooctanoic acid, heavy metals). To understand the prevalence of harmful algal toxins in aquatic environment, for example, single chain fragment variable and single chain antibodies were developed to detect the harmful marine toxins (toxins domoic acid and saxitoxin) using recombinant antibody technology. This funding provided a starting point that can and must be built upon to deliver sensitive and selective biosensor technology for the marine environment [3–8].

2 Challenges The seas and oceans are important in terms of food supply, raw material extraction, waste product disposal, transport and recreational use, it is unsurprising that the range of drivers for marine research and monitoring is very wide. The main difference between freshwater, estuarine and marine environments is salinity. Salinity influences the speciation of compounds and chemical bonds that compounds may be form. This raises an important issue: should samples be adjusted to set salinity values for biosensors or should biosensors be conducted at or closest to the actual field salinity. There are only a few marine bioassays, biomarkers and biosensors that are standardised and regularly applied for the assessment of marine and estuarine water bodies and sediments of harbour sites and for dredged materials. Other fields of interest are the monitoring of “hot spots” of increasing offshore and coastal or landbased aquaculture activities with emissions of nutrients and leftovers by the fish cages and/or commercial mussel beds. Consequently, monitoring and warning of harmful toxic algae blooms, biotoxins and quantifying infectious materials are needed [8]. Cell-based or “whole-cell” biosensors have been introduced in different applications such as pharmacology or as an alternative sub-animal experiment or especially in the so-called in situ effects monitoring of sensitive marine environments [9]. The currently available marine biosensors or bio-analytical tools are used as environmental monitoring tools to assess information on early responses of living organisms to environmental stressors and to deliver signals on marine ecosystems damage and pathology due to both man-made and natural pollution. There are only a few marine bioassays available on the cellular level [10]. A valid alternative are the whole-cell marine biosensors with a sensing marine bio-component. An on-site strategy is yet to be developed to measure effects by cells and complex marine bio-analytical systems. Investigating quantitative alterations in immunity of exposed blue mussels can yield relevant information about the relationship between exposure to environmental contamination and susceptibility to infectious diseases.

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Marine applications [2, 3, 11–14] pose a substantial challenge in the robustness required for remote application, but developments in portable medical devices and receptor design suggest that these demands can now be realistically tackled for environmental applications. Test and deployment infrastructure (Fig. 2), floating platforms and mobile platforms are providing capacity for measuring parameters in the sea as sensors are being developed in parallel [15]. Drivers to improve monitoring of coastal and marine environments include: • Urban Waste Water Treatment Directive (Directive 91/271/EEC) in relation to impact of specific point discharges from sewage and industry; • Nitrates from Agriculture Directive (Directive 91/676/EEC) in relation to impact of diffuse agricultural input; • Water Framework Directive (Directive 2000/60/EC of the European Parliament and of the Council establishing a framework for the Community action in the field of water policy in relation to ecological status); • Habitats Directive in relation to the maintenance of habitat integrity (Directive 92/43/EEC on the conservation of natural habitats and of wild fauna and flora); • United Nations Environment Programme (UNEP) Global Monitoring Programme for persistent organic pollutants (POPs); • OSPAR Strategy to Combat Eutrophication in relation to the quality of the marine environment; • Important developments in the European Marine Strategy.2

3 Components of a Biosensor Biosensors have been developed for a large range of analytes and study areas such as the medical field, food analysis, bioterrorism and environmental applications [4, 6, 9, 16]. The first biosensor, developed by Leland Clark in the 1950s [17], combines glucose oxidase with an amperometric oxygen sensor. Since then, there has been significant progress towards using biosensor technology in diagnostic applications, specifically towards point-of-care analysis of biomarkers. Different sensor formats have been designed and some of the developed instruments have been commercialised. However, few of these innovations have been implemented for measurements in the marine environment. A typical biosensor consists of a biological sensing element attached to a physical transducer. Biosensor mechanisms that have been used in marine sensing applications include the use of the recognition elements highlighted in Fig. 1, which are paired to any of the physical transduction methods shown [18–20].

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https://ec.europa.eu/environment/marine/eu-coast-and-marine-policy/marine-strategy-frameworkdirective/index_en.htm.

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Marine Biosensors

Biological recognion element Enzymes: Catalyc transformaon of the pollutant Aptamers or pseudo-natural modalies: short DNA or RNA oligonucleodes for small molecules. Anbodies: Compound or class-specific affinity toward a pollutant e.g. marine toxin domoic acid.

Molecularly imprinted polymers: synthec biorecognion elements using a templated polymer matrix to achieve analyte specificity through paerns of non-covalent bonding, electrostac interacons, or size. Microorganisms: General inhibion of cellular respiraon by pollutant; promotor recognion by specific pollutant Æ gene expression, enzyme synthesis, and catalyc acvity; Idenficaon and enumeraon of microorganisms through immunocapture or DNA sequence hybridizaon

Physical transducer Electrochemical: e.g. Potenometric, amperometric.

Opcal – Electronic: e.g. Surface plasmon resonance.

Opcal: e.g. Absorbance, luminescence, fluorescence. Acousc: e.g. Quartz crystal microbalance, surface acousc wave, surface transverse wave.

Fig. 1 Summary of biosensor recognition elements and detection modes

Disruptive innovation in sensing is enabled using biological recognition. Omic tools (including eDNA technologies) can provide insight into biodiversity, from microbes to the largest of marine mammals. Optical sensors are being designed for small platforms to measure Primary Productivity, and time-resolved fluorometry now makes this feasible even from (Argo) profiling floats [21, 22]. Microsensors and microfluidics are being developed, enabled by the convergence of biology, physics, chemistry and engineering disciplines. These approaches are reported to perform high-quality assays for a very wide range of variables often resulting in better metrology performance when compared with traditional methods [2, 4, 6, 8, 13]. Systems are now meeting demands of low-cost, pressure balancing and utilising low-cost components (such as simple LED-photodiode optics) [11, 13, 14]. Standard component availability and flexibility will facilitate rapid development of new sensors, such as the combination with new robust bioassays to measure new target analytes or new marine variables. There is an opportunity to develop a small number of additional components to adapt and develop microsystems for cytometers (plankton and microplastic analysis with electrical and optical methods) and nucleic acid (NA) analysers. Common components can enable efficiency in, for example, the bioassay Lab on Chip and NA analyser sharing common sample preparation, reagent cost and storage problems and solutions [23]. Biogeochemical LOC can address 5 nutrient essential ocean variables (EOVs) and a complete description of the ocean Inorganic Carbon (carbonate) system. Bioassay-enabled LOC [24] can address selected organic contaminants and biotoxins as a demonstrator of capability for

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Fig. 2 Observation strategies in a marine test and demonstration facility – SmartBay Ireland

marine strategy framework development (MSFD) contaminants, offshore energy and aquaculture/fisheries monitoring. Microcytometers can address particulate matter, microplastics and phytoplankton including direct measurement of volume/biomass and simultaneous taxonomic discrimination. An important aspect of the synergy between the different types of measurements is to get appropriate spatial and temporal resolution of data. Whilst field trips, ships, remotely operated vehicles or submersibles are useful tools to provide spatial coverage or specific point measurements for short periods of time, moorings and bottom landers can provide excellent time series of concentrations in fixed locations (temporal resolution, resolution of trends and episodic events [25] (Fig. 2). Several major oceanographic programmes have delivered large volumes of NA sequence data from the oceans, leading to major scientific discoveries such as new carbon export pathways to the deep sea and an entirely new tree of life. Further, NA sequencing is the only viable technology to assess the state and functioning of the marine microbiome at scale. These are significant advances to understand roles in regulating ecosystem health, food webs, climate and other planetary processes. Sample collection always poses a problem when operating in remote locations however, advances autonomous marine sample acquisition and .preservation enables ecogenomic (metagenomic, metratranscriptomic, metaproteomic) analyses has been seen at the National Oceanographic Centre in Southampton [26]. Advances on the Marine Autonomous Plankton Sampler (MAPS) from the NOC offer high flexibility

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with sample number starting at 42 samples per deployment. Increasing sample number, sample volume and reducing overall system size provide the capability to develop really disruptive marine monitoring technologies [8, 15, 27].

4 The Future of Marine Sensing Among the topics presented in this book, a transformative approach using a recently developed methodology for biosensing applications uses CRISPR-Cas-RPA. CRISPR-Cas has been applied to eDNA for the first time, using a novel isothermal detection method which combines Recombinase Polymerase Amplification with highly specific CRISPR-Cas detection [28]. It has been proven to work for eDNA by applying the technology to the detection of Salmo salar from eDNA samples collected in Irish rivers, where presence or absence had been previously confirmed using conventional field sampling. The significance of this advance is that it can be applied to any known species in the environment and is highly specific. Re-engineered cytometers developed for the medical domain have been redesigned with pressurised, ruggedised housings. Conventional ruggedised cytometers are generally deployed on-ship though some cytometers are capable of long-term moored deployment or submerged short-term experimentation controlled by cables or underwater vehicles. Two leading submersible cytometers are made by CytoBuoy and McLane Labs [1, 29]. In situ chemical analysis can be achieved using relatively new LOC technologies that allow for exceptionally precise liquid handling in challenging ocean environments. They are highly customisable and commercially viable alternatives to the current detection methods. LOC-based platforms designed for environmental monitoring are currently capable of detecting phosphate and Escherichia coli, whilst also offering the capabilities of detecting polycyclic aromatic hydrocarbons (PAHs), endocrine disruptors (EDCs), inorganic ions and heavy metals, by using existing microfluidic techniques [4, 6, 8, 13]. The topics covered here are innovative, disruptive and the next generation of marine monitoring systems. It is clear that a convergence between biological and chemical disciplines with engineering and machine learning is needed in the future to deliver robust biosensors that can generate data to help understand and protect the marine environment.

References 1. Kröger S, Law RJ (2005) Biosensors for marine applications: we all need the sea, but does the sea need biosensors? Biosens Bioelectron 20:1903–1913. https://doi.org/10.1016/j.bios.2004. 08.036 2. Sunagawa S, Coelho LP, Chaffron S, Kultima JR, Labadie K, Salazar G, Djahanschiri B, Zeller G, Mende DR, Alberti A, Cornejo-Castillo FM, Costea PI, Cruaud C, d’Ovidio F, Engelen S, Ferrera I, Gasol JM, Guidi L, Hildebrand F, Kokoszka F, Lepoivre C, Lima-

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Microgravity Changes Membrane Properties and Triggers Bioluminescence in Pyrocystis noctiluca as an Approach for New Biosensor Concepts Jens Hauslage, Ruth Hemmersbach, and Bernd Schierwater

Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Results . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Material and Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Cell membranes with their well-defined chemo-physical properties play a crucial role in cellular functions and are indispensable for maintaining the physiological status of a cell. Recently, gravity-related changes in membrane fluidity and uptake of pharmaceutical substances have been observed in artificially produced membranes of liposomes. In order to test gravity impacts on in vivo cell membranes, we exposed an unicellular organism, the dinoflagellate Pyrocystis noctiluca, to microgravity conditions. P. noctiluca represents a quickly responding bioassay for membrane disturbances. It generates a luminescence signal in response to mechanical stimulation, which correlates to local changes in membrane fluidity. A free-fall of 4.7 s in a drop tower induced a significant luminescent signal demonstrating that even very short microgravity conditions induce alterations at the membrane level in in vivo cells of P. noctiluca. J. Hauslage (✉) and R. Hemmersbach Department of Gravitational Biology, German Aerospace Center, Institute of Aerospace Medicine, Cologne, Germany e-mail: [email protected]; [email protected] B. Schierwater Institute of Animal Ecology, University of Veterinary Medicine Hannover, Hannover, Germany e-mail: [email protected] Fiona Regan, Peter-Diedrich Hansen, and Damià Barceló (eds.), Biosensors for the Marine Environment: Present and Future Challenges, Hdb Env Chem (2023) 122: 11–20, DOI 10.1007/698_2022_942, © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023, Published online: 29 January 2023

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Keywords Bioluminescence, Dinoflagellates, Drop tower, Membrane fluidity, Microgravity, Pyrocystis noctiluca

1 Introduction Membranes are crucial for the functioning of cells, regulate the interaction with the environment, and are mandatory to keep the cellular system functional. Membrane fluidity is an important property of cells to regulate the interactions with the environment. The composition of a membrane with respect to the relative amounts of different phospholipids, proteins, and cholesterol determines its fluidity. Fluctuations of membrane fluidity occur during the development of cells, as has been shown in neurons [1]. The degree of fluidity is also determined by environmental conditions like temperature [2], chemicals [3], and static magnetic fields [4]. Recently, it has been postulated that membranes may play a crucial role in the perception of altered gravity conditions at the subcellular level [5–7]. It has been suggested that gravity is “sensed” by a cell through changes in membrane fluidity, which then triggers differential physiological responses. This hypothesis has been tested so far for artificial membranes and for isolated cell membranes [8]. In parabolic plane flights, providing 31 × 22 s of 10-2 g microgravity and hypergravity conditions of 1.8 g, Sieber et al. demonstrated that membrane fluidity of liposomes and neuronal cells is increased under microgravity and decreased under hypergravity conditions [8]. Similar results were also obtained in drop tower experiments with plain vesicles [9]. Dinoflagellates are known as very fast reporter systems for the identification of shear forces on mechanosensitive ion channels in membranes [10]. When external forces like sheer or hydrodynamic forces interact with the cell membrane of dinoflagellates, a bioluminescence signal is induced by the reaction of luciferase and luciferin mediated by calcium ions [11]. The bioluminescent reaction occurs in scintillions, small caverns in the vacuole where luciferase and a receptor system are located [12]. Recently dinoflagellates were used to visualize shearing forces and membrane deformation, which triggers a bioluminescence response. The number of emitted photons correlates to the amount of deformation induced by shear forces or mechanical impact [13]. It is also known that the bioluminescent reaction in dinoflagellates is triggered by an increase of the membrane fluidity generated by shear forces on the membrane [13]. Application of this fast biosensor enabled us to validate different ground-based facilities which have been designed to simulate microgravity on ground by using different physical principles, e.g. clinostats and random positioning machines. Dinoflagellates showed distinct responses and proved a low shear stress environment in a fast-rotating clinostat (fast and constant rotation around one axis perpendicular to gravity), however a high shear stress situation during operation of a random positioning machine (rotation of samples around two axis with different speeds

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and changing direction) [14]. To further investigate the influence of microgravity on membrane fluidity this reporter system was now used to visualize this impact on cell membranes during transition from earth gravity into microgravity. We here show for the first time in unicellular organisms that changes in membrane fluidity occur very quickly during the transition phase from 1 g to microgravity conditions.

2 Results The following data sets (Figs. 1, 2, 3 and 4) from four drop tower experiments reveal the influence of a stepless and abrupt change from 1 g to microgravity (free-fall) conditions and prove the suitability of Pyrocystis noctiluca not only as a bioassay for identification of shear stress, but also as an indicator for the status of membrane fluidity and finally the operational quality of drop tower experiments for microgravity research. All graphs show the counted photons emitted via bioluminescence during the 4.7 s lasting free-fall in the drop tower, demonstrated by the logarithmic number of counted photons (unit on the left side marked with big dots) versus the given g level (unit on the right side marked with small lines). All drop tower experiments demonstrate identical behavior of the dinoflagellates in microgravity. After start of

Fig. 1 Drop experiment #1 shows a direct increase of the bioluminescence signal from 540 to 6,591 photon counts (Drop at 5.7 s, Impact of drop capsule at 10.4 s). The signal decreases after approx. 3 s to the initial level

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Fig. 2 Drop experiment #2 shows a direct increase of the bioluminescence signal from 92 to 293,418 photon counts (Drop at 7.6 s, Impact of drop capsule at 12.3 s). The signal decreases during the microgravity phase (4.7 s) to 13,882 photon counts

Fig. 3 Drop experiment #3 shows a direct increase of the bioluminescence signal from 197 to 232,217 photon counts (Drop at 4.7 s, Impact of drop capsule at 9.4 s). The signal decreases within the microgravity phase (4.7 s) to 489 photon counts

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Fig. 4 Drop experiment #4 shows a direct increase of the bioluminescence signal from 66 to 149,551 photon counts (Drop at 5.2 s, Impact of drop capsule at 9.9 s). The signal decreases within the microgravity phase to 757

the transition phase from 1 g to μg (10-5–10-6 g) the dinoflagellates show an intense bioluminescent signal which decreases during the time of free falling. The mathematical function of the decrease of the photon signal is nearly exponential. The high number of photons emitted after the impact of the capsule on ground after each drop is clearly visible, also demonstrating the saturation of detecting photons of the photon counting head. In all graphs the sinusoidal oscillation of the experiment platform after the impact of the experiment capsule at the end of the experiment is visible in the acceleration data.

3 Discussion We have shown that dinoflagellates are a highly sensitive and fast reacting biosensor system for monitoring sudden changes in gravity conditions and membrane fluidity. For this Pyrocystis noctiluca was exposed to the drop tower scenario, which provides a continuous transition – without any detectable mechanical impact – from 1 g to microgravity with a high quality of 10-6 g. Previous experiments on membranes have shown that the diameter and the fluidity of asolectin vesicles and subsequently the uptake of substances were altered under microgravity conditions [8, 9, 15]. These data offered new aspects to the question how single cells manage to perceive altered gravity conditions. Furthermore, changes in membrane fluidity also impact the uptake of pharmaceutical

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substances, which will be important for long-term space missions with respect to human health [7]. All this information so far derived from studying artificial or isolated membranes and organelles. Artificially produced vesicle membranes and neuronal cell membranes demonstrated direct impacts on the biophysics of these systems, however, without so far further understanding on the mechanisms. In the current study, living organisms were applied due to their fast-bioluminescent reaction, which is a result of the deformation of the cell and its membrane. In turn, shear forces can be also interpreted as local changes of the fluidity of a membrane. Control experiments with dimethyl sulfoxide (DMSO) as changer for the membrane fluidity result also in a bioluminescent reaction in dinoflagellates [13]. According to the hypothesis that microgravity changes the fluidity of a biological membrane [8], it can be assumed that a dinoflagellate equipped with a shear force sensitive reporter system will change its membrane fluidity. In all drop experiments Pyrocystis noctiluca shows the same course of reaction after start of the microgravity phase in the drop tower: a clear increase in bioluminescence after onset of microgravity followed by an exponential decrease within the drop microgravity time (4.7 s), terminated by a sudden and maximal signal due to the impact of the capsule determined by the saturation of the photon counting head. A mechanical stimulation of the dinoflagellates occurred during filling of the cuvette and in turn, they were allowed to adapt during 1 h before experiment start to get a stable baseline of a bioluminescence signal [14]. Due to head space-free filling of the cuvette a mechanical stimulation during the free fall can be excluded, which also proved that the operational transition from 1 g to microgravity in the drop tower is not accompanied with mechanical stress. Dinoflagellates like Pyrocystis noctiluca produce bioluminescence in specialized caverns in the acidic vacuole called scintillions. After opening, voltage activated proton channels with a luciferin binding protein (LBP) release luciferin and luciferase will be activated by a conformational change. As a result, luciferin reacts with luciferase producing the visible bioluminescent light [12]. An attempt to explain the observed behavior under microgravity is that the acidic vacuole membrane or the voltage activated proton channels are influenced by an increasing of the membrane fluidity. Also, a leaking of protons across the vacuole membrane reaching the LBP and luciferase can trigger a bioluminescent signal. Furthermore, the vacuolar proton channel LpHv1 was identified in bioluminescent dinoflagellates located in isolated scintillions [16, 17]. The fast decrease of the signal during the microgravity phase can be explained as a fast recovery in form of adaptation of the membrane fluidity or a re-conformation of the activity of the permeability of proton channels. Finally, an increase in intracellular calcium, as observed under microgravity in other cells, might also play a role in triggering the bioluminescent reaction of the dinoflagellates [18]. Further microgravity experiments must be performed with the use of channel inhibitors to identify the different signal pathways which are postulated in the bioluminescent reaction [19]. Jin et al. presented a hypothetical model of the bioluminescence signaling pathway in Lingulodinium polyedrum with two different pathways over stretch and voltage activated channels. In further drop tower experiments Nifedipine for inhibiting

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voltage activated channels or gadolinium chloride to inhibit stretch activated channels are candidates to get a deeper insight of the activation of bioluminescence by gravity changes in dinoflagellate. Membranes are mandatory for life, protecting cells against outer impacts and control its communication and metabolism. It becomes more important to investigate the impact of altered gravity on membranes due to human exploration to Moon and Mars for long-term missions. Several effects of reduced gravity are known in astronauts from altered immune reaction, changed uptake of pharmaceutics to slower wound healing. Cell membranes and their abilities to communicate with other cells or taking up signal molecules or drugs are involved in these processes and must be understood for further long-term space missions. Fast reporter systems like Pyrocystis noctiluca are ideal to provide further insights into the behavior of the membrane fluidity and ion channels under the influence of altered gravity in a noninvasive manner.

4 Material and Methods Pyrocystis noctiluca was obtained from the Culture Collection of Algae of the University of Cologne. The dinoflagellates were cultured in T75 cell culture flasks in F/2 seawater culture media without silicate solution within an incubator with 24°C and a light:dark cycle of 12:12 [20]. The bioluminescence based on a circadian rhythm and was shifted in a way that the bioluminescence which is active approx. 1 h after beginning of the dark starts in the morning hours around 8:00 a.m. All experiments started after 8:00 a.m. in the morning to synchronize the ability of the cells to emit bioluminescent light [14, 21]. During the experiments in the drop tower, dinoflagellates were filled head-space free in a small cylindrical glass cuvette (3 ml volume) with an inner diameter of 4 mm sealed with a rubber plug equipped with a small tube to avoid overpressure. This prevents also bubbles inside of the cuvette. Due to a relative measurement of the emitted bioluminescence during the drop tower experiments, a counting of the cell density was neglected. Before filling the cuvette, the cell culture flask was slightly rotated to mix the media and previously sedimented cells to gain a homogenous distribution of the cells in the media. Filling of the cuvette produces a mechanical stress response and results in a bioluminescence reaction. Thus – after filling the cuvette – approx. 1 h was given for adaptation and to achieve a stable baseline of the bioluminescence level [14]. All experiments were performed in the first 6 h after the artificial dusk controlled by a shifted illumination cycle. This procedure prevents glowing of the dinoflagellates and ensures that the cells will flash after stimulation [21, 22]. The bioluminescent light emitted from the dinoflagellates was documented by means of a photon counting head (Hamamatsu H10682) within a light tight box in front of the experiment cuvette with dinoflagellates (Fig. 5).

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Fig. 5 Experiment setup in the drop tower capsule. (a) Upright profile from the drop tower capsule. (b) 80 mm bracket as support for the measurement unit. (c) Soft foam cube to decouple the measurement unit from mechanical influences during the experiment. (d) Photon counting head. (e) Light tight cylinder with cuvette (white). The arrow indicates the fall direction

The photon counting head sends a single signal for each of four detected photons. These counts were transmitted via an Arduino microcontroller system and a serial connection to a data recording system in the drop tower facility. A counter program using the FreqCounter library from Martin Nawrath from the Academy of Media Arts Cologne was implemented on the Arduino. With this program, the Arduino is able to count frequencies of up to 8 MHz (interface.khm.de/index.php/lab/interfacesadvanced/Arduino-frequencycouter-library/). The photon count was measured with 10 Hz. An additional acceleration sensor (MMA8451; Adafruit Industries) was installed beside the cuvette to record the gravity level during the experiment. To avoid mechanical stimulation during the release of the experiment capsule inside the drop tower the photon counting head with the glass cuvette was mounted in a cube of 3 cm layers of soft foam material (Fig. 5). Ground-based control experiments: The decoupling of the photon counting head was manually tested by giving a strong impulse on the capsule structure (Part A in Fig. 5) via a hammer stroke and simultaneous observation of the counted photons. No bioluminescent reaction was observed directly after the hammer stroke. This ensures that the mechanical stimulation of the release impact was decoupled from the experiment structure. Furthermore, as control, a light mechanical hit was performed directly on the photon counting head with a filled cuvette which induced a high bioluminescent signal in a comparable pattern as observed by the impact of the landing hit of the experiment capsule after the drop (data not shown). Acknowledgments Cell cultures from Pyrocystis noctiluca were obtained from the Cell Culture Collection of Algae at the University of Cologne, Laboratory Melkonian. We also thank the ZARM Team for a perfect support during the drop tower campaign.

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Author Contributions J.H., B.S., and R.H. have designed the study. J.H. built the hardware and performed the drop tower experiments. J.H., B.S., and R.H. wrote the manuscript. All authors read and approved the final manuscript. Competing Interests The authors declare no competing interests. Funding The experiments were funded by the German Aerospace Center and the German Space Agency.

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Addressing Ciguatera Risk Using Biosensors for the Detection of Gambierdiscus and Ciguatoxins Greta Gaiani, Jorge Diogène, and Mònica Campàs Contents 1 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2 Gambierdiscus and Fukuyoa Global Distribution with Particular Focus on the Mediterranean and Macaronesian Regions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 Ciguatoxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 4 Methods for Gambierdiscus and Fukuyoa Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 5 Methods for Ciguatoxin Detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 Biosensors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6.1 Biosensors for the Detection of DNA from Gambierdiscus . . . . . . . . . . . . . . . . . . . . . . . . . . 6.2 Biosensors for the Detection of Ciguatoxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Ciguatera fish poisoning (CFP) is one of the most relevant seafood-borne diseases worldwide. It is caused by the ingestion of seafood contaminated with ciguatoxins (CTXs), potent marine neurotoxins produced by microalgae of the genera Gambierdiscus and Fukuyoa. These genera are endemic of the tropical and subtropical areas of the world, but in recent times are being found more and more in temperate areas, such as the Mediterranean Sea. In order to properly assess and manage CFP risk, the detection of Gambierdiscus/Fukuyoa species and the discrimination between toxic and non-toxic fish specimens are crucial. To tackle these challenges, biosensors have recently been developed for the detection not only of CTXs in fish and microalgae samples but also of the DNA of the responsible microalgae in the environment. These tools are sensitive, specific, rapid, robust, reliable, easy to use, and they could represent a step forward towards in situ detection, helping in the CFP risk management and protecting human health. G. Gaiani, J. Diogène, and M. Campàs (✉) IRTA, La Ràpita, Spain e-mail: [email protected] Fiona Regan, Peter-Diedrich Hansen, and Damià Barceló (eds.), Biosensors for the Marine Environment: Present and Future Challenges, Hdb Env Chem (2023) 122: 21–46, DOI 10.1007/698_2022_943, © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023, Published online: 29 January 2023

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Graphical Abstract

Keywords Ciguatera, Ciguatoxins (CTXs), DNA sensor, Gambierdiscus, Immunosensor

1 Introduction Ciguatera fish poisoning (CFP) is a foodborne disease that causes gastrointestinal, cardiological, and neurological symptoms that can last weeks, months, or even years, and in some cases lead to death [1]. It is caused by the ingestion of seafood containing ciguatoxins (CTXs), a group of cyclic polyether lipophilic compounds produced by microalgae of the genera Gambierdiscus and Fukuyoa, which accumulate into fish flesh and through the food webs. In the CFP process, more than 400 fish species and also sharks are implicated [2, 3], some being more likely to contain CTXs than others [4]. Moreover, it seems that several species of marine invertebrates, such as sea urchins [5], lobsters and octopuses [6], giant clams [7, 8], and sea stars [9], may be involved in CFP pathways. The discrimination between contaminated and uncontaminated specimens is an important challenge, since toxic specimens do not look, smell, or taste any differently from non-toxic ones. This issue has led, in areas endemic for CFP, to the decrease in fish consumption [10], drastic modification of dietary habits [7], and ban of the sale of certain high-risk species, causing important financial losses [11]. Furthermore, in 2020, the International Association for Medical Assistance to Travelers (IAMAT) labeled several countries as “ciguatera at-risk destinations” [12]. CFP is an extremely complex phenomenon to manage, from the detection of CTXs in natural samples to the diagnosis in patients (although some symptoms, such as allodynia, are specific of CFP, some others are common in different food poisonings). Thus, taking into consideration that an antidote for CFP has not been found yet, the efforts of the scientific community must focus on the prevention, by

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providing rapid and reliable tools for the detection not only of CTXs in fish and microalgae samples, but also of Gambierdiscus and Fukuyoa cells in the environment. In this chapter, the development and applicability of such tools will be described and discussed.

2 Gambierdiscus and Fukuyoa Global Distribution with Particular Focus on the Mediterranean and Macaronesian Regions Until 1995, Gambierdiscus was considered as a monotypic taxon with just one species named G. toxicus [13]. Further studies over the past decade have resulted in the identification of 18 Gambierdiscus (G. australes, G. balechii, G. belizeanus, G. caribaeus, G. carolinianus, G. carpenteri, G. cheloniae, G. excentricus, G. holmesii, G. honu, G. jejuensis, G. lapillus, G. lewesii, G. pacificus, G. polynesiensis, G. scabrosus, G. silvae, and G. toxicus) [14–20] and 4 Fukuyoa species (F. paulensis, F. ruetlzeri, F. yasumotoi, and F. koreensis) [21, 22]. Among them, 14 species (G. australes, G. balechii, G. belizeanus, G. caribaeus, G. carolinianus, G. carpenteri, G. excentricus, G. pacificus, G. polynesiensis, G. scabrosus, G. silvae, G. toxicus, F. paulensis, and F. ruetlzeri) are considered able to produce CTXs [23] with different tests and techniques [16, 24–29]. Gambierdiscus and Fukuyoa genera are endemic of the subtropical areas of the world (35°N and 35°S) [30]. In more recent times, they have been identified in temperate areas such as Korea [31], Japan [17], New Zealand [18], Gulf of Mexico [26], coast of North Carolina [15], Brazil [21], and also the Macaronesia region [16, 32, 33] and the Mediterranean Sea [34–38] (Fig. 1). The region that hosts the major diversity of Gambierdiscus and Fukuyoa species is the Pacific, where the presence of 19 out of the 22 currently recognized species has been reported. Thus, it is not surprising that several archipelagos of this region, such as French Polynesia and Cook Islands, are identified as biodiversity “hotspots” of Gambierdiscus [30]. In addition, the Caribbean region also presents a huge variety of Gambierdiscus and Fukuyoa species, and it is quite common to find the co-occurrence of five or six species [39]. The fact that G. excentricus and G. silvae, two of the highest CTX-producing species, are not as frequently found as other species draws the attention. The patchy distributional pattern observed has been related to their thermal tolerance [30]. Another curious circumstance is that of G. australes, which is globally distributed with the exception of the Caribbean region, and the reasons behind are still unknown. Different is the situation in the Indian Ocean, where the distribution of the genera is poorly reported, especially in the coastal areas of Africa. Additionally, most records reported the species as G. toxicus, since the identification was mainly performed with microscopy techniques [40, 41]. Although the presence of G. australes and G. belizeanus has been recently proved with molecular methods [42], more studies are needed to have an appropriate species composition of this

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Fig. 1 Gambierdiscus and Fukuyoa distribution in Mediterranean and Macaronesian waters. Symbol * indicates the places (Greece and Cyprus) where the presence of both Gambierdiscus and Fukuyoa was reported only at genus level. For the global distribution, see Tester et al. [49]

region. Another recently found “hotspot” of Gambierdiscus is the northern Macaronesian region, with the Canary Island hosting the highest biodiversity and the highest number of CTX-producing species (G. australes, G. caribaeus, G. carolinianus, G. excentricus, and G. silvae) [16, 27, 33, 43, 44]. Hence, the variety of species found made Rodriguez and coworkers [43] think that Gambierdiscus settlement in the region can date back to ancient times. Actually, a first report of Gambierdiscus in Cabo Verde can be attributed to Silva [45], initially reported as Goniodoma [16]. Recently, Soler-Onís and coworkers [46] identified several cells of G. excentricus in the waters of the Cabo Verde archipelago, confirming the presence of the genus. Even though the settlement of a Gambierdiscus and Fukuyoa species and the finding event can be separated by several decades, there is a general concern that the geographic range of these two genera, and especially of the CTX-producing species, will expand as a consequence of the rise of sea surface temperature [47]. According to Parsons and coworkers [48], a significant modification in the distribution and the abundances of ciguateric microalgae species is to be expected, with some species becoming more dominant over others.

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3 Ciguatoxins CTXs are secondary metabolites produced by Gambierdiscus and Fukuyoa. CTXs are cyclic polyether compounds with a rigid structure formed by 13–14 rings connected with ether bonds. CTXs target the binding site 5 of the voltage-gated sodium channels (VGSCs) [50], inducing effects at the cellular and physiological levels, such as membrane excitability, release of neurotransmitters [51], increase of intracellular calcium [52], and blockage of voltage-gated potassium channels [53]. The affinity of the different CTX congeners for the binding site of the VGSCs is proportional to their toxicity in mice [54]. Up to date, 34 CTX congeners have been described and grouped in Pacific (P-CTX) (22 congeners), Caribbean (C-CTX) (12 congeners), and Indian (I-CTX) (no congeners described yet), according to their geographical origin [28]. CTX1B (P-CTX-1) was the first one to be identified in 1990 by Murata and coworkers [55], followed by the description of many other congeners. To classify the different P-CTX congeners, Legrand and coworkers [56] proposed to distinguish them into two different groups according to the number of carbons and the structure of the E ring (7 in the CTX1B group and 8 in the CTX3C group) and to the presence (CTX1B) or absence (CTX3C) of the 4-carbon side chain of the left wing (Fig. 2).

Fig. 2 Structure of the two main groups of CTX congeners: CTX1B and CTX3C

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Afterwards, two CTXs from the Caribbean Sea (C-CTXs) were isolated by Vernoux and Lewis [57] and identified structurally in 1998 [58]. Subsequently, other congeners were identified by Pottier and coworkers [59]. Six Indian Ocean CTXs (I-CTXs) were also isolated [2, 60], but their structural determination remains undescribed. Alongside with the CTXs bioaccumulation in fish flesh and through the marine food webs, CTXs undergo metabolization processes in fish [61], resulting in more toxic compounds, as observed in fish samples from the Pacific area [62]. The ratio of these different toxins in fish and microalgae samples can vary. Nevertheless, CTX1B (P-CTX-1) is found as dominant in the toxin profiles of the carnivorous fishes of the Pacific [63]. Toxicity assays in mice (intraperitoneal, i.p.) have provided LD50 (median lethal dose) values of 0.25, 2.3, and 0.9 μg/kg for P-CTX-1, P-CTX-2, and P-CTX-3, respectively [63], classifying CTXs as extremely potent marine toxins. Generally, P-CTXs are more potent than C-CTXs (LD50 of 3.6 and 1 μg/kg for C-CTX-1 and C-CTX-2, respectively) and I-CTXs (5 μg/kg). In humans, it has been estimated that no more than 1 ng of P-CTX-1 per kg of body weight is needed to cause the occurrence of mild CFP symptoms [64]. Moreover, these toxins are heat resistant, so they cannot be inactivated by cooking processes [65]. The United States Food and Drug Administration (US FDA) proposed guidance levels of ≤0.01 μg/kg of CTX1B (P-CTX-1) and ≤0.1 μg/kg of C-CTX-1 equivalent toxicity in fish, and these values represent the only existing suggested thresholds. In fact, New Zealand and Australia provide general guidelines on possible ciguateric fish species and areas [66], and Japan has banned several fish species associated with ciguatera [67, 68]. In European markets, no fish products containing CTXs can be sold [69], but no regulatory limits have been established and no suggestion regarding the analytical methodology to use is given. Nevertheless, the European Food Safety Authority (EFSA) has adopted the US FDA guidance levels for CTXs [70]. Therefore, the development of rapid, reliable, and easy-to-use tools able to detect and quantify such low CTX amounts can be of outmost help for CFP management.

4 Methods for Gambierdiscus and Fukuyoa Detection The presence of Gambierdiscus and Fukuyoa species in an area is likely to contribute to the toxicity of fishes. Therefore, the detection of these microalgae species in the field, which might not be dominant in terms of abundances, but whose contribution to the environmental flux of CTXs is noticeable, is of extreme importance [28]. The most known and used technique is light microscopy. However, it is time consuming, requires trained personnel, and usually does not allow identification at the species level. Hence, the use of molecular methods is almost mandatory to correctly identify to the species level [71]. In fact, in every study concerning the new record of a species in a place, the confirmation with sequencing is highly requested to support the findings obtained with other methods. Among the existing sequencing procedures, certainly the Sanger sequencing has been the most used since its invention in 1977. Other molecular techniques are more and more used for

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the identification of microalgae species in field samples. In fact, quantitative polymerase chain reaction (qPCR), with primers targeting the D1-D2 region [72], the D1-D3 region [20, 73, 74], or the D8-D10 region [20, 75] of the large subunit (LSU) ribosomal gene, has been used for the identification and quantification of Gambierdiscus/Fukuyoa genera, G. belizeanus, G. caribaeus, G. carolinianus, G. carpenteri, G. ruetzleri, G. australes, G. scabrosus, G. excentricus, G. silvae, G. lapillus, and F. paulensis. This method is specific, robust, and reliable, but it requires the use of a thermocycler, which hampers in situ analysis, and it is time consuming. As a solution to these drawbacks, our group used an isothermal DNA amplification technique that allowed obtaining, in 30 min and at a constant temperature of 37°C, the corresponding amplicons. The use of primers modified with oligonucleotide tails resulted in an amplified product flanked with those tails able to be detected in a sandwich hybridization assay, thanks to the use of complementary capture probes and a horseradish peroxidase (HRP)-labeled reporter probe. With this strategy, it was possible to differentiate between Gambierdiscus/Fukuyoa and other microalgae genera and also between two CTX-producing species (G. australes and G. excentricus) and their congeneric species [76]. Despite its advantageous features, this technique is not commonly used, mostly because the kit used to perform the test (commercially available for 96 reactions) is expensive and its efficiency gets lower and lower once opened, compromising the inter-day reproducibility. Meanwhile, researchers have focused on the development of PCR-based techniques that could be implemented into small and portable devices for the in situ detection of microalgae. Recently, a PCR-lateral flow assay (PCR-LFA) has been developed targeting the same two CTX-producing microalgae species [77]. The assay requires less than 1.5 h to be performed, PCR included. Again, tailed primers are exploited and specific detection is achieved with the aid of capture probes and single-chain Cro proteins conjugated with carbon nanoparticles used as labels. This technique represents a step forward towards in situ analysis, although further studies, including the screening of natural samples, need to be performed. The advantage of using tailed primers is undoubtable, and for this reason this strategy has been chosen to develop biosensors.

5 Methods for Ciguatoxin Detection A huge variety of methods have been developed to detect CTXs. Up to date, the techniques developed include animal assays, cell-based assays (CBAs), receptorbinding assays (RBAs), immunoassays, and instrumental analysis [62, 78–80]. Several animal assays have been developed throughout the years, but only one is still in use for the detection of CTXs: the mouse bioassay (MBA) [81]. The MBA is useful because it provides a composite toxicological response, which is very convenient in case of samples with unknown toxicity [70]. However, it is neither specific nor highly sensitive and suffers from ethical problems. Therefore, some laboratories switched to the use of assays based on mammalian cells, instead of entire animals.

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The CBAs developed for CTXs are based on the activity of these toxins on neuronal VGSCs [82] and involve a huge assortment of cells and tissues, from blood, used for the development of hemolytic assays [83], guinea pig ileums [84], guinea pig atrium [85], frog nerve fibers [86], and crayfish nerve cords [87]. Despite this variety of materials, nowadays the most used CBA is the one based on mouse neuroblastoma (N2a) cells [88]. This CBA has demonstrated to be very sensitive and provides a composite toxicity response for the several existing CTXs. Additionally, the assay is relatively easy to perform and interpret. Briefly, it is based on the colorimetric detection of metabolically active N2a cells exposed to CTX in presence of ouabain/veratridine [87]. The detection of CTXs requires the addition of veratridine, that is a VGSC activator with a different binding site than CTX, and ouabain, a sodium/potassium pump inhibitor. The combined effect of these three substances together increases the concentration of intracellular sodium, which has a negative effect on cell viability and can be measured as a function of CTX concentration. The amount of toxin is measured using 3-(4,5-dimethylthiazol-2-yl)-2,5diphenyltetrazolium bromide (MTT), which is reduced by mitochondrial dehydrogenases into a formazan product, which is later solubilized and whose absorbance intensity is proportional to the number of live cells and so inversely proportional to the concentration of CTX, within a certain range. Even if the limit of quantification (LOQ) is different in each experiment, in general it is lower than the FDA suggested threshold. However, a consensus protocol for the MTT-based CBA is still lacking since throughout the last decade users made customized changes to the assay [89]. The modifications included almost every aspect of the test, such as the cell seeding densities, the MTT incubation time, and the ouabain/veratridine treatment. Other N2a CBAs, with no MTT, have been developed. For example, in the study of Fairey and coworkers [90], they used N2a cells that expressed c-fos-Luciferase reporter gene. The c-fos is a response gene and a sensitive biomarker to localize the effects of toxins. Detection and quantification are achieved with luciferasecatalyzed light generation and a luminometer. Additionally, cell lines other than N2a have been exploited for application in CBA. In particular, the human neuroblastoma cell line SH-SY5Y has been used to develop another fluorescence assay [91]. In this assay, cells were loaded with a dye containing calcium adsorbed into the cytoplasm and then incubated with veratridine and subsequently with CTXs. Fluorescence responses to CTXs were measured as the increase of calcium ion influx into cells with a plate reader. These fluorescence assays are not commonly used due to the cost of the fluorescent dye, the need of specialized equipment, and the sensitivity to maitotoxin presence, which can affect the outcome of the test. Indeed, the interfering effects caused by maitotoxins, other toxic compounds or the natural matrix itself (i.e., components from the fish or microalgae extracts) can induce an over or under estimation of the CTXs content. Additionally, CBAs respond similarly to other toxins that block VGSCs (i.e., brevetoxins) and, therefore, it is impossible to distinguish one from another. RBAs, also based on the affinity of CTXs for their binding site on the VGSCs, have been developed. Although in an RBA the response is structure-related, the fact that VGSCs are targeted may involve correlation with toxicity. Since CTXs share

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with brevetoxins the same binding site on the VGSCs (i.e., binding site 5) but with a higher affinity, they can be considered as brevetoxin binding competitors [90, 92]. Therefore, measuring the competition binding of a radioactively labeled brevetoxin ([3H]-brevetoxin-3) and CTXs for the receptor sites in a membrane can be used to estimate the amount of CTXs in an extract. Hence, the concentration of the labeled brevetoxin (that is added constant at all wells) should decrease after the addition of CTXs, and a competition dose-response curve can be obtained. This screening method has been widely used, but it is highly sophisticated, and the comparison between laboratories is quite complicated. Thus, Díaz-Asencio and coworkers [93] made the effort to provide guidance on its quality control checks for the analysis of environmental samples, reaching a limit of detection (LOD) of 0.75 ng/g of P-CTX-3C in fish samples in their optimized assay. However, RBAs involve the use of radioactive compounds. To avoid their use, a fluorescence-based RBA has been developed, where CTXs compete with a brevetoxin-2 labeled with a fluorescent dye [94]. This assay provided biding values that correlated with toxicity values, attained an LOD of 0.075 ng/g of P-CTX-3C and could be performed in approximately 2 h. A commercial kit for CTXs based on this study has been marketed by Sea Tox Research Inc. (Wilmington, NC, USA https://www. seatoxresearch.com/testing-kits/) and can be used as screening tool for fish extracts. Despite the undoubtable utility of the kit described above, it does not allow to know which CTXs are inside a sample. The best solution to obtain toxin profiles is to separate the toxins and high-performance liquid chromatography (HPLC) is the method to perform this task. Since most CTXs do not have a characteristic chromophore group in their structure (i.e., alternating single and double bounds), they do not strongly absorb radiation over the UV/Vis region, and therefore spectroscopy is not viable for their detection. Indeed, the trials with the classical HPLC method with UV detector showed not enough sensitivity to detect the presence of low concentrations of CTXs [95]. Therefore, the HPLC with fluorescence detection has been tried, since some CTX congeners have a primary hydroxyl group available for fluorescent labeling. Even if this technique showed better sensitivity than the previous one, it does not detect CTXs at the recommended level (0.01 μg/kg). Additionally, it does not detect CTXs without a primary hydroxyl group (i.e., P-CTX-3C). Therefore, to increase the sensitivity and specificity of the system, Lewis and Jones [96] combined the HPLC technique with tandem mass spectrometry (HPLC-MS/MS) for the detection of CTXs. Then, Lewis and coworkers [97] combined an electrospray triple quadrupole mass spectrometer with a gradient reverse-phased HPLC and, with this technique, LODs of 0.04 μg/kg and 0.1 μg/kg for P-CTX-1 and C-CTX-1, respectively, were achieved [97]. Right after this first trial, LC-MS/MS becomes one of the most used, if not the most used, techniques to detect and quantify CTXs. It must be underlined that CBA is the most used technique to perform sample screening, even though LC-MS/MS is the one that may confirm the presence of CTXs. Although instrumental analysis techniques are highly sensitive, their application to monitoring programs is hampered by the cost of the machinery, the time needed to prepare the samples, and the need of highly trained personnel to perform the analysis.

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Additionally, the analysis of CTXs in natural samples is limited by the lack of CTX standards, certified materials, and the chemical complexity of the CTXs. These limitations have also hindered the development of immunoassays, based on antibodies (Abs). These assays take advantage from the high specificity of the antigen–antibody interaction. The first group to produce anti-CTX Abs was the one of Hokama and coworkers [98]. In their work, they produced an anti-CTX polyclonal Ab (pAb) and labeled it with a radioactive compound to subsequently perform a radioimmunoassay directly on fish tissues from the Hawaiian Islands [98, 99]. The same pAb was labeled with HRP and exploited in an immunoassay also for fish extracts [100]. The authors decided to simplify the enzyme immunoassay by formatting it into a faster stick test that did not require any instrumentation [101]. These last findings were used to build two commercial kits named CiguaCheck [101, 102] and Ciguatect [103]. Although these achievements represent an advance in the development of easy-to-use tests, since no extraction whatsoever was needed to perform the assay, these kits showed cross-reactivity with okadaic acid and brevetoxin [102, 104]. This cross-reactivity together with the low sensitivity led to false positive and false negative results, respectively [105]. Therefore, the only fish reported as ciguateric up to date in the Mediterranean, which was analyzed with the Cigua-Check kit [106], is still pending of confirmation. Due to the disadvantages observed when using these pAbs, Hokama and coworkers [107] decided to focus on the production of monoclonal antibodies (mAbs) using purified CTX, that were subsequently exploited in a similar system but using colored latex beads as labels. Results were promising but even the authors claimed that refinement was necessary. Another approach to produce mAbs is the one based on the use of synthetic haptens instead of natural CTXs. The first work using this strategy is the one of Campora and coworkers [108], which developed a sandwich enzyme-linked immunosorbent assay (ELISA), using one specific mAb for the left wing of P-CTX-1 and one specific mAb for the right wing labeled with HRP. No cross-reactivity was observed with other marine toxins such as okadaic acid or domoic acid, and only very slightly with brevetoxin-3. Subsequently, Tsumuraya and coworkers [109–113] immunized mice with haptens that mimic the left and right wings of the four principal congeners of Pacific CTXs: CTX1B, 54-deoxy-CTX1B, CTX3C, and 51-hydroxy-CTX3C. The resulting mAbs were used to develop colorimetric sandwich ELISAs. The immunoassays demonstrated the high specificity and sensitivity that was expected, showing no cross-reactivity with other marine toxins such as okadaic acid, maitotoxin, brevetoxin A, and brevetoxin B. Additionally, a fluorescence ELISA was developed, whose LOD was of less than 1 pg/mL for both CTX1B and CTX3C [114, 115]. Moreover, CTX1B was spiked into a fish extract at the suggested threshold of 0.01 μg/kg and successfully detected. Based on this fluorescence assay, a kit named “CTXELISA 1B” for the detection of CTXs from the CTX1B group has been marketed and can be bought from Fujifilm Wako Corporation (Osaka, Japan). Since the results obtained with this strategy are very encouraging, the mAbs produced by the group of Tsumuraya were exploited in the development of biosensors.

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6 Biosensors Biosensors are practical and reliable tools to detect biological and chemical hazards. They are composed of a biorecognition element that interacts specifically with a target molecule, and a transducer that converts the biorecognition event into a quantifiable signal, both in intimate contact. The biorecognition element is the one that gives specificity to the system, and it could be an enzyme, antibody, oligonucleotide, aptamer, receptor, cell, tissue, or microorganism. The transducer can be electrochemical, optical, gravimetric, or thermometric, according to the type of signal they transform in a measurable unit. Even if biosensors represent an interesting and useful tool for the detection of different type of analytes, they have been rarely used to detect DNA of toxin-producing microalgae. This has been the case for Karenia brevis [116], Karlodinium armiger [117], Ostreopsis ovata [118] and for some species of Gambierdiscus (G. australes, G. excentricus, and G. silvae), Coolia (C. monotis, C. tropicalis, and C. cf. canariensis), Ostreopsis genus, and Prorocentrum lima [119]. On the contrary, several biosensors have been developed for the detection of marine toxins, such as surface plasmon resonance immunosensors (palytoxins, tetrodotoxins), surface plasmon resonance receptorbased biosensors (palytoxins), electrochemical immunosensors (tetrodotoxins, okadaic acid, azaspiracids, domoic acid, saxitoxins, palytoxins, brevetoxins), electrochemical enzyme-based sensors (okadaic acid), electrochemical aptasensors (okadaic acid, brevetoxin-2, saxitoxin, tetrodotoxins), electrochemical cell-based biosensors (palytoxin), and electrochemiluminescence immunosensors (palytoxins) (for more details, see [78, 120, 121]). In the following paragraphs, the recently reported biosensors for the detection of DNA from Gambierdiscus and CTXs will be described and commented (Table 1).

6.1

Biosensors for the Detection of DNA from Gambierdiscus

The current increase in the reports of Gambierdiscus species in Mediterranean and Macaronesian waters [34, 37, 122] has raised the need to have appropriate tools to detect these species in the field. In recent years, rapid and reliable molecular-based biosensors for the detection and enumeration of marine microalgae species have been developed as an alternative to the traditional light microscopy technique. In the first work, Medlin et al. [119] designed probes for the detection of several species of the genera Gambierdiscus, Ostreopsis, Coolia, and Prorocentrum. These probes were used in the development of hybridization assays, where streptavidin-coated magnetic beads (MBs) were used for capture probe immobilization and an HRP-labeled anti-FITC Fab fragment was used to recognize the FITC-labeled sandwich DNA/DNA (for the detection of DNA) or a ProtA-polyHRP was used to recognize an Ab able to interact with RNA/DNA hybrids (for the detection of RNA).

Target G. australes G. excentricus G. silvae

G. australes G. excentricus

CTX1B 54-deoxyCTX1B CTX3C 51-OH-CTX3C

CTX1B 54-deoxyCTX1B CTX3C 51-OH-CTX3C

Biosensor type Electrochemical DNA-based sensor

Electrochemical DNA-based sensor

Electrochemical immunosensor

Electrochemical immunosensor

Abs immobilized on multiwalled carbon nanotubes-modified screen-printed carbon electrodes

Immobilization support Biotinylated capture probes conjugated to streptavidincoated MBs and immobilized on screen-printed carbon electrodes Thiolated capture probes conjugated to maleimide-coated MBs and immobilized on screen-printed carbon electrodes Abs conjugated to carboxylic acid-modified MBs and immobilized on screen-printed carbon electrodes Amperometry

Amperometry

Amperometry

Electrochemical technique Amperometry

Table 1 Existing biosensors for the detection of Gambierdiscus species and CTXs

6 pg/mL of CTX1B

1.96 pg/mL of CTX1B 3.59 pg/mL of 51-OH-CTX3C

10 cells

LOD 1 pM of RNA (10–444 cells)

Genomic DNA of microalgae cultures; field samples from the Balearic Islands Fishes from La Réunion Gambierdiscus/Fukuyoa cultures Field samples from the Balearic Islands Fishes from Japan and Fiji

Samples RNA or synthetic DNA

[124]

[122]

[123] [125]

[122]

Reference [119]

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Fig. 3 Schematic representation of the biosensor developed for the dual detection of G. australes and G. excentricus [122]. Field samples collection (a) was followed by a fast DNA extraction technique (b) and amplification with a multiplex PCR. Amplified products were exposed to the capture probes specific for G. excentricus (c1) and G. australes (c2) conjugated with maleimidecoated magnetic beads and to an HRP-labeled reporter probe. Electrochemical detection was achieved by immobilizing the oligocomplexes on the working electrodes of a dual array, adding TMB substrate, and measuring the reduction current with amperometry (d)

Oligocomplexes were immobilized on the surface of screen-printed carbon electrodes (SPCEs) with the use of a customized case containing a magnet placed underneath the working electrode and immersed in an electrochemical cell containing 1 mM hydroquinone (HQ) under constant agitation. The reduction current intensity, arising from the enzymatic reduction of H2O2 mediated by HQ and measured with amperometry, was proportional to the concentration of the RNA/DNA target and, consequently, to the number of microalgae cells. The specificity towards G. australes, G. excentricus, and G. silvae was tested. The LOD achieved was close to 1 pM of RNA, but no corresponding quantification in terms of cell abundance was established. Nevertheless, for the other dinoflagellates targeted in the study, the number of cells corresponding to 1 pM of RNA ranged from 10 to 444 cultivated cells, so probably the LOD for Gambierdiscus species would be similar or even lower. The technique presented by Medlin and coworkers is fast, since no PCR step is performed (although it would be necessary to detect DNA from field samples), cost-effective, and reliable, but it has been tested only with synthetic DNA and it has not been applied yet to the analysis of genomic DNA/RNA or the screening of field samples. More recently, our group [122] described the first electrochemical biosensor for the simultaneous detection of the two CTX-producing species G. australes and G. excentricus (Fig. 3). Like Medlin and coworkers [119] for DNA, we used a sandwich configuration, although the strategy was different. We designed speciesspecific capture probes for G. australes and G. excentricus, which were immobilized on the surface of maleimide-coated MBs. Subsequently, PCR was performed using primers for both G. australes and G. excentricus at the same time. These primers included oligonucleotide tails that, after PCR, gave amplified products flanked with single-stranded oligonucleotides at each end. Afterwards, the amplified products were incubated with the MBs modified with the specific capture probes, and then an

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HRP-labeled reporter probe, common to G. australes and G. excentricus, was added. For the electrochemical detection, the G. australes oligocomplexes were immobilized on one of the working electrodes of a dual electrode array, and the G. excentricus oligocomplexes were immobilized on the other one. As in Medlin et al. [119], a customized plastic case with magnets placed underneath the working electrodes was used for MBs immobilization. Nevertheless, our electrodes were used in a horizontal configuration, which requires smaller sample volumes to operate. Finally, 3,3′,5,5′-tetramethylbenzidine (TMB) Enhanced One Component HRP Membrane Substrate was added and incubated for 10 min. The use of this compound allowed the simultaneous detection of amplified DNA on both electrodes because, when oxidized, it precipitates on the electrode on which the chemical reaction is happening, not interfering on the other one. The reduction current intensity, measured with amperometry, was proportional to the amount of amplified product and consequently to the number of microalgae cells present in the samples. To determine the LOD of the system, calibration curves were constructed starting from a 104-cell pellet, performing 1/10 serial dilutions and extracting each pellet dilution with a newly developed fast technique, based on a portable bead beater device and combined with specific MBs for DNA capture. Since the DNA extraction part is crucial, in order to assess its efficiency, DNA dilutions (i.e., extraction of DNA from a 104 cell pellet, and subsequent 1/10 serial dilutions of the extract) were also tested and compared. In both cases, an LOD of 10 cells was reached for the target species. Additionally, the simultaneous detection of these two species at different cell concentrations and ratios was successfully achieved. Finally, field samples collected in Majorca (Balearic Islands, Spain) were screened, and the abundances obtained with the biosensor were similar to the estimations provided by light microscopy. In comparison with Medlin et al. [119], the strategy presented by our group is longer and less species are targeted. Nevertheless, it allowed not only the detection of genomic DNA extracted from laboratory cultures, but also the screening of field samples, providing the first report of G. excentricus in the Balearic Islands. The detection strategy combined with the fast extraction technique represents an important step forward towards the practical application of biosensors for in situ detection of CTX-producing species. Moreover, this system can be easily modified for the simultaneous detection of other microalgae species (pending the design of tailed primers that do not cross-react with each other), and therefore it could be useful for the assessment of other marine-related diseases.

6.2

Biosensors for the Detection of Ciguatoxins

The availability of sensitive, specific, rapid, reliable, and easy-to-use tools for the screening and quantification of CTXs in natural samples is of extreme interest, not only for scientific purposes but also to promote the sustainability and safety of fisheries and guarantee public health. In the next paragraphs, a description of the few existing biosensors for CTXs is provided.

Addressing Ciguatera Risk Using Biosensors for the Detection. . .

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Detection of Ciguatoxins in Fish Samples

The detection of CTXs in fish samples is a challenging task. Researchers are trying to focus on the development of sensitive, specific, simple, rapid, and cheap methodologies for sample screening and quantification. Biosensors, which additionally can be made portable, may be the solution, which would require analytical instrumentation only as a confirmation technique. Recently, the first biosensor for the detection of CTXs was developed by our group [123], targeting four congeners belonging to the CTX1B and the CTX3C groups. Three different mAbs, two capture ones, which specifically bind to the right wing of the CTX1B and 54-deoxyCTX1B (3G8) and of the CTX3C and 51-hydroxyCTX3C (10C9), and a detector mAb, which either specifically binds or show appropriate cross-reactivity with the left wing (8H4) of all four congeners, were used [109–114] (Fig. 4). The capture mAbs were conjugated to MBs and then exposed to CTX standards (CTX1B or 51-OH-CTX3C) or extracts of fishes naturally contaminated with CTXs, followed by the addition of the detector mAb previously biotinylated. Subsequently, polyHRP-streptavidin was incubated and, finally, the immunocomplexes were placed on the working electrodes of an 8-electrode array. Again, a plastic support with magnets underneath each working electrode was used to block MBs in the right position. Then, TMB liquid substrate was incubated, and the reduction current intensity was measured with amperometry. The LODs obtained were 1.96 pg/mL forCTX1B and 3.59 pg/mL for51-OH-CTX3C. The effects of the fish matrix compounds on the detection of CTX1B were evaluated and recovery values calculated. Additionally, an extract of Variola louti, negative for CTXs, was spiked with CTX1B at the threshold value suggested as safety guidance level by the FDA (0.01 μg/kg), and then screened with the immunosensor, which successfully detect this CTX1B concentration. Finally, fishes naturally contaminated from La Réunion island were tested and the amount of CTXs detected correlated well with the results obtained with MBA and CBA. Even if this strategy detects only four of the many existing CTX congeners, the detection was not affected by the presence of marine toxins other than these four CTXs, thanks to the high specificity of the mAbs. Therefore, with this strategy, samples do not require many purification steps (to remove other marine toxins), shortening the assay time compared to other techniques, such as LC-MS/MS. This immunosensor strategy has been simplified even more in one of the last works of our group [124]. In this work, capture mAbs were immobilized directly on carbon electrodes modified with multiwalled carbon nanotubes instead that on MBs. The sandwich assay was then performed and amperometric signals were measured with a ready-to-go smartphone potentiostat. The achieved LOD was 0.001 μg/kg of CTX1B, 10 times lower than the FDA suggested threshold. In addition, recovery values around 100% were obtained, indicating that the fish flesh matrix did not interfere with the performance of the assay, which represents a step forward in comparison with the previous work. However, it is fair to mention that this was probably due to the extraction protocol used in this work, rather than the biosensor configuration, which resulted in very clean samples but involved many purification

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Fig. 4 Schematic representation of the biosensor developed for the detection of the four target CTXs congeners in fish and microalgae samples [123, 125]. Epiphytic Gambierdiscus species (a) were grazed by herbivorous fishes which were then preyed by carnivorous fishes, sampled and CTXs extracted (b1) or isolated, cultured, and CTXs extracted with a fast toxin extraction technique (b2). Fish extracts were exposed to both capture mAbs together (3G8 and 10C9) (c1). Microalgae extracts were incubated with the capture mAbs separately (c2, c3) and together (c4). The biotinylated detector mAb (8H4) and polyHRP-streptavidin were added. Electrochemical detection was achieved by immobilizing the immunocomplexes on the working electrodes of an 8-electrode array, adding TMB substrate, and measuring the reduction current with amperometry

steps. Therefore, in front of a fish suspected for CTXs a compromise would have to be made: a long extraction protocol that gives a more purified extract but that may involve CTXs losses, or a short extraction protocol that may cause undesirable matrix effects. Probably, to take the best decision, also the facilities at disposal for the assay execution would have to be considered. In fact, the long extraction procedure would require more sophisticated instruments and reagents that are not always available. However, in this work, a long extraction protocol was applied to fish samples from Japan and Fiji and the extracts were analyzed with the biosensor,

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sandwich ELISA, CBA, and LC-MS/MS, obtaining comparable results. It is important to explain that the correlation between CBA and the immunosensor was excellent, and the quantifications provided by this last strategy were only slightly lower than the ones by CBA, in contrast to what we had observed in the previous work [123]. This cannot be due to the quality of the extracts, the same for both assays, and probably the geographic origins of the fishes analyzed in the different works may play a role. In fact, in this last work, the fishes were from the Pacific Ocean, and so most likely contained P-CTXs, which are the specific target of the mAbs used. Anyhow, the strategies proposed by our group are sensitive, rapid, easy to perform, and reliable, and they definitely represent a step forward towards the development and applicability of portable devices for the in situ detection of CTXs, especially the last one, in which the instruments required for the analysis are a compact potentiostat and a smartphone.

6.2.2

Detection of Ciguatoxins in Microalgae Samples

After the success in detecting CTXs in fish samples, our group decided to extend the strategy to the detection of CTXs in microalgae samples. Therefore, several strains of Gambierdiscus and Fukuyoa were selected to investigate the differences in CTX production among species [125]. With this purpose in mind, 20.000 cells from 9 Gambierdiscus and 4 Fukuyoa strains were cultured, extracted, and analyzed with the strategy previously developed [123]. Nevertheless, since the capture mAbs (3G8 and 10C9) targeted two different groups of CTX congeners, they were used together, but also separately (Fig. 4). Our results showed the presence of CTXs in 11 out of the 13 strains analyzed. Higher CTX contents were detected when the two capture mAbs were combined, in comparison with the results obtained with just one. A predominance of CTX1B equiv. was observed in 4 out of 6 G. excentricus strains (0.06 to 0.21 fg/cell), and 1 out of 4 F. paulensis strains (0.33 fg/cell). On the other hand, G. australes and the other 2 G. excentricus strains showed a higher abundance of CTX3C equiv. (0.16 fg/cell and 0.04–3.54 fg/cell, respectively). The unique strain of G. caribaeus tested showed an equal amount of both types of congeners (0.13 fg/cell). Additionally, the same microalgae extracts were screened with CBA, which identified CTX-like activity only in 4 out of the 6 G. excentricus strains. It should be kept in mind that the strategy, although detects only four CTX congeners, is not affected by the presence of the other toxic compounds produced by Gambierdiscus and Fukuyoa, such as maitotoxins, and so provides reliable results. In fact, maitotoxins are known to affect in the execution of the CBA, if no pretreatment of the extract is performed. It is also important to mention that, in this study, a new fast CTXs extraction technique was developed, allowing to operate with as low as 20.000 cells of Gambierdiscus, an amount that can be easily found in natural environments, making it suitable for the screening of field samples. This strategy was also used to characterize a G. belizeanus strain first reported in the Canary Islands, detecting the production of congeners of both the CTX1B and CTX3C groups [126].

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To make further advances in the direction of in situ detection of CTXs, this strategy was also applied to the analysis of field samples [122]. In this work, macroalgae were collected in Majorca, shaken for microalgae cell detachment, extracted, and then exposed to both capture mAbs simultaneously, to maximize the probabilities to detect CTXs. Results showed CTX contents in one sample (13.35 ± 0.5 pg CTX1B equiv./cell) and traces of CTX (below LOQ) in three out of nine analyzed samples, demonstrating that the system is suitable for the screening of field samples.

7 Conclusions CFP is one of the most common seafood-borne diseases, whose real incidence is difficult to estimate. Many phenomena can co-occur during a CFP event, making its prediction very difficult. In order to get a correct CFP risk assessment and management in a specific area, the detection of Gambierdiscus/Fukuyoa species and the discrimination between toxic and non-toxic fish specimens are crucial. Light microscopy has been commonly used to identify microalgae cells and estimate their abundances, but it is time consuming, requires trained personnel, and usually does not allow identification at the species level. Regarding CTXs, several methodologies have been developed during the years to determine the toxin content in an extract either from microalgae or fish, but they involve expensive and time-consuming procedures. Additionally, these methodologies require the use of standards that, in the case of CTXs, are scarce and expensive. In this picture, there is an extreme need for rapid and reliable bioanalytical devices able to detect the DNA of CTX-producing microalgae species and CTXs in fish and microalgae samples. Biosensors could be the answer to this need. They are sensitive, specific, rapid, robust, cost-effective, and do not need highly trained personnel to be operated. The availability of biosensors that detect CTXs in fish samples at a level even lower than the suggested guideline is of outmost interest. They can be helpful to rapidly screen and discriminate contaminated specimens, and consequently to take quick and right decisions. Moreover, the detection of CTXs in microalgae pellets with low cell concentrations from Gambierdiscus and Fukuyoa cultures has been proved to provide useful information about the toxin-producing behavior of these species without requiring large-scale cultures. When the ability to detect CTXs from few microalgae cells is exploited in the analysis of field samples, the utility of the biosensors is even more evident. In fact, the application of these techniques to the analysis of this kind of samples could help to identify areas of interest for surveillance of marine specimens. Additionally, the instrumentation needed for the analysis can be easily miniaturized at a relatively low cost, and therefore these biosensors can be easily integrated into portable devices. The biosensors able to detect CTX-producing microalgae species in field samples at low cell concentrations is also very useful to know their geographical distribution and consequently where fishes can accumulate CTXs. This information could contribute to design the

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sampling strategy more rationally and therefore improve CFP risk assessment and management. In conclusion, the results obtained so far with the biosensors are promising and underline the high performance of these bioanalytical tools together with their successful applicability to the analysis of natural samples. The biosensors for the detection of CTXs can be considered as a useful screening method, although also able to provide toxin quantifications, complementary to instrumental analysis, which would be required for confirmation purposes. The biosensors for the detection of Gambierdiscus and Fukuyoa can be complementary to light microscopy, identifying at the species level. Up to date, such devices are not yet routinely used, as further validation studies are needed, but sure, the integration of biosensors in the monitoring of fish and environmental samples will certainly contribute to better assess CFP risk and prevent outbreaks. Acknowledgments This work was supported by the Agencia Estatal de Investigación (AEI) and the Fondo Europeo de Desarrollo Regional (FEDER) through the CIGUASENSING (BIO201787946-C2-2-R) and the CELLECTRA (PID2020-112976RB-C21) projects.

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Antibody, Aptamer and Affimer-Based Affinity Tools for Marine Toxin Biosensing Caroline Murphy

Contents 1 Introduction: Overview of Affinity Tools and Marine Toxin Detection . . . . . . . . . . . . . . . . . . . . 2 Harmful Marine Toxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 The Use of Affimer, Aptamer and Antibody-Based Systems to Detect Marine Toxins . . . . 4 Discussion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

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Abstract Antibody-based detection systems have long been chosen to facilitate on-site monitoring of environmental markers such as marine toxins. Antibodies were the first detection molecules to be seen as effective replacements to mouse bioassays, high-performance liquid chromatography (HPLC) and mass-spectrometry (MS)based systems in the detection of harmful marine toxins. Thereby satisfying European Union regulatory standard 853/2004, legislation that states that all shellfish produced must be routinely monitored and tested for the presence of regulated marine toxins before they can reach the market. Antibodies have progressed significantly in their capabilities over the past 70 years, with much investigative research being carried out with regard to their production, purification, sensitivity enhancement and their incorporation into sensor platforms. However, reports of new smallmolecule detectors such as affimers and aptamers over the past 20 years may now herald a ‘changing of the guard’. Many reports of aptamers being used for the detection of marine biotoxins have been produced and suggest that their sensitivities equal or surpass that of antibodies. The next stage for the use of aptamers lies in assessing and enhancing their incorporation into sensor platforms as well as investigating their capacity to detect multiple forms of marine congeners, of which there are many.

C. Murphy (✉) School of Biotechnology, Dublin City University, Dublin, Ireland e-mail: [email protected] Fiona Regan, Peter-Diedrich Hansen, and Damià Barceló (eds.), Biosensors for the Marine Environment: Present and Future Challenges, Hdb Env Chem (2023) 122: 47–62, DOI 10.1007/698_2022_953, © The Author(s), under exclusive license to Springer Nature Switzerland AG 2023, Published online: 29 January 2023

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Graphical Abstract

Keywords Affimer, Antibody, Aptamer, Environmental monitoring, Marine toxin, Toxin congener

1 Introduction: Overview of Affinity Tools and Marine Toxin Detection Marine bacteria were the first forms of life to exist and thrive on Earth for almost two billion years. Their presence in the world’s oceans is not a new phenomena, however, Earth’s overpopulation is! Increased pressure on coastal regions to provide clean food whilst being a dumping ground for waste is a significant paradox. An algal bloom is a sizable growth of microscopic algae or algae-like marine bacteria, blooms can occur in salt, brackish and fresh water bodies. Some algal blooms produce toxins, these are known as harmful algal blooms or HABs. Algal blooms are naturally occurring, however, contributing factors to more intense and extensive algal blooms include intensification of agriculture resulting in run-off and waste products entering water courses, industrial waste, climate change, marine transportation, an ever-increasing population requiring more food and many more. While it is still not fully understood, the reason the algal blooms produce these harmful toxins,

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the prevalence of poisoning due to seafood is being increasingly felt due to a rising human population requiring more food. Concomitantly, seafood originating from algal-rich water is being consumed without knowing or without fully understanding its toxification status. Bivalve molluscs (such as mussels, scallops, clams, snails and oysters, among others) must be considered as they feed by filtering water and bio-accumulate harmful particulates in their tissues such as algal cells (which may contain toxins), viruses, bacteria, parasites and industrial wastes. In addition, bivalve molluscs are commonly eaten raw, thereby contributing to a higher consumer risk. Of particular concern is the ingestion of marine biotoxins due to their acute and chronic effects (Table 1). Marine biotoxins are classified according to their chemical properties and the symptoms they elicit when consumed. Marine biotoxins commonly found in Europe are legislated for, with established regulatory limits agreed, however, ‘newly emerging biotoxins’ are not fully legislated for, but are becoming more prevalent, and thus require attention. Additional tools are required to reduce seafood poisonings and mitigate economic losses in the seafood industry. Routine toxicity measurements are carried out using mass spectrophotometric (MS) analysis, this requires transporting samples to laboratories, thus, increasing cost and time [1, 2]. By measuring marine toxins with tools that facilitate on-site measurement, more immediate action can be taken to prevent seafood poisonings and mitigate against economic losses to fishing industries. While the use of antibodies in effective sensing technology has dominated, in the past 20 years, the use of aptamer-based sensor technology is contributing to the advancement of marine biotoxin monitoring capabilities with affimer-based technology currently warranting major interest.

2 Harmful Marine Toxins Hydrophilic marine toxins include paralytic shellfish poisoning (PSP) toxins and amnesic shellfish poisoning (ASP) toxins. They are commonly produced by Pseudonitzschia and Alexandrium marine phytoplankton species, respectively. As they are water soluble, they are removed more readily and do not persist as long as lipophilic toxins within the tissues of shellfish [1]. Saxitoxin (STX) and up to 30 of its analogues are potent neurotoxins and members of the PSP group of toxins, they block voltage-gated sodium channels, interrupting nerve signal transmission paralysing muscles and causing paralytic shellfish poisoning [24]. Symptoms of PSP range from a tingling sensation on the lips/mouth to nausea, vomiting, diarrhoea to muscular paralysis and respiratory difficulties which can be fatal [25]. The EU regulatory limit for the consumption of STX is 800 μg/kg [26, 27]. PSP toxin was identified in Castlemaine in the South West Coast of Ireland in 2021, a number of official bodies including the Irish Food Safety Authority of Ireland (FSAI), the Health Service Executive and the Marine Institute warned the public not to collect shellfish from this area. Local businesses were advised of the toxic occurrence and appropriate action was taken, monitoring the presence of the PSPs until they subsided [28] (FSAI, 13.July.2021).

Yes, monoclonal antibody to BTX1 (LOD 14 ng/mL), BTX2 LOD 0.106 μg/L

Dinophysis, Prorocentrum

Karenia brevis, Chattonella

Yes, polyclonal antibody (LOD 75 μg/kg)

Prorocentrum reticulatum

Okadaic acid (OA) (diarrhetic shellfish poisoning (DSP)) Emerging toxins Brevetoxin (BTX)

Yes. (available commercially)

Yes, competitive ELISA, quantitative range 0.45–8.46 ng/mL, LOQ 57 μg/kg Yes, linear range of ELISA to detect OA: 20–750 pg/mL, and the limit of detection (LOD) was 12 pg/mL

Yes. (LOD 0.03 ng/mL) (available commercially)

Dinoflagellates (Alexandrium tamarense, Gymnodinium catenatum) Pseudo-nitzschia

Azadinium spinosum

Antibody detection

Producer species

Azaspiracid (AZA)

Domoic acid (DA) (amnesic shellfish poisoning) Lipophilic Yessotoxin (YTX)

Harmful marine toxin Hydrophilic Saxitoxin (STX) (paralytic shellfish poisoning)

Table 1 Detection of harmful marine toxins by antibodies, aptamers and affimers

No

No

No

Yes. LOD 70 pg/mL

Yes. LOD of 0.106 μg/L detection range 0.01–2,000 μg/L.

No

No

No

No

None identified.

Yes. (LOD 0.45 ng/mL)

Yes. 30 min detection. AuNPs. LOD 10 fM (3 fg/ mL), range 10 fM to 0.1 μM.

Aptamer detection

Affimer detection

No EU legislation

3.75 mg YTX eq./ kg 160 μg AZA eq./ kg 160 μg OA eq./kg

20 mg/kg

800 μg/kg (STX eq.)

EU Regulatory limit

[18–20]

[13–17]

[11, 12]

[10]

[6] [7–9]

[3–5]

References

50 C. Murphy

Yes, immunosensor available

Yes, polyclonal antibody.

Gambierdiscus sp.

Pseudomonas spp. in puffer fish. Vibrio sp. Alteromonas sp.

eq. equivalent, b.w. body weight, LOD limit of detection, ARfD acute reference dose

Ciguatoxin (CTX) (ciguatera fish poisoning (CFP)) Tetrodotoxin (TTX) Yes. (LOD 1.21 ng/mL)

No

No

No

[21, 22]

[9, 23]

No EU legislation (ARfD – 0.25 μg/kg)

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Domoic acid (DA) causes amnesic shellfish poisoning (ASP), symptoms include abdominal cramping, vomiting, diarrhoea and more severe cases may cause neurological symptoms such as disorientation, seizures, memory loss, and some cases have been fatal [1, 25, 29]. The EU regulatory limit (Regulation (EC) No. 853/2004) for the consumption of DA is 20 μg/kg [29]. Lipophilic toxins are ‘fat-friendly’ compounds, which can dissolve in the body’s lipids, oils and fats and for this reason, they are more persistent than hydrophilic toxins in tissues, consequently causing more damage. Lipophilic toxins include yessotoxins (YTX), azaspiracids (AZAs) and okadaic acid (OA). Their consumption mainly causes gastrointestinal disorders such as nausea, diarrhoea and vomiting [25]. Yessotoxins are polycyclic ether compounds, they are distributed globally and are produced by phytoplankton dinoflagellate Prorocentrum reticulatum [30]. YTX accumulates in filter feeding shellfish and when ingested by mammals primarily targets cardiac muscle, but can also affect the liver and pancreas [30]. The EU has imposed a YTX regulatory limit in shellfish of 3.75 μg/g expressed as YTX equivalents (Regulation (EC) No 853/2004) [1, 31]. Azaspiracid poisoning (AP) is caused by the consumption of shellfish contaminated with azaspiracid (AZA) or one of its 30 currently known analogues. AZA is produced by dinoflagellate Azadinium spinosum [32]. The EU has imposed a regulatory limit of 0.16 μg/g AZA equivalents in bivalve molluscs (Regulation (EC) No 853/2004) [1, 33]. Diarrhetic shellfish poisoning (DSP) is caused by okadaic acid (OA), dinophysistoxins (DTX) and pectenotoxins (PTX). They are predominantly produced by Dinophysis and Prorocentrum lima. OA causes diarrhoea by stimulating intestinal cells to release sodium, leading to an accumulation of gastrointestinal fluid and abdominal cramping [34, 35]. The EU regulatory limit for OA is 0.16 μg/g expressed as OA equivalents (Regulatory (EC) No 853/2004) [1]. While the acute effects of OA poisoning are known, the chronic effects of long-term exposure to OA are not fully understood, chronic exposure to OA toxins is pointing towards an increased risk of cancer due to its tumour promoting properties [36, 37]. Emerging toxins that are not routinely detected in a specific area must be considered due to the risk of the spread of their causative organisms due to climate change and the movement of these causative organisms from their native environment to a new environment caused by the movement of ships. Brevetoxin (BTX) is a cyclic, polyether lipophilic marine neurotoxin which causes neurotoxic shellfish poisoning (NSP), BTX is produced by ‘red-tide’ causing dinoflagellates from genera Karenia and Chattonella commonly found in the Gulf of Mexico and the Caribbean. A related species Karenia mikimotoi, a dinoflagellate phytoplankton is known to cause extensive blooms off the Irish coast [38]. The BTX neurotoxin is released from the dying cells and if consumed or inhaled (through aerosolized toxin) by mammals or birds binds to and blocks voltage-gated sodium channels in muscle and nerve cells, impairing their function and causing neurotoxic shellfish poisoning, which can be fatal [18]. Tetrodotoxin (TTX) is one of the most dangerous toxins in the marine environment. TTX targets sodium channels, blocking nerves and muscles. Its presence has been identified in the Tetraodontidae family which includes the puffer fish. The

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puffer fish is eaten as a delicacy in Japan known as ‘fugu’. Members of the Tetraodontidae family are most commonly found in south-east Asia, but in recent years, some species have been identified in the Mediterranean and in the English Channel [39, 40]. Ciguatera toxins (CTXs) are produced by the dinoflagellate Gambierdiscus toxicus. CTXs cause gastrointestinal problems such as vomiting and diarrhoea, and neurological problems such as tingling [22]. CTX poisoning is becoming more prevalent, with Friedman et al. reporting that ciguatera fish poisoning (CFP) is the most commonly reported seafood-associated illness [21].

3 The Use of Affimer, Aptamer and Antibody-Based Systems to Detect Marine Toxins Affimers are engineered protein scaffolds and have been highlighted as the next alternative to antibodies [41]. Affimer technology is provided by a company called Avacta® Life Sciences [42]. Affimers are selected from phage display libraries of scaffold proteins. They are developed from two parent scaffolds: type I is based on the human protease inhibitor stefin A and type II is based on a phyto-cystatin protein (Adhiron scaffold) [41]. Each scaffold is composed of four beta-sheets and an alphahelix, they contain a short random sequence composed of 9 amino acids within 2 variable loops, and these loops facilitate interactions with potential binders of interest. In 2018, S. Kyle identified that affimers are advantageous over antibodies such that they are simpler and easier to develop, can be generated more rapidly, and possess superior stability in a broad pH range. Affimers can provide comparable affinities to antibodies, but have higher reported specificities and can be identified without the use of animals [41]. Affimers have demonstrated excellent potential as future therapeutics, as they can act as excellent cell signalling blockers [43] and in vivo imaging tools. There are currently no reported affimers towards marine biotoxins, which may merit investigation in the future. Aptamers are single-stranded nucleic acid (deoxyribonucleic acid (DNA) and ribonucleic acid (RNA)-based molecules (approximately 40–100 nucleotides long)) first introduced in the 1990s [13], each oligonucleotide is composed of a central region composed of random nucleotides (approximately 20–28 nucleotides in length), these are flanked by two constant sequences (15–25 nucleotides in length), which facilitate primer annealing during polymerase chain reaction (PCR) [44, 45]. The word ‘Aptamer’ comes from the Latin words ‘aptus’ and ‘meros’ meaning ‘to fit’ and ‘particle’, respectively [44]. Aptamers can form many different three-dimensional conformational structures such as hairpin, stem ring, convex ring, false knot and many more, the formation of these three-dimensional structures forms the basis of the ‘lock-and-key’ interaction between the aptamer and the target molecule [13]. Aptamers are specifically developed to detect small molecules, therefore, they may serve as highly suitable tailor-made detection agents for marine biotoxins. They

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have the capacity to be engineered post-production and have been heralded as a viable replacement to antibodies. The use of aptamers for environmental monitoring is challenging due to a number of hurdles 1: poor sensitivities achieved in targeting small molecules and 2: difficulty in developing sensing platforms [46]. DNA aptamers may be more suitable for environmental monitoring than RNA-based aptamers, as the DNA aptamers are more stable. Aptamers are selected according to their specificity towards target molecules from a large random sequence pool (1014 to 1015 random sequences) using ‘Systematic Evolution of Ligands by Exponential Enrichment’ or ‘SELEX’. A DNA or RNA library is challenged by the target molecule of interest (for example, protein, virus, peptides, antibiotics, nucleic acids, toxins, small molecules and chemical compounds [44]), separating steps remove the weakly bound and unbound random sequences, from the tightly bound sequences, the eluted DNA sequences are amplified using polymerase chain reaction (PCR) and the eluted RNA sequences are amplified by reverse transcription PCR [47]. The resulting selected binding nucleic acid sequence of interest is called an ‘aptamer’ and exhibits high sensitivity towards its target. The reported advantages of aptamers over monoclonal antibodies include smaller size, prolonged shelf life, lower batch-to-batch variation, thermal stability, lower immunogenicity and toxicity, lower development costs and more convenient conjugation and modification [44]. In 2020, Qiang et al. reported the development of a STX specific aptamer [3]. The colourimetric sensor was based on the aggregation of gold nanoparticles (AuNPs). The STX specific aptamer was functionalised onto AuNPs, and in the presence of STX the aptamer binds and aggregation occurs, thus inducing a colour change, and producing a result in a rapid time of 30 min. This is comparable and in direct competition with antibody-based immunosensor developed against STX of 36 min [4]. Li et al. subsequently developed a ‘terminal fixed’ aptamer, which involved taking the original aptamer [3] against STX and extending the two ends of the aptamer to include a spacer followed by bases that upon interaction with its complementary partner form a double helix [5]. The upgraded aptamer displayed 145-fold higher affinity for STX compared to the parent molecule [5]. This work highlighted the post-modifications that are possible with aptamers. Qi et al. reported the development of a label-free electrochemical system for the successful detection of STX using aptamers [48]. The DNA-based aptamer-triplex against STX was immobilised on the surface of the screen-printed electrode using a nano-tetrahedron structure [48]. Upon detection of STX in seawater samples, the aptamer bound to the STX, changing the structure of the aptamer, which was subsequently released, thus changing the electrochemical signal. The system showed a good limit of detection (LOD) with high repeatability in seawater samples [48]. A significant volume of work was carried out on the development of aptasensors towards okadaic acid (OA). OA is a low molecular weight compound that causes diarrhetic shellfish poisoning (DSP), the regulatory limit for OA is 0.16 μg/g (199.75 nM) (EC regulation No 853/2004) [1]. In 2016, Gu et al. developed an aptamer to detect OA using SELEX assisted by graphene oxide (GO) [14]. The single-stranded DNA (ssDNA) aptamer was incorporated into a competitive

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enzyme-linked aptamer assay (ELAA) and exhibited an LOD