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Laboratoire Interdisciplinaire des Environnements Continentaux LIEC – UMR 7360 CNRS, Metz

THESE

Pour l’obtention du titre de:

DOCTEUR DE L’UNIVERSITE DE LORRAINE

Mention: Ecotoxicologie, Biodiversité, Ecosystèmes

Kahina MEHENNAOUI

Understanding the impact of engineered nanoparticles

Gammarus sp. as a valuable non-vertebrate model?

Soutenue publiquement Le 20 décembre 2017 devant la commission d’examen :

Laure Giamberini Université de Lorraine – UMR 7360 CNRS - LIEC

Arno Gutleb Luxembourg Institute of Science and Technology

Catherine Mouneyrac Université Catholique de l’Ouest – MMs - Angers

Mélanie Auffan CNRS – CEREGE – Aix en Provence

Erik Ropstad Norwegian University of Life Sciences

Elise David Université Reims Champagne-Ardenne – UMR-I 02 SEBIO

François Guérold Université de Lorraine – UMR 7360 CNRS - LIEC

Sebastien Cambier Luxembourg Institute of Science and Technology

Ecole Doctorale Sciences et Ingénierie Ressources Procèdes Produits Environnement,

RP2E, ED Nº 410, Nancy

Environmental Research and Innovation (ERIN)

Department,

Luxembourg Institute of Science and Technology,

(LIST), Luxembourg

Directeur de thèse

Directeur de thèse

Rapporteur

Rapporteur

Examinateur

Examinateur

Examinateur

Examinateur

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To my parents

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ABSTRACT The potential toxicity of nanomaterials is of high societal and scientific interest due to the promise of ground-breaking innovations for many technical applications. However, toxicity can often not be related to the actual size, mass or surface area of the single nanoparticles (NPs) or the NP agglomerates. Therefore, it can be proposed that the toxicity is greatly influenced by other inherent and non-understood properties of the particles to which ions dissolving from the particle, surface or molecules adhering to the surface interfering with the uptake of NPs into cells, may have important contributions. The PhD project “NANOGAM”, closely linked up to CORE2012 NANION project that aims to obtain knowledge to understand some of the processes and factors involved in NP uptake and toxicity as such understanding is a prerequisite for the development of nanomaterials following the safer-by-design philosophy. This PhD project aims to investigate, based on known characteristics of the key physico-chemical parameters; as size and surface functionalities, of a well-chosen list of silver and gold NPs, the uptake, and dependent biological effects of different complexity (mortality, behavioural effects, physiological effects, transcriptomic effects, etc.), on a sensitive species; Gammarus fossarum (Crustacea Amphipoda), in order to understand to which extent toxicity of nanomaterials is due to intrinsic material properties or ion leaching. Such understanding will contribute to the prediction of toxicity based on material properties rather than repetitive testing of an indefinite number of new nanomaterials. G. fossarum were exposed at low concentrations of AgNPs and AuNPs for 72h or 15 days in presence or absence of food. The obtained results showed that (i) surface coating is the main factor governing AgNPs and AuNPs uptake by G. fossarum, (ii) both released ions and NPs themselves play a role in the potency of the studied AgNPs and AuNPs and (iii) chemical composition led to different effects at the sub-individual levels (target genes expression) and different tissue distribution as AgNPs were found in G. fossarum gills while AuNPs were found in the intestinal caeca. Additionally, this work shows that Gammarus sp. are valuable models for the study of the effects of AgNPs and AuNPs.

Keywords: Gammarus sp.; silver nanoparticles; gold nanoparticles; ions release; multi-biomarker approach;

transcriptomic

RESUME La toxicité potentielle des nanomatériaux présente un intérêt sociétal et scientifique élevé en raison de la promesse d'innovations pour de nombreuses applications techniques. Cependant, elle n’est pas forcément liée à la taille réelle, à la masse, à la surface des nanoparticules (NP) ou à leurs agglomérats. La toxicité des NPs pourrait être fortement influencée par d'autres propriétés inhérentes et encore incomprises telles que le relargage d’ions, de la particule elle-même, sa surface, ou des molécules adhérentes à la surface, qui interfèreraient avec l'absorption cellulaires des NPs. Le projet « NANOGAM» étroitement lié au projet « FNR CORE2012 NANION », vise à définir certains processus et facteurs impliqués dans l'absorption des NPs et leur toxicité. Une telle compréhension est une condition préalable au développement des nanomatériaux, fondement de la philosophie « safer-by-design ». Les objectifs de ce projet de thèse sont multiples. En tenant compte des caractéristiques des principaux paramètres physico-chimiques tels que la taille et l’aspect de la surface, l’étude a porté sur l'absorption de NPs d'argent et d'or, et leurs effets biologiques via une approche multi-biomarqueurs (mortalité, effets comportementaux, effets physiologiques, effets transcriptomiques, etc.) sur une espèce sensible, Gammarus

fossarum (Crustacea Amphipoda). Le but de cette investigation est de comprendre si la toxicité des nanomatériaux est inhérente aux propriétés intrinsèques des NPs ou plutôt aux ions relargués, ce qui contribuera à la prédiction de la toxicité des NPs en rapport avec leurs propriétés physico-chimiques et ce afin de limiter le nombre d’essais répétitifs sur de nouveaux nanomatériaux. G. fossarum ont été exposés à de faibles concentrations d'AgNPs et AuNPs pendant 72h à jeun et 15 jours nourris. Les résultats obtenus ont montré que (i) la nature de l’enrobage de surface est le principal facteur responsable de l'absorption d'AgNPs et d'AuNPs par G. fossarum ; (ii) les ions libérés et les NPs elles-mêmes jouent un rôle dans la toxicité des AgNPs et AuNPs étudiées ; (iii) la composition chimique des NPs a conduit à des effets différents aux niveaux sub-individuels (transcriptomique), ainsi qu’à une distribution différente dans les tissues selon la nature métallique de la NP. Les AgNPs ont été localisées dans les branchies de G. fossarum tandis que les AuNPs ont été observées dans les caeca intestinaux. Cette étude a également révélé que Gammarus sp. est un excellent modèle pour l'étude de la toxicité et des effets des AgNPs et des AuNPs.

Mots-clé : Gammarus sp. Nanoparticules d’argent, nanoparticules d’or, relargage d’ions, approche multi-biomarquers, transcriptomiques

AVANT PROPOS/ FOREWORD

The research performed within the project” NanoGAM” have been funded by the National Fund for Research

(FNR Luxembourg- AFR-PhD-9229040). This project was closely linked up to CORE2012 NANION and FP7

FUTURENANONEEDS (FNN). These two projects allowed the funding of ICP-MS and NanoSIMS analyses.

Additionally, NanoGAM project were closely linked to PhD projects at Université de Lorraine (ANR P2N

MESONNET and ANR nanoSALT) and PhD programs at Oslo University (NanoZebra).

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ACKNOWLEDGEMENTS

These last four years were such a rewarding and meaningful experience. I would never have succeeded in

this work without the support of colleagues, relatives and friends.

First of all, I would like to thank Arno Gutleb and Pr. Laure Giamberini. Thank you for giving me the

opportunity to make this PhD and research possible. Your advices, ideas and inputs helped me throughout

this process. You also taught me how to face the difficulties encountered during this work. Thank you, Laure,

for your trust and for accepting to be my supervisor. Thank you for your availability, your precious advices

and the good moments shared during the different conferences over Europe. Thank you, Arno, for giving me

the opportunity to make a “PRE-doc” in the TOX group that resulted in this present PhD project. Thanks a lot

for your availability, trust and optimism. Thank you also for keeping my ideas structured and realistic.

Working with you was such an amazing experience.

I also would like to thank Pr. François Guérold and Sebastien Cambier for accepting to be my co-supervisors.

Thank you, François, for your advices and our discussions about the Gammarus. Sebastien, thank you for

your availability, for teaching me molecular biology, for your help in the lab even on Sundays, for criticizing

my work, and helping me finding solutions. Thanks also for all the discussions we had during these last 4

years.

I would like to address my sincere acknowledgments to the opponents who accepted to evaluate my present

work: Catherine Mouneyrac, Mélanie Auffan, Elise David and Erik Ropstad. I am also deeply grateful to the

Luxembourg Institute of Science and Technology, Lucien Hoffmann, the Fond National de la Recherche

(Luxembourg) and the Laboratoire Interdisciplinaire des Environnements Continentaux at Lorraine University

for making this research possible.

This work would have been impossible without the help of colleagues. Tommaso, thank you for correcting

my works, for your help and support, especially during these last very stressful months. I learned a lot with

you. I am also grateful to Natasa, who supervised my work during the internship and who kept helping me

from NIVA Oslo where she is now. Working with you was a great experience. I will never be thankful enough

to Aline, Boris and Sylvain. Thank you all for the support. Sylvain thank you for your precious help and

advices, for all the discussions we had on the way back to Metz. Thanks also to Aurélie for her support during

difficult moments. Thanks, the both of you for your presence and support. Aline you’ve been a precious

colleague and friend. I’ll be always grateful to you. You’ve always been here and it was so nice to work with

you. Thank you very much for your support. I am also grateful to Boris for his good spirit, optimism, and

enthusiasm. You’ve always made the days funnier in the lab… there’re so many things to say that I do not

even know from where to start… Thank you for being here. I have been very lucky to share the office with

you girls; Anouk, Joanna and Blandine. I wish you all the best for your future careers. I also thank all my PhD

fellows: Marc, Marie B, Sebastien L, Rodolphe and Benoit, I wish you the best and I’m sure you will succeed.

I am also indebted to all the LISRA group and the nice moments shared at SETAC. Thank you, Enrico, for your

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availability. I am also grateful to all the ERIN and MRT colleagues: Gea, Xavier, Magda, Christelle, Elisa,

Vincent R, and Brahim.

This work would not be possible without the help of all the ERIN support. Thank you Servane, Marie F,

Delphine and Lionel for helping me in the lab and in the field for Gammarus sampling. Servane, it was so

nice to work with you. Thank you also Johanna, Sebastien P, Audrey L, Audrey J, Aude, Laurent, Francois,

Marine, Celine, Cecile, Cyril, Jeff, Cedric and Jenny. Thank you, Alain, for all your help in the beginning of my

PhD, for helping me finding the nice “spot” for the Gammarus sampling. I am also greatly thankful to Anaïs

C for your support during difficult moments and availability and for helping me in the lab. Thank you also

Jean-Nicolas, Nathalie, Esther, Gaëlle, Jean-Sebastien and Patrick for your help and fruitful collaboration. I

learned a lot with you concerning new techniques. I also thank Maryline and Vincent for your trust and for

the collaboration.

I also would like to thank my colleagues from LIEC; Maël Garaud, Jennifer Andreï, Vanessa Koehle-Divo and

Alice Gossiaux for your help during the lab work. Thank you, Carole, Sandrine, Vincent and Simon, for your

advices these last years.

I would like to dedicate this work to the late Pr. Stephen J Klaine who inspired me for many years. I am so

grateful to have known you.

I am also deeply grateful to my friends in France and Algeria. Thank you all for being here. I thank also Hanne

for all the great moments and very interesting discussions we had. I also thank Greg for your support, your

help your advices and kindness. Thank you for keeping me in the right behaviour to achieve my objectives.

Last but not least, I would like to thank my family; my parents, my brother and his wonderful wife; Mehdi

and Fairouz for your great support and all the good moments spent with you each time I came back home.

My parents, Smaïl and Fatima, thank you for allowing me to achieve this work. Thank you for giving me this

opportunity despite the very hard moments 7 years ago when I left home. I am deeply grateful to you, you

always supported me, helped me in the difficult moments. I learned so much with you and I hope that I make

you proud. I love you.

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THESIS ACHIEVEMENTS

Articles in the thesis frame

Gançalo Vale, Kahina Mehennaoui, Sébastien Cambier, Giovanni Libralato, Stéphane Jomini, Rute F. Domingos. Manufactured nanoparticles in the aquatic environment – biochemical responses on freshwater organisms: a critical overview. 2016. Aquatic toxicology 170, 162-174.

Kahina Mehennaoui, Anastasia Georgantzopoulou, Vincent Felten, Jennifer Andreï, Maël Garaud, Sébastien Cambier, Tommaso Serchi, Sandrine Pain-Devin, François Guérold, Jean-Nicolas Audinot, Laure Giamberini, Arno C. Gutleb. 2016. Gammarus fossarum (Crustacea, Amphipoda) as a model organism to study the effects of silver nanoparticles. Science of the Total Environment 566, 1649-1659.

Kahina Mehennaoui, Sébastien Cambier, Sylvain Legay, Tommaso Serchi, François Guérold, Laure Giamberini, Arno C. Gutleb. Identification of reference genes for RT-qPCR data normalization in Gammarus fossarum (Crustacea Amphipoda). Submitted to Scientific reports

Kahina Mehennaoui, Sébastien Cambier, Tommaso Serchi, François Guérold, Johanna Ziebel, Jean-Sebastien Thomann, Nathalie Valle, Laure Giambérini, Arno C. Gutleb. Influence of size and surface coating on silver and gold nanoparticles uptake and their molecular effects on Gammarus fossarum (Crustacea Amphipoda). Submitted to Science of the Total Environment

Kahina Mehennaoui, Sébastien Cambier, Tommaso Serchi, François Guérold, Laure Giambérini, Arno C. Gutleb. Sub-chronic effects of silver and gold nanoparticles on Gammarus fossarum (Crustacea Amphipoda): from molecular to behavioural responses. In prep.

Book chapter

Arno C. Gutleb, Sébastien Cambier, Teresa Fernandes, Anastasia Georganztopoulou, Thomas A.J. Kuhlbusch, Iseult Lynch, Ailbhe Macken, Kahina Mehennaoui, Ruth Moeller, Carmen Nickel, W. Peijnenburg, Tommaso Serchi. 2016. Chapter 4: Environmental Fate and Effects of Nanomaterials in aquatic Freshwater Environments. 96-114. Nanomaterials: A guide to fabrication and applications. Edited by Sivashankar

Krishnamoorthy. CRC Press, Taylor & Francis Group.

Articles in the nanotoxicology topic

Jennifer Andreï, Sandrine Pain-Devin, Vincent Felten, Simon Devin, Laure Gaimberini, Kahina Mehennaoui, Sébastien Cambier, Arno C. Gutleb, François Guérold. 2016. Silver nanoparticles impact the functional role of Gammarus roeseli (Crustacea Amphipoda). Environmental Pollution 208, part B, 608-618.

Articles out of the topic

Vincent Rogé, Anastasia Georgantzopoulou, Kahina Mehennaoui, Ioana Fechete, François Garin, Aziz Dinia, Arno C. Gutleb, Damien Lenoble. 2015. Tailoring the optical properties of ZnO nano-lazers and their effect on in vitro biocompatibility. RCS advances 5, 97635-97647.

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Oral presentations

Effets de différentes tailles de nanoaprticles d’argent sur le comportement et la physiologie de Gammarus sp. Mehennaoui K., Georgantzopoulou A., Felten V., Garaud M., Andreï J., Cambier S., Serchi T., Contal S., Balachandran Y.L., Pain-Devin S., Giamberini L., Gutleb A.C. Colloque ARET-SFTG, Paris France, 3-4.06.2014.

Influence of size and surface coating on silver and gold nanoparticles uptake by Gammarus fossarum. Mehennaoui K., Cambier S., Serchi T., Ziebel J., Chauvière A., Lentzen E., Valle N., Thomann J.S., Guérold F., Giamberini L., Gutleb A.C., 32nd international conference on environmental geochemistry and health, Brussels, Belgium. 4-8.07.2016.

Influence of size and surface coating on silver and gold nanoparticles uptake by Gammarus fossarum. Mehennaoui K., Cambier S., Serchi T., Ziebel J., Chauvière A., Lentzen E., Valle N., Thomann J.S., Guérold F., Giamberini L., Gutleb A.C., ES1205 final conference, Aveiro, Portugal. 7-8.02.2017.

Do size and surface coating of AgNPs and AuNPs influence their uptake and molecular effects on Gammarus

fossarum? Mehennaoui K., Cambier S., Serchi T., Ziebel J., Chauvière A., Lentzen E., Valle N., Thomann J.S., Guérold F., Giamberini L., Gutleb A.C., PRIMO19 international conference, Matsuyama, Ehime, Japan. 30.6-3.07.2017.

Posters

Effects of different sizes of silver nanoparticles on physiological and behavioural responses of Gammarus

fossarum. Mehennaoui K., Georgantzopoulou A., Felten V., Garaud M., Andreï J., Cambier S., Serchi T., Contal S., Balachandran Y.L., Pain-Devin S., Giamberini L., Gutleb A.C. Geel, Belgium. 4.12.2014.

Surface and Size-dependent effects of silver nanoparticles on Gammarus fossarum: Link between physiological and behavioural responses. Mehennaoui K., Georgantzopoulou A., Felten V., Garaud M., Andreï J., Cambier S., Serchi T., Contal S., Ziebel J., Guignard C., Balachandran Y.L., Pain-Devin S., Giamberini L., Gutleb A.C. Nanoposter Virtual conference. 04.2015.

Surface and Size-dependent effects of silver nanoparticles on Gammarus fossarum: Link between physiological and behavioural responses. Mehennaoui K., Georgantzopoulou A., Felten V., Garaud M., Andreï J., Cambier S., Serchi T., Contal S., Ziebel J., Guignard C., Balachandran Y.L., Pain-Devin S., Giamberini L., Gutleb A.C. 25th SETAC Europe Annual meeting, Barcelona Spain. 3-7.05.2015.

Effects of Ag nanoparticles on two aquatic invertebrates. S. Cambier, K. Mehennaoui, E.J. Keuzenkamp, A. Georgantzopoulou, T Serchi, L Giamberini, A.C. Gutleb. SOT Annual meeting, New-Orleans, Louisiana, United States. 13-17.03.2016.

Surface and size-dependent effects of silver nanoparticles on behavioural and physiological responses of Gammarus fossarum and uptake evaluation with NanoSIMS50. Mehennaoui K., Georgantzopoulou A., Felten V., Garaud M., Andreï J., Cambier S., Serchi T., Contal S., Balachandran Y.L., Pain-Devin S., Giamberini L., Gutleb A.C. SOT Annual meeting, New-Orleans, Louisiana, United States. 13-17.03.2016.

Influence of Size and Surface coating on silver nanoparticles uptake by Gammarus fossarum. K. Mehennaoui, S. Cambier, J. Ziebel, N. Valle, J.S. Thoamnn, F. Guérold, L. Giamberini, A.C. Gutleb. 26th SETAC Europe Annual meeting, Nantes, France. 22-26.05.2016.

Influence of Size and Surface coating on silver and gold uptake by Gammarus fossarum. K. Mehennaoui, S. Cambier, J. Ziebel, N. Valle, J.S. Thoamnn, F. Guérold, L. Giamberini, A.C. Gutleb. Beltox Annual meeting, Louvain-La-Neuve, Belgium, 12.2016.

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Viualisation of the assimilation of nanoaprticles in biological tissues by SIMS nano-analyses. E. Lentzen, J.N. Audinot, N. Valle, A. Georgantzopoulou, K. Mehennaoui, S. Cambier, A.C. Gutleb. 6th international NanoSIMS user meeting, Utrecht, The Netherlands. 26-27.09.2016.

Influence of Size and Surface coating on silver and gold uptake by Gammarus fossarum. K. Mehennaoui, S. Cambier, J. Ziebel, N. Valle, J.S. Thoamnn, F. Guérold, L. Giamberini, A.C. Gutleb. Nanosafety 2017, Saarbrucken, Germany, 10-13.10.2017.

Identification of reference genes for RT-qPCR data normalization in Gammarus fossarum (Crustacea Amphipoda). K. Mehennaoui, S. Legay, T. Serchi, F. Guérold, L. Giamberini, A.C. Gutleb, S. Cambier. Beltox Annual Meeting, Leuven, Belgium, 01.12.2017.

Lectures

Utilisation des biomarqueurs en écotoxicologie : effets de différentes tailles de nanoparticules d’argent sur le comportement et la physiologie de Gammarus sp. K. Mehennaoui. Mentouri University, Constantine, Algeria. 2.01.2015.

Utilisation des biomarqueurs en écotoxicologie : effets de différentes tailles de nanoparticules d’argent sur les organismes aquatiques. K. Mehennaoui. Mentouri University, Constantine, Algeria. 3.01.2016.

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TABLE DES MATIERES/ TABLE OF CONTENT

ABSTRACT .................................................................................................................................................... 4

RESUME ....................................................................................................................................................... 4

AVANT PROPOS/ FOREWORD....................................................................................................................... 6

ACKNOWLEDGEMENTS ................................................................................................................................ 7

THESIS ACHIEVEMENTS .............................................................................................................................. 10

ARTICLES IN THE THESIS FRAME ...................................................................................................................... 10

BOOK CHAPTER .......................................................................................................................................... 10

ARTICLES IN THE NANOTOXICOLOGY TOPIC ....................................................................................................... 10

ARTICLES OUT OF THE TOPIC .......................................................................................................................... 10

ORAL PRESENTATIONS ................................................................................................................................. 11

POSTERS ................................................................................................................................................... 11

KEYNOTE LECTURES ..................................................................................................................................... 12

LIST OF FIGURES......................................................................................................................................... 19

LIST OF TABLES .......................................................................................................................................... 23

ABBREVIATIONS ......................................................................................................................................... 25

GENERAL INTRODUCTION .......................................................................................................................... 27

CHAPTER 1 ................................................................................................................................................. 30

PART 1: STAT OF THE ART BASED ON THE PUBLISHED REVIEW: .................................................................. 30

ABSTRACT .................................................................................................................................................. 32

1. INTRODUCTION .................................................................................................................................. 32

2. NPS TRANSFORMATIONS IN AQUATIC SYSTEMS ................................................................................. 33

3. NANOTOXICITY TOWARD AQUATIC ORGANISMS ................................................................................ 36

3.1. GENERATION OF ROS ...................................................................................................................... 36

3.2. OMICS ENDPOINTS ......................................................................................................................... 38

4. NPS TOXICITY ON FRESHWATER ORGANISMS ..................................................................................... 40

4.1. SILVER NPS (AGNPS) ........................................................................................................................ 40

4.2. GOLD NANOPARTICLES (AUNPS) ..................................................................................................... 44

5. GENERAL REMARKS AND CONCLUSIONS ............................................................................................. 46

ACKNOWLEDGEMENTS .............................................................................................................................. 48

REFERENCE ................................................................................................................................................ 48

CHAPTER 1 ................................................................................................................................................. 57

PART 2: G. FOSSARUM AS A MODEL ORGANISM IN NANOTOXICOLOGY: .................................................... 57

1. SYSTEMATIC AND IDENTIFICATION OF GAMMARUS FOSSARUM ......................................................... 58

1.1. SYSTEMATIC POSITION .................................................................................................................... 58

1.2. IDENTIFICATION OF GAMMARUS FOSSARUM .................................................................................. 60

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2. MORPHOLOGY.................................................................................................................................... 62

2.1. PROSOMA ....................................................................................................................................... 63

2.2. MESOSOMA .................................................................................................................................... 63

2.3. METASOMA .................................................................................................................................... 64

2.4. UROSOMA ...................................................................................................................................... 64

3. ANATOMY OF GAMMARUS FOSSARUM .............................................................................................. 64

3.1. NERVOUS SYSTEM AND CIRCULATORY ORGANS .............................................................................. 64

3.2. INTESTINAL CAECA AND DIGESTION SYSTEM ................................................................................... 65

3.3. GAMMARUS GILLS AND RESPIRATION ............................................................................................. 66

3.4. REPRODUCTION ORGANS, LIFE CYCLE AND DEVELOPMENT ............................................................. 67

3.4.1. ANATOMY OF THE REPRODUCTION ORGANS ............................................................................... 67

3.4.2. REPRODUCTION AND LIFE CYCLE ................................................................................................. 67

3.4.3. DEVELOPMENT ............................................................................................................................ 68

4. GAMMARUS SP. ECOLOGY .................................................................................................................. 69

5. FORAGING PLASTICITY ........................................................................................................................ 69

6. GAMMARUS FOSSARUM AS A MODEL ORGANISM IN ECOTOXICOLOGY .............................................. 70

REFERENCES .............................................................................................................................................. 71

CHAPTER 2 ................................................................................................................................................. 77

GAMMARUS FOSSARUM (CRUSTACEA AMPHIPODA) AS A MODEL ORGANISM FOR NANO-ECOTOXICOLOGY

.................................................................................................................................................................. 77

AIM OF THE STUDY .................................................................................................................................... 78

EXPERIMENTAL DESIGN ............................................................................................................................. 78

KEY FINDINGS ............................................................................................................................................ 78

ARTICLE 1 ................................................................................................................................................... 79

COMPLEMENT 1......................................................................................................................................... 91

SUPPLEMENTARY MATERIAL ...................................................................................................................... 92

PARTICLE DISPERSION AND CHARACTERISATION IN VOLVIC® WATER ....................................................................... 92

ENERGY RESERVES, DETOXIFICATION AND LPO LEVEL MEASUREMENTS ................................................................... 92

CHAPTER 3 ................................................................................................................................................. 96

G. FOSSARUM MOLECULAR RESPONSES ..................................................................................................... 96

AIM OF THE STUDY .................................................................................................................................... 97

EXPERIMENTAL DESIGN ............................................................................................................................. 97

KEY FINDINGS ............................................................................................................................................ 97

ARTICLE 2 ................................................................................................................................................... 98

1. INTRODUCTION ................................................................................................................................ 100

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2. RESULTS ........................................................................................................................................... 101

2.1. STABILITY OF THE CANDIDATE REFERENCE GENES IN G. FOSSARUM .............................................. 101

2.2. OPTIMAL NUMBER OF REFERENCE GENE FOR DATA NORMALIZATION IN G. FOSSARUM USING

GENORM ................................................................................................................................................. 103

2.3. VALIDATION OF THE SELECTED REFERENCE GENES FOR G. FOSSARUM .......................................... 104

3. DISCUSSION ...................................................................................................................................... 105

4. MATERIALS AND METHODS .............................................................................................................. 107

4.1. ORGANISMS SAMPLING AND ACCLIMATION ................................................................................. 107

4.2. AGNO3, AGNPS AND AUNPS CONTAMINATION ............................................................................. 108

4.3. GENE IDENTIFICATION AND QPCR PRIMER DESIGN ....................................................................... 108

4.3.1. DNA EXTRACTION ...................................................................................................................... 108

4.3.2. LIBRARY PREPARATION AND SEQUENCING ................................................................................ 108

4.3.3. DE NOVO ASSEMBLY.................................................................................................................. 108

4.3.4. GENE IDENTIFICATION ............................................................................................................... 109

4.3.5. PRIMER DESIGN ......................................................................................................................... 110

4.4. RNA EXTRACTION, CDNA AND RT-QPCR ........................................................................................ 110

4.5. STABILITY OF THE CANDIDATES’ REFERENCE GENES ...................................................................... 110

4.6. STATISTICAL ANALYSES ................................................................................................................. 111

5. CONCLUSIONS .................................................................................................................................. 111

ABBREVIATIONS ....................................................................................................................................... 112

REFERENCES ............................................................................................................................................ 113

COMPLEMENT 2....................................................................................................................................... 117

CHAPTER 4 ............................................................................................................................................... 118

AIM OF THE STUDY .................................................................................................................................. 119

EXPERIMENTAL DESIGN ........................................................................................................................... 119

KEY FINDINGS .......................................................................................................................................... 119

ARTICLE 3 ................................................................................................................................................. 120

1 INTRODUCTION ................................................................................................................................ 123

2 MATERIALS AND METHODS .............................................................................................................. 124

2.1 PARTICLES AND CHEMICALS ............................................................................................................. 124

2.2 PARTICLE CHARACTERIZATION ........................................................................................................... 124

2.3 ORGANISM SAMPLING AND ACCLIMATION ........................................................................................... 125

2.4 ACUTE TOXICITY TEST ...................................................................................................................... 125

2.4.1 Total and dissolved silver and gold measurements ............................................................. 126

2.4.2 Silver and gold bioaccumulation ......................................................................................... 126

2.4.3 Nanoparticle uptake........................................................................................................... 127

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2.4.4 Molecular responses .......................................................................................................... 128

2.5 STATISTICAL ANALYSES .................................................................................................................... 132

3 RESULTS ........................................................................................................................................... 132

3.1 NANOPARTICLE CHARACTERIZATION ................................................................................................... 132

3.2 ACUTE TOXICITY TEST ...................................................................................................................... 133

3.2.1 Survival .............................................................................................................................. 133

3.2.2 AgNP exposure ................................................................................................................... 133

3.2.3 AuNP exposure ................................................................................................................... 136

3.2.4 Particle uptake ................................................................................................................... 137

3.2.5 Molecular responses .......................................................................................................... 140

4 DISCUSSION ...................................................................................................................................... 144

5 CONCLUSION .................................................................................................................................... 148

COMPLEMENT 3....................................................................................................................................... 157

SUPPLEMENTARY MATERIAL .................................................................................................................... 158

1- COATING OF AGNPS AND AUNPS BY WET CHEMISTRY ...................................................................... 158

2- PARTICLE CHARACTERIZATION .......................................................................................................... 159

3- PRELIMINARY TEST: DAPHNIA MAGNA STRAUS MOBILITY INHIBITION TEST (ISO 6341:1996) ............ 163

3.1- TEST ORGANISMS .............................................................................................................................. 163

3.2- BIOLOGICAL ASSAY ............................................................................................................................. 163

3.2.1- Reference test ......................................................................................................................... 163

3.2.2- Final test ................................................................................................................................. 164

3.2.3- Results .................................................................................................................................... 164

4- ACUTE TOXICITY OF AGNO3 ON GAMMARUS FOSSARUM .................................................................. 165

CHAPTER 5 ............................................................................................................................................... 167

AIM OF THE STUDY .................................................................................................................................. 168

EXPERIMENTAL DESIGN ........................................................................................................................... 168

KEY FINDINGS .......................................................................................................................................... 168

1. INTRODUCTION ................................................................................................................................ 171

2. MATERIALS AND METHODS .............................................................................................................. 172

2.1. PARTICLES AND CHEMICALS ............................................................................................................. 172

2.2. PARTICLE CHARACTERIZATION ........................................................................................................... 173

2.3. ALDER LEAVES CONDITIONING .......................................................................................................... 173

2.4. ORGANISMS SAMPLING AND ACCLIMATION.......................................................................................... 173

2.5. TROPHIC EXPOSURE ........................................................................................................................ 174

2.5.1. EXPERIMENTAL DESIGN ............................................................................................................... 174

2.5.2. TOTAL AND DISSOLVED SILVER AND GOLD MEASUREMENTS .................................................................. 174

2.5.3. SILVER AND GOLD BIOACCUMULATION ............................................................................................ 175

2.5.4. PARTICLES UPTAKE: CYTOVIVA ® ANALYSES ...................................................................................... 175

2.5.4.1. SAMPLE PREPARATION ............................................................................................................. 175

2.5.4.2. CYTOVIVA® DARK FIELD HYPERSPECTRAL IMAGING ......................................................................... 175

2.5.5. MOLECULAR EFFECTS .................................................................................................................. 176

18

2.5.5.1. RNA EXTRACTION AND CDNA SYNTHESIS .................................................................................... 176

2.5.5.2. PRIMER DESIGN AND QUANTITATIVE REAL-TIME PCR ...................................................................... 176

2.5.6. BEHAVIOURAL RESPONSES AND OSMOREGULATION ........................................................................... 177

2.5.6.1. LOCOMOTOR ACTIVITY ............................................................................................................. 177

2.5.6.2. VENTILATION ACTIVITY............................................................................................................. 177

2.5.6.3. OSMOREGULATION ................................................................................................................. 177

2.5.6.3.1. HAEMOLYMPH SAMPLING ..................................................................................................... 177

2.5.6.3.2. HAEMOLYMPH NA+, CL- AND CA2+ CONCENTRATIONS MEASUREMENTS ........................................... 178

2.6. STATISTICAL ANALYSES .................................................................................................................... 178

3. RESULTS ........................................................................................................................................... 178

3.1. PARTICLE CHARACTERIZATION ...................................................................................................... 178

3.2. SURVIVAL ...................................................................................................................................... 178

3.3. SILVER AND GOLD BIOACCUMULATION......................................................................................... 179

3.4. CYTOVIVA® DARK FIELD HYPERSPECTRAL IMAGING....................................................................... 181

3.5. MOLECULAR EFFECTS .................................................................................................................... 182

3.6. BEHAVIOURAL RESPONSES AND OSMOREGULATION .................................................................... 182

3.6.1. LOCOMOTION AND VENTILATION ACTIVITY ............................................................................... 182

3.6.2. OSMOREGULATION ................................................................................................................... 185

4. DISCUSSION ...................................................................................................................................... 185

4.1. PARTICLE CHARACTERIZATION ...................................................................................................... 186

4.2. BIOLOGICAL ENDPOINTS ............................................................................................................... 186

5. CONCLUSIONS .................................................................................................................................. 190

CHAPTER 6 ............................................................................................................................................... 198

1. NPS SIZE, SURFACE COATING AND CHEMICAL COMPOSITION INFLUENCE THE UPTAKE ..................... 203

2. SYNTHESIS METHODS AND CHEMICAL COMPOSITION INFLUENCE AGNPS AND AUNPS EFFECTS ....... 205

3. CONTRIBUTION OF SOLUBLE IONS TO THE EFFECTS .......................................................................... 206

4. LIMITS OF THE MULTI-BIOMARKERS APPROACH ............................................................................... 207

5. CONCLUDING REMARKS AND FUTURE PERSPECTIVES ....................................................................... 208

REFERENCES ............................................................................................................................................ 209

REFERENCE .............................................................................................................................................. 213

19

LIST OF FIGURES

Chapter 1 Part 1

Figure P1. 1. Representative chemical and physical transformations of NPs when entering in natural aquatic systems:

dissolution, phosphatization, sulfidation, homo- and hetero-aggregation, and sedimentation. Important constituents

with which NPs can interact governing their fate and transport includes hardness cations (e.g., Ca2+, Mg2+), alkalinity,

phosphate and sulfide anions, pH, dissolved organic carbon (DOC), organic matter (OM) and mineral surfaces (such as

iron and manganese oxides, and clays). Legend: blue circles: Engineered NPs; yellow circles: humic substances (HM);

brown circles: natural inorganic colloids; blue lines: rigid biopolymers; gray surroundings: representing sulfidation; Mz+:

free metal ion. Adapted from(Domingos et al., 2015b) ............................................................................................... 34

Figure P1. 2. Antioxidant defense system in an animal cell (Cossu et al., 1997; Sroda, 2011; Garaud, 2015). CAT:

catalase; G6PD: glucose-6-phospho dehydrogenase; SeGPx: selenium dependent glutathione peroxidase, GPx:

glutathione peroxidase; GR: glutathione reductase; GSH/GSSG: reduced/oxidised glutathione; NAPD+/NAPDH:

oxidised/reduced nicotinamide dinucleotide phosphate; NOS: nitric oxide synthase; SOD: superoxide dismutase......... 37

Figure P1. 3. Potential routes for the generation of ROS due to the presence of NPs. 1) Internalization of NPs ROS

generation could occur due to the NPs dissolution inside the cells and/or due to the NPs photocatalytic activity. 2)

Dissolution of the NPs leads to an increase concentration of metal ions in the media; some of these metals can also be

uptake by the organisms. 3) NPs and/or their surrounding coatings can adsorb/complex other metals present in the

media, being taken up by the cells. 4) Photocatalytic activity of the NPs in the presence of UV and/or natural light. ... 38

Chapter 1 Part 2

Figure P2. 1. (A) Tree of life of the order Amphipoda (Yellow arrow) and its constituents’ suborders Gammaridea,

Senticaudata and Hyperiidea. (B) Tree of life of Gammarus fossarum (Yellow arrow) (Lifemap)............................... 58

Figure P2. 2. Geographical distribution of Gammarus fossarum in (A) Europe and (B) Luxembourg (Dohet et al., 2008;

Fauna Europaea). ...................................................................................................................................................... 59

Figure P2. 3. Determination key [in French] of Gammarus fossarum (Felten, 2003) .................................................. 60

Figure P2. 4. Distribution of Gammarus fossarum cryptic species in Europe (Adapted from Westram et al., 2011). Grey

shaded zone indicates the contact zone between Type A and B. ............................................................................... 62

Figure P2. 5. General morphology of Gammarus sp. Adapted from Felten 2003. ...................................................... 63

Figure P2. 6. Lateral view of the anatomy of Gammarus sp. Illustrating the principal organs (Schmitz, 1992). ......... 65

Figure P2. 7. Cross section of (A) Gammarus fossarum illustrating the main organs. (B) midgut and intestinal caeca

and (B) gills of G. fossarum observed with optical microscope at 60x magnification. MG: midgut, IC: intestinal caeca

(Pictures: Chauvière A. and Mehennaoui K.) ............................................................................................................... 66

Figure P2. 8. Reproduction organs of a male Gammarus and a female (Trapp, 2015). cmu: mucus cells, GA: androgen

gland, ci: incubation chamber, mt: non-differentiated mesenchymal tissue, ov1: primer vitellogenese oocyte, ov2:

secondary vitellogenese oocyte, ovd: oviduct, ovd vst: vestigial oviduct, spc: spermatocyte, spg: spermatogonium, spz:

spermatozoid, vd: spermiduct .................................................................................................................................... 67

Figure P2. 9. Gammarus fossarum male and female forming a prepopulate pair (Picture: Untereiner B. and Mehennaoui

K.) ............................................................................................................................................................................. 68

20

Chapter 2 Article 1

Figure 1. Experimental design (adapted from(Arce Funck et al., 2013). CAT: Catalase; GPx: Glutathione

Peroxidase; TAC: Total Antioxidant Activity; ACP: Acid Phosphatase; GST: Glutathione S-Transferase; CHOL:

Cholesterol; TRIG: Triglyceride: PROT: Proteins: ETS: Electron Transport System; LDH: Lactate dehydrogenase; LPO:

Lipid peroxidation; CASP-3: Caspase 3………………………………………………………………………………………………………………..82

Figure 2. Elemental distribution of 12C14N- cluster and 109Ag- ion in 300 nm cuts of gills. G. fossarum were exposed

to AgNPs (A) 23 nm, (B) 27nm, (C) AgNO3 and (D) Control (Volvic® water). Scale bar is 5 µm…………………………..85

Figure 3. G. f2 survival rates (Mean ± SD) after 72h of exposure to Ag, AgNPs 23 nm, AgNPs 27 nm, AgNPs 20

nm. Data were arcsin root square transformed. Letters (a-c) illustrate significant differences (One-way ANOVA +

Fisher LSD post hoc test at P<0.05 level of significance, n=15). ……………………………………………………………………………86

Figure 4: G .f2 haemolymph osmolality (Mean ± SD) after 72h exposure to AgNO3, AgNPs 20 nm, AgNPs 23 nm

and AgNPs 27 nm. Different letters illustrate significant differences between different treatments (One-way

ANOVA + LSD Fisher post hoc test at P < 0.05 level of significance, n=15)……………………………………………………………87

Figure 5: Behavioural responses of G.f2 after 72h of exposure to AgNO3, AgNPs 20 nm, AgNPs 23 nm, AgNPs 27

nm. (A) Ventilation (Mean pleopod beat frequency ± SD). No significant differences were detected for any of

exposure conditions (One-way ANOVA + Fischer LSD post hoc test, P < 0.05, n=10). (B) Locomotor activities

(Mean percentage of moving G.f2 ± SD). Letters illustrate significant differences. (One-way ANOVA + Fisher LSD

post hoc test at P < 0.05 level of significance, n=10)…………………………………………………………………………………………….88

Chapter 3 Article 2

Figure 3. 1. Global ranking of candidate reference genes in G. fossarum. A number (from 1 to 6) was assigned to each

stability coefficient. A mean rank was generated and error bars .............................................................................. 103

Figure 3. 2 Determination of the optimal number of reference gens for data normalization in G. fossarum exposed to

AgNO3, AgNPs 40 nm and AuNPs 40 nm. The pairwise variation (Vn/Vn+1) was calculated between normalization factors

NF/NFn+1. The recommended cut-off threshold of 0.15 was applied in this study....................................................... 104

Figure 3. 3 HSP90 expression analysis using different normalization strategies. Error bars indicate the standard errors

of the means (n=4). Different letter (a-c) indicate significant differences at P < 0.05. ................................................ 105

Chapter 4 Article 3

Figure 4. 1. G. fossarum Ag bioconcentration (mean ± SD) after 72h exposure to CIT and PEG-AgNPs 20, 40 and 80 nm.

................................................................................................................................................................................ 135

Figure 4. 2. G. fossarum au bioconcentration (mean ± SD) after 72h exposure to CIT and PEG AuNPs 20, 40 and 80 nm.

................................................................................................................................................................................ 137

Figure 4. 3. A) 300 nm cross section of G. fossarum observed under optical microscope. Red square represents the area

observed with NanoSIMS 50. Scale bar is 250 µm. B) Elemental distribution of 12C14n- clusters. C) 109Ag- ions in 300 nm

cross sections of G. fossarum gills. Animals were exposed to 10 µg. L-1 of CIT-AgNPs 40 nm and PEG-AgNPs 40 nm in

Volvic water. Scale bar is 2 µm. ................................................................................................................................ 137


observed with NanoSIMS 50. Scale bar is 250 µm. B) Elemental distribution of 12C14N- clusters. C) 109Au- ions in 300 nm

cross sections of G. fossarum intestinal caeca. Animals were exposed for 72h to 10 µg. L-1 of CIT-AuNPs 40 nm and PEG-

AuNPs 40 nm in Volvic water. Scale bar is 2 µm ........................................................................................................ 138


observed with Cytoviva darkfield hyperspectral microscope. Scale bar is 250 µm. B) G. fossarum gills viewed with

Cytoviva (60x oil immersion magnification). C) AgNPs accumulation in G. fossarum gills (red spots and white arrows).

21

Animals were exposed for 72h to 10 µg. L-1 of CIT-AgNPs 40 nm and PEG-AgNPs 40 nm in Volvic water. Scale bar is 6 µm

................................................................................................................................................................................ 139


observed with Cytoviva darkfield hyperspectral microscope. Scale bar is 250 µm. B) G. fossarum intestinal caeca viewed

with Cytoviva (60x oil immersion magnification). C) AuNPs accumulation in G. fossarum intestinal caeca (green spots

and white arrows). Animals were exposed for 72h to 10 µg. L-1 of CIT-AgNPs 40nm and PEG-AgNPs 40 nm in Volvic water.

Scale bar is 6 µm ...................................................................................................................................................... 140

Chapter 5 Article 4

Figure 5. 1. Survival rates (mean ± SD) of G. fossarum exposed for 15 days to 0.5 and 5 µg. L-1 of CIT and PEG-AgNPs

and PEG-AuNPs 40 nm ............................................................................................................................................ 179

Figure 5. 2 A) 300 nm cross section of G. fossarum observed under optical microscope. Red square represents the area

observed with cytoviva darkfield hyperspectral microscope. Scale bar is 250 µm. B) G. fossarum gills or intestinal caeca

viewed with Cytoviva (60x oil immersion magnification). C) AgNPs accumulation in G. fossarum gills (red spots and

white arrows). Animals were exposed for 72h to 10 µg. L-1 of CIT-AgNPs 40nm and PEG-AgNPs 40 nm in Volvic water.

Scale bar is 6µm. ...................................................................................................................................................... 181


observed with CytoViva darkfield hyperspectral microscope. Scale bar is 250 µm. B) G. fossarum intestinal caeca viewed

with CytoViva (60x oil immersion magnification). C) AuNPs accumulation in G. fossarum caeca (red spots and white

arrows). Animals were exposed for 72h to 10 µg. L-1 of CIT-AgNPs 40nm and PEG-AgNPs 40 nm in Volvic water. Scale bar

is 6µm. .................................................................................................................................................................... 182

Figure 5. 4. Behavioural responses of G. fossarum exposed for 15 days to CIT and PEG-AgNPs and AuNPs 40 nm. A)

Locomotor activity (mean percentage of moving G. fossarum ± SD) and b) Ventilation (mean pleopods beat frequency

± SD). Different letters (a-f) indicates significant differences (one-way ANOVA + Tukey HSD post hoc test at P < 0.05 level

of significance, n = 10). ............................................................................................................................................ 184

Chapter 6

Figure 6. 1. Physiological and behavioural effects of synthetic and biological AgNPs on G. fossarum exposed for 72h:

summary of results presented in Chapter 2. G. fossarum were exposed for 72h to synthetic AgNPs 20 and 200 nm and

biologically synthetized from plant leaf extract AgNPs 23 and 27 nm. Effects on antioxidant responses (GPx, TAC, CAT),

defense mechanism (GST, ACP), cellular damage (LDH, CASP3, LOOH), energy reserves (Prot, Chol, Trig, ETS),

osmoregulation and behaviour (Locomotion and ventilation) were assessed............................................................. 200

Figure 6. 2. Influence of size (20, 40 and 80 nm) and surface coating (CIT and PEG) of AgNPs and AuNPs on their uptake,

tissue distribution and molecular effects of G. fossarum exposed for 72h: summary of the results presented in Chapter

4. G. fossarum were exposed for 72h to up to 50 µg. L-1 of AgNPs and AuNPs in absence of food. Influence of size and

surface coating on bioaccumulation was assessed using ICP-MS, internal distribution of AgNPs and AuNPs were

evaluated using NanoSIMS50 and Cytoviva, and molecular effects were assessed using RT-qPCR. A set of stress-related

genes expression including genes implied in cytoskeleton trafficking (Actin, TUB, UB), exoskeleton cuticle (Chitinase),

antioxidant defence (CAT, MnSOD, CuZnSOD, GPx7), general stress (GST, HSP90, GAPDH), DNA damage and repair

(Gadd45, NfkB, c-jun, P53), lysosomes (Cathepsin L), osmoregulation and respiration (Na+K+ATPase, HEM) was used.

................................................................................................................................................................................ 201

Figure 6. 3. Sub-chronic toxicity of CIT and PEG-AgNPs and CIT and PEG-AuNPs 40 nm on molecular, physiological and

behavioural responses of G. fossarum: summary of the results presented in Chapter 5. G. fossarum were exposed for

15 days to 0.5 and 5 µg.L-1 of CIT and PEG-AgNPs and AuNPs 40 nm in presence of food. Bioaccumulation was assessed

using ICP-MS, internal distribution of AgNPs and AuNPs were evaluated using Cytoviva and molecular effects were

assessed using a set of stress-related genes expression including genes implied in cytoskeleton trafficking (Actin, TUB,

22

UB), exoskeleton cuticle (Chitinase), antioxidant defence (CAT, MnSOD, CuZnSOD, GPx7), general stress (GST, HSP90,

GAPDH), DNA damage and repair (Gadd45, NfkB, c-jun, P53), lysosomes (Cathepsin L), osmoregulation and respiration

(Na+K+ATPase, HEM). ............................................................................................................................................... 202

Supplementary materials

Figure S 1. Size distribution of Ag NPs 23 nm (A), 27 nm (B), 20 nm (C) and 200 nm (D) in Volvic® water expressed as

particle concentration x 106.mL-1. The red error bars indicate the ± SD of the mean of triplicate measurements......... 96

Figure S 2 PEG coating on metal Nanoparticles (here silver NPs). SEM shows clearly the conformal silver coating with

the PEG layer. NTA analysis shows a small size increasing for the 40 and 80 nm silver nanoparticles as a consequence of

their coating with the PEG layer. The sensitivity of NTA was not enough to resolve the size increase for 20 nm

nanoparticles. .......................................................................................................................................................... 158

Figure S 3. Size distribution of CIT-AgNPs 20 nm (A), CIT-AgNPs 40 nm (B) and CIT-AgNPs 80 nm (C) in Volvic Water

(T0h) and CIT-AgNPs 20 nm (D), CIT-AgNPs 40 nm (E) and CIT-AgNPs 80 nm (F) in Volvic water after 24h of incubation.

Size distribution is expressed as particle concentration E6.mL-1. The red errors bars indicate ±SD of the mean of triplicate

measurements ......................................................................................................................................................... 159

Figure S 4. Size distribution of PEG-AgNPs 20 nm (A), PEG-AgNPs 40 nm (B) and PEG-AgNPs 80 nm (C) in Volvic Water

(T0h) and PEG-AgNPs 20 nm (D), PEG-AgNPs 40 nm(E) and PEG-AgNPs 80 nm (F) in Volvic water after 24h of

incubation. Size distribution is expressed as particle concentration E6.mL-1. The red errors bars indicate ±SD of the mean

of triplicate measurements ...................................................................................................................................... 160

Figure S 5. Size distribution of CIT-AuNPs 20 nm (A), CIT-AuNPs 40 nm (B) and CIT-AuNPs 80 nm (C) in Volvic Water

(T0h) and CIT-AuNPs 20 nm (D), CIT-AuNPs 40 nm (E) and CIT-AuNPs 80 nm (F) in Volvic water after 24h of incubation.

Size distribution is expressed as particle concentration E6.mL-1. The red errors bars indicate ±SD of the mean of triplicate

measurements ......................................................................................................................................................... 161

Figure S 6. Size distribution of PEG-AuNPs 20 nm (A), PEG-AuNPs 40 nm (B) and PEG-AuNPs 80 nm (C) in Volvic Water

(T0h) and PEG-AuNPs 20 nm (D), PEG-AuNPs 40 nm(E) and PEG-AuNPs 80 nm (F) in Volvic water after 24h of

incubation. Size distribution is expressed as particle concentration E6.mL-1. The red errors bars indicate ±SD of the mean

of triplicate measurements ...................................................................................................................................... 162

Figure S 7. Effects AgNO3 on survival of Gammarus fossarum collected in A) June, B) September and C) November after

72h of exposure....................................................................................................................................................... 165

23

LIST OF TABLES

Chapter 1 Part 2

Table P2. 1. Classification of the species Gammarus fossarum .................................................................................. 59

Chapter 2 Article 1

Table 2.1. Particles size distribution (mode ± SD, 3 replicates) and ζ potential of AgNPs in Volvic® water………..84

Table 2.2: LC50 values with 95% confidence intervals for G.f1 and G.f2 exposed to AgNO3 and AgNPs20 nm,

AgNPs 23 nm, AgNPs 27nm and AgNPs 200 nm for 72h……………………………………………………………………………………84

Table 2.3: Total and dissolved Ag concentrations (mean ± SD) and dissolution rates of AgNO3 and AgNPs after

72h of exposure in Volvic water. Different letters illustrate significant differences between different treatments

(Two-way ANOVA + Tukey HSD post-hoc test at P < 0.05 level of significance, n=3)…………………………………………….85

Table 2.4: G.f2 Ag Bioconcentration (Mean± SD) after 72h exposure to AgNO3, AgNPs 20 nm, AgNPs 23 nm,

AgNPs 27 nm. Different letters illustrate significant differences between different treatments (Kruskal-Wallis

ANOVA + Mann-Whitney U test at P < 0.05 level of significance, n=3)…………………………………………………………………86

Table 2.5: Mean values (±SD) of biomarkers measured in Gf2 exposed to AgNO3, AgNPs 20nm, AgNPs 23nm and

AgNPs 27 nm for 72h. Biomarkers in italic were analysed using One-way ANOVA and Tukey post hoc test, whereas

the others were analysed using Kruskal-Wallis ANOVA and Mann-Whitney U test. No significant differences were

detected for any of the exposure conditions. a: mg. g fresh weight-1; b: µmol O2. g proteins-1. h-1.; c: µmol p-

nitrophenol. g protein-1.h-1; d: µmol CDNB.min-1.g-1 proteins; e: µmol NADPH.g protein-1.min-1; f: mmol Trolox

equivalent.g protein-1; g: mmol H2O2.g proteins-1.min-1; µmol NADH. g proteins-1. h-1; h: µmolpNA. g proteins-1.h-1;

j: nmol TBH. g proteins-1……………………………………………………………………………………………………………………………………..87

Chapter 3 Article 2

Table 3. 1.Ranking of candidate reference genes according to the five algorithms used ......................................... 102

Table 3. 2 Identification of Gammarus fossarum gene sequences ........................................................................... 108

Table 3. 3 List of primers of the candidate reference genes and target gene HSP90 ................................................ 109

Chapter 4 Article 3

Table 4. 1. Specific primer pairs used for RT-qPCR analyses on Gammarus fossarum exposed for 72h to AgNPs and

AuNPs (coated with CIT or PEG, sizes: 20, 40 and 80 nm, F: forward sequence, R: reverse sequence) ...................... 130

Table 4. 2. Size distribution of particles (mode ± SD, 3 replicates) and ζ potential of AgNPs and AuNPs in Volvic water

(exposure medium) at T0h and T24h. ...................................................................................................................... 133

Table 4. 3. Total ag concentrations (mean ± SD) and recovery rates (mean ± SD) of AgNO3, CIT-AgNPs and PEG-AgNPs

in Volvic water (exposure medium). ........................................................................................................................ 134

Table 4. 4. Relative gene expression of G. fossarum exposed for 72h to CIT-AgNPs and PEG-AgNPs 20, 40 and 80 nm

................................................................................................................................................................................ 142

Table 4. 5. Relative gene expression of G. fossarum exposed for 72h to CIT-AuNPs and PEG-AuNPs 20, 40 and 80 nm

................................................................................................................................................................................ 143

24

Chapter 5 Article 4

Table 5. 1. G. fossarum Ag and Au uptake (mean ± SD) after 15 days of exposure to CIT and PEG-AgNPs and AuNPs 40

nm. ......................................................................................................................................................................... 180

Table 5. 2. Relative gene expression of G. fossarum exposed for 15 days to CIT- and PEG-AgNPs 40 nm and CIT and

PEG-AuNPs 40 nm ................................................................................................................................................... 183

Table 5. 3. Haemolymph [Cl-], [Na+] and [Ca2+] of G. fossarum exposed for 15 days to CIT and PEG-AgNPs 40 nm and

CIT and PEG-AuNPs 40 nm. ...................................................................................................................................... 185

Supplementary materials

Table S 1. Physico-chemical parameters of Volvic® water ......................................................................................... 95

Table S 2 D. magna reference test ........................................................................................................................... 163

Table S 3. EC50 values obtained after 48h exposure of D. magna to AgNPs and AuNPs .......................................... 164

25

ABBREVIATIONS

ATP Adenosine triphosphate

cDNA Complementary DNA

CYP450 Cytochrome P450

DTT Dithioreitol

EF1 Elongation factor 1 alpha

EMBL European Molecular Biology Laboratory

EST Expressed Sequence Tag

FC Fold change

FNR National Research Funds – Luxembourg

MIQe Minimum information for publication of qPCR experiments

mRNA Messenger ribonucleic acid

PCR Polymerase Chain Reaction

qPCR Quantitative real-time polymerase chain reaction

RIN RNA integrity number

RNA Ribonucleic acid

RNA-seq RNA sequencing

SI Supporting information

TEM Transmission electron microscope

TF Transcription factor

UV Ultraviolet

AgNPs Silver nanoparticles

AuNPs Gold nanoparticles

AgNO3 Silver nitrate

Ag2S Silver sulphide

DMSO Dimethyl Sulfoxide

NTA Nanoparticles tracking analysis

DLS Dynamic light scattering

HIM-SIMS Helium Ion microscopy – secondary ion mass spectrometer

EDTA Ethylenediaminetetraacetic acid

HNO3 Nitric acid

TAC Total antioxidant capacity

HCl Chloride acid

CIT Citrate

PEG Polyethylene glycol

NanoSIMS 50 Nano secondary ion mass spectroscopy

ICP-MS Inductively coupled plasma – mass spectrometer

ACP Acid phosphatase

CAT Catalase

OS Oxidative stress

NPs Nanoparticles

ENPs Engineered nanoparticles

NMs Nanomaterials

HM Humic acid

DOC Dissolved organic carbon

TOC Total organic carbon

OM Organic matter

FPOM Fine particle organic matter

CASP Caspase

PEC Predicted environmental concentration

L(E)C 50 Lethal or effective concentration that impact 50% of the exposed population

ETS Electron transport system

GR Glutathione reductase

GSH Reduced glutathione

GST Glutathione S-transferase

NOEC Non-observable effects concentrations

H2O2 Hydrogen peroxide

HSP Heat shock protein

MDA Malondialdehyde

ROS Reactive oxygen species

SOD Superoxide dismutase

LPO Lipid peroxidation

GPx Glutathione peroxidase

27

GENERAL INTRODUCTION

In the recent decades, nanotechnology has emerged as a fast-growing sector impacting key economical fields

and providing new engineered nano-enabled products, constituted by nanoparticles (NPs), with novel and

unique functions that reach the market every day. NPs are defined as compound presenting at least one

dimension less than 100 nm (Klaine et al., 2008).

The use of nanomaterials is increasingly and continuously growing as they are used in different areas such as

electronics, medicine, environmental technology, etc. Silver nanoparticles (AgNPs) are among the most

promising group of NPs. AgNPs are used in different kind of daily-life products such as textiles, food

packaging, healthcare products, etc., mostly for their antibacterial properties. AuNPs are being investigated

for their unique optical properties and are used as contrast agents in electron microscopy, optical sensors,

catalysts, and for therapeutic uses. The increasing use of AgNPs and AuNPs lead to their inevitable release in

the environments and may reach the aquatic ecosystems where they may represent a threat for aquatic

organisms. The potential toxicity of nanomaterials is of high societal and scientific interest and can be related

to many properties such as the size, the mass, surface area and characteristics, aggregation and

agglomeration (Georgantzopoulou et al., 2013). Their toxicity could also be linked to less known or less

understood parameters like released ions and adhering molecules which could have an influence on their

toxicity by interfering with their uptake and fate on living organisms.

The PhD project “NANOGAM” is a FNR funded project closely linked up to the “CORE2012 NANION” and to

“FP7 FUTURENANONEEDS (FNN)” projects that aim to obtain knowledge to understand some of the

processes and factors involved in NPs uptake and toxicity. The specific objectives of NANION are:

• Leaching ions from NPs have their own spectrum of toxicity and effects can be clearly separated

between free ions in the exposure media and the NPs (Georgantzopoulou, 2015).

• Within NANION, a set of AgNPs and AuNPs comprising different size classes and coated with various

surfaces (citrate and polyethylene glycol) are used. Prior to any testing, these NPs were carefully

characterized (zeta-potential, agglomeration, aggregation, etc.) in the relevant exposure media. The

ion release from uncoated and coated NPs (Au does not release ions) were studied in dependency of

exposure media and related to toxicity endpoints ranging from molecular to organism level.

• Biomolecules attach easily and quickly to the surface of NPs. This so-called corona influences uptake,

kinetics, distribution within the organism and thereby finally impact endpoints such as biochemical

biomarkers but also reproduction, behaviour of the organism, and other relevant whole organism

endpoints. The uptake kinetics, distribution of the NPs within the organism and toxicity for a range

of relevant endpoints were studied and related to physico-chemical properties of the NPs.

28

Generally, NANOGAM aims at contributing to the understanding in how far surface properties of NPs

(physical parameters, ion leaching, biological molecules attached, etc.) affect and interfere with the toxicity

of NPs in a very relevant freshwater model organism, the crustacean Gammarus fossarum. G. fossarum was

selected as test species due to its large distribution in Europe (Barnard and Barnard, 1983), high abundance

(Felten et al., 2008; Kunz et al., 2010), clear sexual dimorphism, easiness of identification to the species level,

collection and handling, the high sensitivity to a large range of toxicants and their major functional role in

ecosystems. Indeed, as shredders, they play a key role in the litter breakdown process and thus in freshwater

food chain and nutrient cycling (Forrow and Maltby, 2000; Vellinger et al., 2012a). Consequently, Gammarus

sp. could provide valuable information about the potential effects of studied contaminants on some other

taxa in aquatic ecosystem communities (Vellinger et al., 2012b) To our knowledge, only few studies have

been done on the ecotoxicity of AgNPs and AuNPs in these species (Andreï et al., 2016; Baudrimont et al.,

2017; Bundschuh et al., 2011; Park et al., 2015).

The “NanoGAM” project aims at investigating the characteristics of the key physico-chemical parameters and

surface functionalities of a well-chosen list of AgNPs and AuNPs that control uptake, and dependent

biological effects of different complexity (using a battery of biomarkers) on G. fossarum. Starting from a set

of commercially available NPs that will form the baseline for the studies proposed within NANOGAM.

The following questions have been raised within the present project:

• Is G. fossarum a valuable model to study the effects of nanoparticles?

• How do particle size, surface coating, synthesis method and chemical composition contribute to the

toxicity and effects of AgNPs and AuNPs?

• To what extent does leaching ions play a role in the toxicity of AgNPs?

• What are the acute effects of AgNPs and AuNPs, with different sizes (20, 40 and 80 nm) and two

different coatings (CIT and PEG), on G. fossarum?

• What are the sub-chronic effects of AgNPs and AuNPs on molecular, physiological and behavioural

responses of G. fossarum?

These research questions are addressed in the following chapters:

• Chapter 1 is a critical overview on biochemical effects of AgNPs and AuNPs on aquatic organism (Part

1) with a focus on G. fossarum, the model organism used in the present PhD project (Part 2)

• In Chapter 2, the effects of a well-characterized and well-studied set of AgNPs are used in order to

evaluate their effects on G. fossarum through a multi-biomarker approach. This study allows also the

assessment of the contribution of leaching ions on the toxicity and effects of AgNPs. Furthermore,

the internal distribution of AgNPs in G. fossarum is discussed.

• As G. fossarum is to our best knowledge a non-sequenced species, it was necessary to identify target

stress-related genes for RT-qPCR experiments. For accurate analyses, it was necessary to identify a

set of reference genes for RT-qPCR normalization and results are presented in Chapter 3.

29

• Chapter 4 provides evidence of the influence of AgNPs and AuNPs size and surface-coating on their

uptake and internal tissue distribution in G. fossarum. Furthermore, the acute effects on the

molecular responses are investigated.

• Chapter 5 deals with the sub-chronic effects of AgNPs and AuNPs on G. fossarum. A multi-biomarker

approach including molecular, physiological and behavioural responses was applied to assess the

effects of AgNPs and AuNPs. Additionally, tissues distribution of NPs is evaluated

• Finally, Chapter 6 discuss the knowledge acquired in the present work and the relevance of the

current findings. Conclusions and future perspectives are also addressed.

Chapter 1 Part1: NPs in the aquatic environment

30

CHAPTER 1

Part 1: Stat of the art based on the published review:

Gançalo Vale, Kahina Mehennaoui, Sébastien Cambier, Giovanni Libralato, Stéphane Jomini, Rute F.

Domingos. Manufactured nanoparticles in the aquatic environment – biochemical responses on

freshwater organisms: a critical overview. 2016. Aquatic toxicology 170, 162-174.

The published review is presented in Annexe 1

CHAPTER 1

Part 1: Stat of the art based on the published review:

Gançalo Vale, Kahina Mehennaoui, Sébastien Cambier, Giovanni Libralato, Stéphane Jomini,

Rute F. Domingos. Manufactured nanoparticles in the aquatic environment – biochemical

responses on freshwater organisms: a critical overview. 2016. Aquatic Toxicology 170, 162-174.

The published review is presented in Annexe 1


31

Manufactured nanoparticles in the aquatic environment –Biochemical responses on freshwater

organisms: A critical overview1

Gonçalo Valea,b*, Kahina Mehennaouic,e, Sebastien Cambierc, Giovanni Libralatod, Stéphane

Jominie, Rute F. Domingosa,f

a) Centro de Química Estrutural, Instituto Superior Técnico, Universidade de Lisboa, Torre Sul, Av. Rovisco Pais,

1049-001 Lisboa, Portugal. Gonçalo Vale: [email protected]

b) Department of Molecular Genetics, University of Texas Southwestern Medical center, Harry Dallas, TX 75390,

USA (present address)

c) Luxembourg Institute of Science and Technology, Environmental Research and Innovation (ERIN) Department,

Belvaux, Luxembourg. Kahina Mehennaoui: [email protected]; Sebastien Cambier:

[email protected]

d) Department of Environmental Sciences, Informatics and Statistics, University Ca’ Foscari Venice, Via Torino, 155,

30172, Mestre, Venice, Italy. Giovanni Libralato: [email protected]

e) Laboratoire Interdisciplinaire des Environements Continentaux (LIEC), Université de Lorraine, UMR 7360,

Campus Bridoux rue du Général Delestraint, 57070 Metz, France. Stéphane Jomini:

[email protected]

f) Institut de Physique du Globe de Paris, Sorbonne Paris Cité, UMR CNRS 7154, Université Paris Diderot, 75205

Paris Cedex 05, France. Rute F. Domingos: [email protected] (present address)

*Corresponding author

Email: [email protected]

1 Vale, G., Mehennaoui, K., Cambier, S., Libralato, G., Jomini, S., Domingos, R.F., 2016. Manufactured

nanoparticles in the aquatic environment-biochemical responses on freshwater organisms: A critical overview.

Aquat. Toxicol. 170, 162–174. doi:10.1016/j.aquatox.2015.11.019


32

ABSTRACT

The enormous investments in nanotechnology have led to an exponential increase of new manufactured

nano-enabled materials whose impact in the aquatic systems is still largely unknown. Ecotoxicity and

nanosafety studies mostly resulted in contradictory results and generally failed to clearly identify biological

patterns that could be related specifically to nanotoxicity. Generation of reactive oxygen species (ROS) is one

of the most discussed nanotoxicity mechanism in literature. ROS can induce oxidative stress (OS), resulting

in cyto- and genotoxicity. The ROS overproduction can trigger the induction of anti-oxidant enzymes such as

catalase (CAT), superoxide dismutase (SOD) and glutathione peroxidases (GPx), which are used as biomarkers

of response. A critical overview of the biochemical responses induced by the presence of NPs on freshwater

organisms is performed with a strong interest on indicators of ROS and general stress. A special focus will be

given to the NPs transformations, including aggregation, and dissolution, in the exposure media and the

produced biochemical endpoints.

Keywords: nanoparticles, transformations, freshwater organisms, nanotoxicity, reactive oxygen species

(ROS), anti-oxidant enzymes

1. INTRODUCTION

Nanotechnology has emerged as a fast growing sector impacting key economical fields and providing new

engineered nano-enabled products, constituted by nanoparticles (NPs), with novel and unique functions that

reach the market every day (Bour et al., 2015). NPs are defined as materials with a size between 1 and 100

nm on at least one dimension, having unique physico-chemical properties differing from their bulk forms due

to their greater surface area to volume ratio (Hood, 2004; The European Commision, 2011). The size related-

properties results in larger reactivity and higher mobility, leading to numerous applications in medical

diagnostics, electronics, computers, cosmetics and environmental remediation (Rauscher et al., 2015). The

worldwide consumption of NPs is expected to grow from 225,060 metric tons in 2014 to nearly 584,984

metric tons in 2019 representing an annual growth rate of 21.1 % (BCC RESEARCH, 2017). Although

impressive, these numbers are in fact “expected” values obtained by estimation or modelling. The lack of

legislation for nanotechnologies gives the manufacturers no onus to reveal the real figures, thus, indeed,

these predicted values are most probably significantly higher. The absence of real numbers hinders the

prediction of the NPs amount that are actually being released into the environment (Piccinno et al., 2012).

Even though several studies have been performed with the goal of modelling NPs environmental

concentrations (Gottschalk et al., 2009, 2013; Yang et al., 2016), they should only be considered as guidelines,

since they derive from uncertain data about the NPs production (often obtained by surveys to producers)

and extrapolations used to scale up regional to worldwide amounts (Gottschalk et al., 2013; Keller et al.,

2010; Piccinno et al., 2012).


33

When released in natural media NPs will be subjected to a dynamic physical and chemical environment that

consequently results in different and unknown endpoints far from their pristine or as released state.

Therefore, environments and living organisms are not facing pristine manufactured NPs but rather

transformed nano-enabled products, which is factually accepted but so far neglected. In fact, the large

majority of the physico-chemical and toxicity data obtained so far was focused on simple nanoscale particles

and not on relevant nano-enabled products. This includes not only the NP embedded in the manufactured

matrix but also the materials resulting from the interaction with biotic and abiotic (bio)molecules composing

the natural systems (Nasser and Lynch, 2016). To further complicate the interpretation of the NPs studies,

there are two distinct mechanisms that should be considered but are not easily differentiated:

• Chemical toxicity by the release of possible ions and/or formation of reactive oxygen species (ROS)

(Fu et al., 2014)

• Physical stress or stimuli caused by NPs size, shape and surface properties (Libralato, 2014; Vale et

al., 2014).

These materials are generally associated with cellular perturbations such as ROS generation, gene expression

and proteome profiles alterations. For these reasons, the NPs escalating production and applications has

raised concerns about their environmental and human safety, which have led to large investments in

nanosafety-related projects resulting in a considerable amount of data assessing their potential hazard

(Savolainen et al., 2013). However, the establishment of relationships between bioavailable NP-containing

species and the specific bioadverse or biocompatible endpoints is still lacking, mainly since the effects are

NP-dependent and also specie-dependent (Burić et al., 2015).

This work provides an overview of the latest studies on the impact of NPs onto aquatic environment with a

focus on freshwater ecosystems, considered by many as the ultimate sink of these particles, with a special

focus:

• NPs transformations and characterization in the different test media,

• Toxicological effects such as generation of ROS, genotoxicity, transcriptomic and proteomic changes.

This survey is focused on metallic NPs including silver nanoparticles (AgNPs) and gold nanoparticles

(AuNPs), mostly due to the great number of studies dedicated to these particles.

2. NPs TRANSFORMATIONS IN AQUATIC SYSTEMS

NPs can enter in an aquatic compartment from:

• Wastewater treatment plants effluents,

• Direct use (e.g., application of NPs-containing paintings on boats),

• Deposition from the air compartment.


34

When entering aquatic compartment, NPs will be exposed to a highly dynamic physical and chemical

environment that leads to several transformations that will change their pristine or as released physico-

chemical properties (Figure P1. 1). These transformations, including dissolution, aggregation and

sedimentation, are dependent on both physico-chemical properties of the NPs and those of the environment

into which they were released (Auffan et al., 2012).

Colloidal particles, including organic and inorganic matter, are ubiquitous in the aquatic environment and can

be originated from both natural and anthropic sources. These colloids can strongly interact with NPs, thereby

determining their forms over space and time (dynamic speciation), and greatly affecting their bioavailability.

Thus, the NPs will have a specific speciation in each environmental compartment, and this speciation is always

dynamic with reaction rates that depend upon the chemical nature and physical sizes of the engineered and

natural colloids (Levard et al., 2012). Although it is clear that dynamic speciation must be considered in order

to make relevant predictions of NPs fate, toxicity and risk, was mostly neglected (Domingos et al., 2015b).

Figure P1. 1. Representative chemical and physical transformations of NPs when entering in natural aquatic

systems: dissolution, phosphatization, sulfidation, homo- and hetero-aggregation, and sedimentation.

Important constituents with which NPs can interact governing their fate and transport includes hardness cations (e.g., Ca2+, Mg2+), alkalinity, phosphate and sulfide anions, pH, dissolved organic carbon (DOC), organic matter (OM) and mineral surfaces (such as iron and manganese oxides, and clays). Legend: blue circles: Engineered NPs; yellow circles: humic substances (HM); brown circles: natural inorganic colloids; blue lines: rigid biopolymers; gray surroundings: representing sulfidation; Mz+: free metal ion. Adapted from(Domingos et al., 2015b)


35

Dissolution, which is one of the main transformations of metallic NPs such as AgNPs, is mainly due to:

• The formation of partially soluble metal-oxide (Domingos et al., 2013; Heinlaan et al., 2008; Wang et

al., 2015),

• The oxidation of the particle constituents (Dale et al., 2013; Ma et al., 2013; Wang et al., 2013),

• The complexation of the metal particle constituent by complexants present in the environmental

compartment or even in the NPs embedded matrix (including the manufactured stabilizers)

(Domingos et al., 2013, 2015a).

The sulfidation of the metallic NPs can retard their oxidation and, thus, their dissolution (Dale et al., 2013;

Ma et al., 2013; Wang et al., 2013). This dissolution mechanism results in the release of toxic cations, such

that their persistence is reduced but the toxicity is increased. Evidently, complete dissolution of the NPs

allows the prediction of their impact using already existing models for metal speciation and toxicity.

Aggregation is another critical transformation, which, mainly is occurring by interaction with naturally bio- or

geo-macromolecules, affect NPs size and surface chemistry. For example, organic matter (OM) provides both

charge and steric stabilization (Domingos et al., 2009) of the NPs, although they may also result in bridging

flocculation in presence of multiple charged cations and anions (Domingos et al., 2010). OM effects are

complex and difficult to predict; however, it is of extremely importance to explore these interactions since

OM concentrations are typically orders of magnitude higher in concentration than engineered NPs, and, thus,

likely to substantially modify their properties and behaviours.

Dissolution and aggregation are dynamic processes that can decrease the NPs available surface area, thereby

decreasing their reactivity. However, this decrease is dependent on the surface properties, particle number,

size distribution, and the fractal dimensions of the aggregate (Hotze et al., 2010). The NP size will affect its

bioavailability to the organisms; when aggregates become too large for direct transport across the cell wall

and/or membrane, uptake may be prevented, whereas partial dissolution, which will lead to smaller sizes,

would facilitate this cellular transport. Since these transformations are most often not in equilibrium, they

require real-time kinetic measurements, limiting the methodology to be used:

• The storage of whole unfractionated samples for ion analysis may not be possible since the

dissolution rate may be fast or not attaining the equilibrium during the experimental time,

• The aggregation rate can be fast or the aggregates size distribution may not reach equilibrium within

the experimental time window.

Despite the large number of studies focused on nanotoxicology, most of them disregard the particles kinetic

physicochemical characterization under the exposure conditions, hindering the establishment of crucial

predictive structure-activity relationships that can be used afterwards in the categorization and function for


36

risk assessment studies. In absence of these realistic studies, two less constrain conditions were used to select

the literature studies that will be discussed in this critical overview:

• Studies where the NPs characterization was performed in the same media as the bioassays;

• Quantification of the NPs dissolution for studies using metallic NPs with propensity for dissolution

such as AgNPs.

3. NANOTOXICITY TOWARD AQUATIC ORGANISMS

The increasing use of engineered NPs in consumers’ products such as cosmetics, paints, food, computers,

medicine, etc. lead to their increasing release in aquatic environments and raise concerns about their

potential toxicity of aquatic organisms (Klaine et al., 2008; Manzo et al., 2013; Nowack et al., 2012).

3.1. Generation of ROS

Despite the large number of studies on NPs toxicity both in cell line systems and organisms, a complete

understanding about the mechanisms behind is still lacking (Bour et al., 2015; Fu et al., 2014; Manke et al.,

2013; Schultz et al., 2014). ROS generation, whose overproduction can lead to oxidative stress (OS) in the

organism tissues, is unquestionably the most studied nanotoxicity mechanism.

Molecular oxygen is used as an oxidizing agent for the production of adenosine triphosphate (ATP) in the

organism cells, being afterwards reduced to water. The non-reduced oxygen results in the formation of

superoxides (O"#.) that can be further converted to hydroxyl radicals(HO.), which has the highest reduction

potential of all the physiological relevant ROS. When under control, these species are easily scavenged by

• Antioxidant agents such as polyphenols (Fu et al., 2014; Lipinski, 2011) (e.g., elimination ofHO.),

• Enzymes such as superoxide dismutase (SOD), catalase (CAT) and glutathione peroxidase (GPx)

(Barata et al., 2005; Regoli et al., 2002; Vasseur and Leguille, 2004).

The SOD enzymes catalyse the dismutation of O"#. into oxygen or hydrogen peroxide (H2O2), which is

decomposed by CAT into water and oxygen. Even though H2O2 is less reactive than the radical species, is still

a strong oxidant that needs further elimination. The GPx, also plays a role in the detoxification of H2O2 by

using glutathione (GSH) as a reductant. During the process, GSH is oxidized and converted to glutathione

disulfide (GSSG) being latter reduced back to GSH by glutathione S-transferase enzymes (GST), thus

completing the cycle (Brigelius-Flohé and Maiorino, 2013; Deponte, 2013). The antioxidant-enzymes activity

is considered a reflection of the redox state of the cells and is frequently studied as a biomarker of OS (Figure

P1. 2).


37

Figure P1. 2. Antioxidant defense system in an animal cell (Cossu et al., 1997; Sroda, 2011; Garaud, 2015). CAT: catalase; G6PD: glucose-6-phospho dehydrogenase; SeGPx: selenium dependent glutathione peroxidase, GPx: glutathione peroxidase; GR: glutathione reductase; GSH/GSSG: reduced/oxidised glutathione; NAPD+/NAPDH: oxidised/reduced nicotinamide dinucleotide phosphate; NOS: nitric oxide synthase; SOD: superoxide dismutase.

When ROS is overproduced beyond the organism antioxidant response capacity, it leads to several

deleterious effects on the cells components such as lipids, proteins and DNA, possibly resulting in lipid

peroxidation, apoptosis and/or cancer initiation processes, respectively. The production of ROS can be

enhanced by the presence of NPs, depending mainly on their size, aggregation, solubility and coating. It is

commonly accepted that smaller particles can easily penetrate cell membranes, and thus induce cytotoxicity

(Klaine et al., 2008). However, this correlation between size and toxicity is still controversial. For instance, a

size dependent toxicity of 5-10 nm AgNPs on Tetrahymena pyriformis compared to 15-25 nm AgNPs was

reported (Shi et al., 2013) while no size-dependent response on Danio rerio was obtained when exposed to

20, 50 and 110 nm (Bowman et al., 2012). Most studies show that toxicity increases with decreasing particle

size. However, others reported that either the size has no role on toxicity or that smaller NPs are less toxic

(Ivask et al., 2014). A consensus about the size effect is still lacking, and, most probably, will be unlikely to be

attained since the effects seem to be NP- and even specie-dependent (Burić et al., 2015).

The dissolution of metallic NPs such as AgNPs and CuNPs results in the release of Cu and Ag ions, which are

known to catalyse Fenton, Fenton-like and Haber-Weiss reactions, leading to the formation of ROS (Fu et al.,

O2

O2⦁ -

H2O2

OH⦁

H2O

R ⦁

R’

H2O 2GSH

GSSG

NADP+

NADPH

2GSH

GSSG

NADP+

NADPH

ROO⦁ /ROOH

ROH/ONOH

GPx Grd G6PD

GR G6PDSeGPx

SODO2

2H+

e-

CAT H2O+½O2

Men+

Me(n+1)+ +OH-

Fenton

reaction

NO⦁

ONOO-/ONOOH

NOSe-

nitrocompounds

Organicmolecules


38

2014; Lipinski, 2011; Wang et al., 2013). Moreover, the highly reactive surface of NPs and the presence of

manufactured and/or natural coatings can lead to the adsorption/complexation of trace metals present in

the environmental compartment also acting as a catalyzer platform to the above-mentioned reactions, and

thus increasing the concentration of ROS in the system. Photoactive NPs such as nTiO2 and nZnO, can also

induce the formation of ROS. When exposed to visible or UV light, these NPs can be photo excited resulting

in the formation of electron-holes, which are powerful oxidants that can react with surface bounded

molecules forming radicals (Clemente et al., 2014). All these processes are schematized in Figure P1. 3.

Figure P1. 3. Potential routes for the generation of ROS due to the presence of NPs. 1) Internalization of NPs ROS generation could occur due to the NPs dissolution inside the cells and/or due to the NPs photocatalytic activity. 2) Dissolution of the NPs leads to an increase concentration of metal ions in the media; some of these metals can also be uptake by the organisms. 3) NPs and/or their surrounding coatings can adsorb/complex other metals present in the media, being taken up by the cells. 4) Photocatalytic activity of the NPs in the presence of UV and/or natural light.

3.2. Omics endpoints

The omics tools, such as toxicogenomic and proteomic, are very useful on the establishment of toxic

endpoints. A toxicogenomic approach allows the identification of gene and protein activities in organisms’

cells induced when in the presence of a certain xenobiotic. A central assumption is that chemicals generating

toxicity by the same mechanism will produce similar gene expression responses under a given set of

conditions, bringing new insights about their mode of action that can be linked to their specific

physicochemical properties. A metallomic and proteomic approach will allow a complete analysis on the

metal and metalloid species composition within a cell or tissue and the establishment of metallo-proteins

profiles leading to the identification of new biomarkers (e.g., proteins expressed by the NP itself (Shepard et


39

al., 2000). This will allow a better understanding and profiling of NPs toxic mechanisms discriminating them

from their bulk components.

Despite the large number of nanotoxicology studies, few of them have reported the use of omics tools to

evaluate NPs toxic effects at the molecular level on freshwater organisms (Kuznetsova et al., 2014; Rainville

et al., 2014; Revel et al., 2017). The “omics” approach generates a huge amount of data whose interpretation

is not always straightforward. The large amount of data together with unappropriated physico-chemical

characterization prior to the biological assays results in an escalating number of unknown variables impeding

a comprehensible understanding of the biochemical responses.

Despite the lack of genetic information needed for applying proteomic techniques to freshwater organisms

(Larkin et al., 2003), these approaches are interesting in nanotoxicology as they allow a better understanding

and profiling of toxic mechanisms of nanoparticles and discriminate them from their bulk components

(Rainville et al., 2014). In addition, it has been proved that, to use biomarker measurements, it is better to

have a deep knowledge of mode of actions of pollutants (Vioque-Fernández et al., 2009) and to avoid the

influence of confounding factors such as gender (Sornom et al., 2010), parasitism (Gismondi et al., 2012a,

2012b) and seasonal variations (Sroda and Cossu-Leguille, 2011). Therefore, the use of proteomic approaches

can allow the identification of unbiased responses for nanoparticle exposures (Shepard et al., 2000).

Even though the number of studies concerning nanoparticles-induced toxicity in aquatic organisms under

laboratory conditions continues to increase, the mode of action behind NPs toxicity in freshwater organisms

need further clarification (Canesi et al., 2017; Moore, 2006; Scown et al., 2010). Some studies on metallic

nanoparticles reported, for instance, the effects of copper nanoparticles on the proteomes of aquatic

organisms by considering the effects of CuNPs and Cu2+ on two marine bivalves M. galloprovincialis (Gomes

et al., 2014) and M. edulis (Hu et al., 2014). M. galloprovincialis were exposed for 15 days to CuNPs and Cu2+

(10 μg. L-1). CuNPs showed a high tendency to up-regulate proteins in the gills tissues and down-regulate

them in the digestive gland, while Cu2+ showed the opposite tendency. The effects were metal-dependent

with either common or distinctive response mechanisms induced by CuNPs and Cu2+. Both CuNPs and Cu2+

affected cytoskeleton and cell structure (actin, α-tubulin, paramyosin), stress response (heat shock cognate

71, putative c1q domain containing protein), transcription regulation (zinc finger BED domain-containing

protein 1, nuclear receptor subfamily 1G), and energy metabolism (ATP synthase FO subunit 6). CuNPs

specific mechanisms were also identified with oxidative stress (GST), proteolysis (Cathepsin L) and apoptosis

(caspase3/7-1) while Cu2+ impacted percollagen-D, an adhesion and mobility protein associated with the

detoxification mechanism of Cu2+. This study demonstrated that CuNPs effects are not only due to ion release

but are also due to mitochondrial and nucleus stress induced cell-signalling cascade that can lead to apoptosis

(Gomes et al., 2012, 2013a). M. edulis were exposed to 100 nm CuNPs (1000 ppb) for 1h. CuNPs accumulated


40

in gills and caused an alteration of the expression of six proteins: α and β-tubulin, actin, tropomyosin,

triosephosphate isomerase and CuZnSOD. Actin and triosephosphate showed a decrease in proteins thiol

levels while alpha-tubulin, tropomyosin and CuZnSOD showed an increase in carbonylation indicating protein

oxidation of cytoskeleton and antioxidant enzymes in response to CuNPs. This study showed that CuNPs

affect first cytoskeletal protein and disrupt lysosomal membrane stability (Hu et al., 2014). Other studies

reported the effects of ZnO NPs at the transcriptomic level of D. magna (Poynton et al., 2012). Exposure of

adult D. magna to very high concentration (9 mg.L-1) of 20 nm ZnO-NPs for 24h and to 0.52 mg.L-1 of 30 nm

of ZnO-NPs for 96h led to induction of cytoskeletal transport proteins and a repression of the expression of

genes linked to the reproduction and antioxidant responses (Adam et al., 2015; Poynton et al., 2012). Within

metallic NPs, TiO2 NPs are among the most studied ones. Molecular effects of TiO2 NPs on zebrafish were

reported. TiO2 NPs 20 nm were showed to cause a down-regulation of genes involved in ribosomal functions

of D. rerio gills (Griffitt et al., 2009). Other studies reported that microinjection of TiO2 NPs (25 nm; anatase;

8.5 ng.g-1 bw; 48h at 28.5°C) in zebrafish embryo significantly modified the expression of genes involved in

circadian rhythm, cell signalling, exocytosis and vesicular trafficking (Jovanović et al., 2011). However, no

induction of ROS induction was detected in these two studies.

4. NPS TOXICITY ON FRESHWATER ORGANISMS

4.1. Silver nanopartic les (AgNPs)

AgNPs are known for their antifungal and antimicrobial properties, being extensively used in several products

such as clothing, cosmetics, medical devices, paints, etc. (Fabrega et al., 2011; McGillicuddy et al., 2017;

Vance et al., 2015). Several databases have been created in order to collect information of nano-

functionalised products. Two reviews compiled lists of these inventories (Hansen et al., 2016; Vance et al.,

2015). According to the Woodrow Wilson database, more than 400 consumer products contain AgNPs which

make them among the most used NPs in consumer products (Hansen et al., 2016; Vance et al., 2015). The

global annual production of silver has been estimated around 27,000 tons (US Geological Survey, 2016) with

an annual consumption of 55 tons/year to 450 tons/year as AgNPs (Zhang et al., 2016). The increasing

application of AgNPs will lead to their release in the environment through different routes from synthesis,

manufacturing, distribution, end product use and end of life disposal of every day consumer products

containing AgNPs (Blaser et al., 2008; Mueller and Nowack, 2008). Release of AgNPs in the environment may

occur via wastewater treatment plants (WWTP) even though AgNPs may be retained in sewage sludge (Kaegi

et al., 2013). In wastewater, it was suggested that AgNPs are in contact with sulfides leading to the formation

of an important amount of silver sulfide (Ag2S) that are kept in sewage sludge (Kaegi et al., 2013; Völker et

al., 2015) leading to the release of about 15% of AgNPs in surface water (Keller et al., 2013). Thus, the


41

predicted environmental concentrations in surface water are usually in the low ng. L-1 to µg. L-1 (Mueller and

Nowack, 2008) and range from 0.088 to 2.16 ng. L-1 in European and North American surface waters

(Gottschalk et al., 2009). Recent studies reported predicted environmental concentrations, in surface waters,

in the range from 0.01 and 0.32 µg. L-1 (Tiede et al., 2009; Yang et al., 2016).

Many questions have been raised in order to determine the fate of AgNPs once they reach the environment

(Klaine et al., 2008). Several factors including size, structure and environmental physico-chemical parameters

may affect the behaviour of AgNPs in aquatic environment and thus influence their toxic effects. The

speciation and concomitant interference is of particular note to researchers attempting to risk assess AgNPs

as their toxic mechanisms towards aquatic organisms are still not fully clear (McGillicuddy et al., 2017). It was

possible to identify two distinct routes that could induce biochemical responses: (Auffan et al., 2009; Lowry

et al., 2012; Schultz et al., 2014):

• Presence of the NPs per se

• Presence of both AgNPs and dissolved Ag+.

The identification of responses that are uniquely due to the presence of AgNPs and do not occur in matched

Ag+ exposures are crucial. This is possible by:

• Using AgNPs with low dissolution rates, so, that the leached Ag+ in the media is insufficient to induce

toxicity to the organisms,

• Identify endpoints specific to AgNPs, such as internalization of NPs, cytotoxicity and genotoxicity.

The work performed by Kumar et al., (2014) is an example of the application of these strategies. They reported

a significant increase of ROS and SOD in addition to morphological alterations on freshwater bacteria exposed

to polyvinyl-pyrrolidone PVP-AgNPs. Since the NPs dissolution was very low (leached Ag+ < 1 µg. L-1), it was

concluded that the results were related to AgNPs form. Morphological changes and alteration in genes

profiles, related to the presence of AgNPs itself, were also observed in carp (Cyprinus carpio) (Lee et al., 2012),

zebrafish (Danio rerio) (Choi et al., 2010; Griffitt et al., 2009), D. magna (Poynton et al., 2012), medaka

(Oryzias latipes) (Pham et al., 2012) and rainbow trout (Oncorhynchus mykiss) (Gagné et al., 2012). The

exposure of 7 days old D. magna to citrate-coated AgNPs (size 10 nm, 30 μg L-1) and AgNO3 (2.5 μg L-1) during

24h showed that both AgNPs and Ag+ increased proteins thiol content, while only particles increased proteins

carbonyl levels (Gündel et al., 2007). Similarly, to D. magna, AgNPs and Ag+ also impacted different proteins

in the mussel M. gallopronvinciallis. Although M. galloprovincialis is a marine organism, the observed effects

on this species exposed to AgNPs was highly significant. M. galloprovincialis were exposed for 15 days to

AgNPs and Ag+ (10 μg. L-1). AgNPs affected similar cellular pathways than Ag+ with common responses

mechanisms in cytoskeleton and cell structure (catchin, myosin heavy chain), stress responses (heat shock

protein 70), oxidative stress (GST), transcriptional regulation (nuclear receptor subfamily 1G), adhesion and


42

mobility (percollagen P), and energy metabolism (ATP synthase FO subunit 6 and NADH dehydrogenase

subunit 2). Exposure to AgNPs altered the expression of protein associated with stress response (major vault

protein and ras partial) and proteins involved in cytoskeleton and cell structure (paramyosin) while Ag+ had

a strong effect in proteins related to stress response (putative c1q domain containing protein) and proteins

involved in cytoskeleton and cell structure (actin and a-tubulin). The identification of altered protein

suggested that AgNPs toxicity is mediated by oxidative stress-induced cell signalling cascade (including

mitochondria and nucleus) that can lead to cell death. In fact, it has been described that oxidative stress

caused by AgNPs and Ag+ induce cytoskeleton disorganization (Gomes et al., 2013b). Transcriptomic effects

of two kinds of AgNPs (CIT-AgNPs 40 nm and PVP-AgNPs 35 nm) were investigated on 10 days old D. magna

after 24 h of exposure to different concentrations corresponding to 1/10 of the LC50 of each AgNPs (CIT-

AgNPs: 0.43 µg.L-1; PVP-Ag NPs: 1.1 µg.L-1) and also at the LC25 (CIT-AgNPs: 3.5 µg.L-1; PVP-AgNPs: 8.1 µg.L-1).

This work revealed common effects of these two types of AgNPs with an induction of the ubiquitination and

proteolysis pathways and a repression of ribosomal genes suggesting a general effect on the protein

metabolism. This work also revealed an effect of the PVP coated AgNPs on the expression of immune and

circadian genes of D. magna (Poynton et al., 2012). Other studies reported a significant decrease in lysosomal

membrane stability and the presence of vacuolization and necrosis in D. rerio exposed through the diet to

100 µg.L-1 of PVP/PEI-AgNPs 5 nm was reported (Lacave et al., 2017). Additionally, DNA damage, total

degeneration of hepatocytes and down regulation of stress related genes in L. rohita exposed to PVP-AgNPs

20 and 30 nm were observed. These studies suggested that the observed effects were caused by the AgNP

itself.

It is well known that Ag+ has a great propensity to bioaccumulate in tissues leading to ROS generation,

genotoxicity and inhibition of Na+/ K+-ATPase activity by blocking the Na+ uptake by the cells (Arce Funck et

al., 2013; Luoma et al., 2014; Morgan et al., 1997). For instance, toxic effects were observed in juvenile

Oncorhynchus mykiss exposed to CIT-AgNPs, with claimed low propensity for dissolution (Schultz et al., 2012).

However, it was not possible to confidently establish if the observed effects were caused by the AgNPs per

se or by dissolved Ag+. In fact, it is very difficult to distinguish between the toxic effects induced by particulate

or ionic Ag, and, thus, is crucial not only to evaluate the size distribution but also to quantify the AgNPs

dissolution in the medium.

Both forms, NPs and ionic Ag, can induce OS and genotoxicity being the distinction between these effects a

truly challenge. For example, in the algae Chlorella vulgaris a positive correlation between ROS production

and LPO on the tissues after 24h of exposure to uncoated AgNPs (1 and 10 mg L-1) (Oukarroum et al., 2012)

was found, but with no possibility to establish a positive correlation with solely the particulate or ionic Ag.

Similar difficulties were also observed in the snail Lymnaea luteola (Ali et al., 2014), as exposure to AgNPs


43

resulted in DNA damage and induction of OS but without the possibility to identify which Ag forms has the

dominant role on the observed effects. In these situations, a complete physicochemical characterization of

the NP in the exposure media, localisation within biological tissues along with the use of controls containing

ionic Ag are crucial to understand which Ag form (or both) is responsible for the observed biochemical

responses. Ions release form AgNPs can directly bind to cell structure or interact with sulfhydryl groups of

proteins (such as Ca2+-ATPase) and disrupt calcium homeostasis through ROS formation (AshaRani et al.,

2009; AshaRani et al., 2009; Gomes et al., 2013b). Mechanical injury caused by AgNPs in mitochondria may

be another reason of important changes in membrane permeability, disruption of the respiratory chain and

contributes to oxidative stress (Asharani et al., 2009b). The up-regulation on the nuclear receptor sub-family

1G in the gills were reported after exposure to Ag+ indicates the ability of silver ions (released from AgNPs)

to interfere with signal transduction in DNA-related functions and induce genotoxicity (Gomes et al., 2013b).

Localisation of NPs in biological tissues is a step forward in understanding uptake route and internal

distribution. For instance, PVP-AgNPs 50 nm were observed in the brain of the fish Piaractus mesopotamicus

(Bacchetta et al., 2017) and poly-N-vynil-2-pirrolidone/polyethylenimine (PVP/PEI) AgNPs 5 nm were found

in liver and intestine of D. rerio (Lacave et al., 2017) as CIT-AgNPs 20 nm were observed in liver and gills of

rainbow trout (Bruneau et al., 2016). Other studies reported a dose and time-dependent uptake of PVP-

AgNPs 20 and 30 nm in the fish L. rohita exposed to up to 800 µg.L-1 for 168h (Sharma et al., 2016). Usually,

localisation of NPs is performed using transmission electron microscopy (Heinlaan et al., 2011; Lovern et al.,

2008). This method allows the detection of NPs without identifying the chemical nature of the observed

element. Thus, more sensitive techniques have been used in order to better define AgNPs internal

distribution in aquatic organisms. For example, a secondary ion mass spectrometry instruments, NanoSIMS

50, was used to define AgNPs internal distribution in D. magna exposed for 48h to up to 10 mg.L-1 of

chemically synthesised AgNPs 20 and 200 nm and biologically synthesised from plant leaf extract AgNPs 23

and 27nm (Georgantzopoulou et al., 2013). AgNPs were observed as big aggregates in the gut lumen of D.

magna with AgNPs 20 nm being able to cross the gastrointestinal tract (GIT), AgNPs 200 nm passing the

peritrophic membrane and possibly being present within the epithelial layer of the cells and AgNPs 23 nm

being observed around developing oocytes. However, AgNPs 27 nm were not found to have cross D. magna’s

GIT (Georgantzopoulou et al., 2013). These results were linked to surface properties of the studied AgNPs as

the biological AgNPs 23 and 27 nm were synthesised from to different plants, Ocinum sanctum and

Azadirachta indica, respectively. AgNPs 23 nm appeared as the one to release the highest amount of Ag ions.

Thus, although NanoSIMS 50 allowed the internal distribution of AgNPs in D. magna it did not allow

distinction between the AgNPs per se and the released ions.


44

4.2. Gold nanoparticles (AuNPs)

Compared to AgNPs, AuNPs are considered as chemically stable NPs as they are not expected to release ions

(Auffan et al., 2012). AuNPs are being investigated for their unique optical properties and are used as contrast

agents in electron microscopy, optical sensors, catalysts, and for therapeutic uses. AuNPs have one of the

broadest range of synthetic strategies (Thompson et al., 2017) resulting in cubes, spheres, pyramids, stars

octopods and other variants (Xia et al., 2008). Au metal is extremely resistant to oxidation and essentially

insoluble under ambient conditions. Thus, using AgNPs and AuNPs as model NPs in the present work allowed

us to integrate the effects of ions and NPs on one hand and effects of NPs only on the other hand.

A major challenge in nano-ecotoxicology is finding suitable methods to determine the uptake and localisation

of NPs on a whole organism level (Skjolding et al., 2017). For nanoparticles in order to cause toxicity, it is

assumed that they are taken up and distributed within an organism, therefore a better understanding of how

they behave in an in vivo system and what factors influence their uptake by living organisms is necessary

(Wepener et al., 2011). Some studies reported a size and coating dependent bioaccumulation of AuNPs. CIT-

AuNPs and mercaptoundecanoic acid (MUDA)-AuNPs 10 and 30 nm were reported to accumulate in Daphnia

magna with CIT-AgNPs being more taken up then MUDA-AuNPs and MUDA-AuNPs 10 nm being more

bioconcentrated than MUDA-AuNPs 30 nm (Skjolding et al., 2014). One aspect on NPs structure that may

influence toxicity is the identity and charge of ligand molecules used for functionalization of NPs surface.

Thus, three different types of capping agent of 4-5 nm AuNPs were tested on D. magna. Animals were

exposed to CIT-AuNPs, MPA-AuNPs and PAH-AuNPs, and acute and chronic toxicity tests were performed.

The negatively charged AuNPs appeared to be less toxic while after chronic exposure, both positively and

negatively charged AuNPs affected reproduction (Bozich et al., 2014). However, the mechanisms underlying

the observed effects could not be clearly determined even though the surface chemistry appeared as a critical

factor influencing NPs toxicity on D. magna (Bozich et al., 2014). Uptake of different sizes and surface-coated

AuNPs was modelled using D. magna as model organism (Wray and Klaine, 2015). Size and surface charges

resulted as the main influencing factors controlling AuNPs uptake by D. magna and elimination, whereas

shape had no significant effects (Wray and Klaine, 2015). Examination of intestinal microvilli indicate no

assimilation of AuNPs by D. magna as AuNPs remained in the gut lumen and the carapace (Wray and Klaine,

2015). One study used an innovative technique that allows the localisation of NPs but not of ions to localise

and determine the internal distribution of AuNPs in D. magna. Animals were exposed for 14 days to up to 20

mg. L-1 of AuNPs. Cytoviva darkfield imaging microscope was used to localise AuNPs in D. magna. AuNP

aggregates were observed in the gut of D. magna when animals were fed as no AuNPs were observed in the

body cavity. However, a direct exposure to AuNPs led to the adsorption of AuNPs on the carapace of D.


45

magna and led also to their ingestion and uptake into the gut. This significant uptake had no significant effect

on reproduction and moulting pattern of D. magna (Botha et al., 2016).

Few other studies investigated the effects of AuNPs on freshwater invertebrates. For instance, asiatic clams

Corbicula fulminea were exposed to algae contaminated with 10 nm of amino-acid AuNPs for 7 days. A

concentration-dependent mortality of algae was observed after 24h of exposure. Trophic exposure led to a

significant bioaccumulation with amino-acid AuNPs were observed in the gills and digestive gland. AuNPs

were also observed in the lysosome and led to an oxidative stress in C. fulminea (Renault et al., 2008). Very

few studies have investigated the effects of AuNPs on Gammarus sp. G. pulex were exposed to differently

functionalized AuNPs, namely CIT-AuNPs, MUDA-AuNPs, NH2-AuNPs and PEG-AuNPs. A surface-coating-

dependent uptake of AuNPs was observed in G. pulex with CIT-AuNPs and MUDA-AuNPs being the most

bioaccumulated by G. pulex while no size effects were observed (Park et al., 2015). To our best knowledge

there is only one study investigating effects of AuNPs on G. fossarum. Individuals were exposed through a

contaminated biofilm to up to 45 mg.L-1 of 10 nm amino-acid AuNPs (Baudrimont et al., 2017). Cellular

damage linked to an oxidative stress and significant effect on mitochondrial respiration was observed after 7

days of exposure with a daily renewal of contaminated food. Additionally, modulation of digestive enzymes

was also observed suggestion a modification of digestive functions (Baudrimont et al., 2017).

Within freshwater vertebrates, the model organism zebrafish is one of the most studied species. Zebrafish

embryo were exposed to different metal-based nanoparticles (Ag, Au, CuO and ZnO). AuNPs appeared as the

most bioconcentrated ones in the eggs followed by Ag, ZnO and CuO. However, when the whole organism

was assessed, Ag appeared as the highest bioaccumulated NPs followed by Zn (Böhme et al., 2017). A

previous study also reported presence of AuNPs in ovaries of female zebrafish exposed to AuNPs 15 nm and

47 nm (Dayal et al., 2016). Zebrafish was exposed to fluorescent AuNPs via aqueous or dietary route (Skjolding

et al., 2017). Internal distribution was assessed using light sheet microscopy (LSM) at different time point (1,

3 and 7 days). AuNPs were observed within the gut of zebrafish after trophic exposure. AuNPs were not

observed in the gut epithelia indicating the absence or limited uptake of AuNPs through the intestinal villi.

Direct exposure of zebrafish to AuNPs led an increase in relative swimming distance. This study showed the

importence of exposure routes and subsequent localisation of AuNPs in zebrafish (Skjolding et al., 2017).

Deleterious effects were reported as AuNPs caused modulation of genes linked to oxidative stress,

mitochondrial metabolism, detoxification and DNA repair in zebrafish exposed to AuNPs 14 nm (Dedeh et al.,

2015). Other studies using estuarine and marine organisms reported further toxic effects of AuNPs. Estuarine

copepod Eurytemora affinis were exposed for 30 min to 11.4 µg. L-1 of AuNPs and a decrease in swimming

behaviour and a lower velocity and acceleration were observed in females whereas males appeared to be

less sensitive. However, the mechanisms underlying these effects were not clearly determined as neither


46

adsorbed nor ingested AuNPs could be localised in E. affinis (Michalec et al., 2017). AuNPs showed a higher

uptake in the digestive gland than in the gills of the clam R. philippinarum. Changes in oxidative stress and

inflammatory response markers as phase II antioxidant enzymes and qPCR changes were observed (Volland

et al., 2015). However, no strong oxidative damage was reported and a significant depuration from digestive

tract after 7 days in cleans medium was observed (Volland et al., 2015). The oligochaetes Tubifex tubifex were

exposed for 5 days to 10 and 60 µg. g-1 dry weight of sediments of 5 nm AuNPs. A concentration dependent

uptake of Au was observed but the internal concentration did not cause significant mortality of T. tubifex

(Zhang et al., 2017). Marine blue mussels Mytillus edulis were exposed to 750 ppb of 5 nm AuNPs for 24h

which mainly accumulated in the digestive gland. A significant decrease in lysosomal membrane stability in

addition to a significant increase in lipid peroxidation was observed together with a decrease in thiol-

containing proteins. All these results suggest that the observed effects are linked to an increase in ROS leading

to oxidative stress (Tedesco et al., 2010).

5. GENERAL REMARKS AND CONCLUSIONS

Most of the biochemical responses reported are related to the organism’s ROS defence mechanisms, mainly

through gene expression or changes on anti-oxidant enzyme activities. As mentioned before (section 3.2),

data related with changes in freshwater organism genome and proteome due to the presence of NPs is very

scarce. This is indeed surprisingly since -omic techniques have already proved a great potential on the

recognition of signatures related to specific stress, eventually leading to the discover of new biomarkers

(Revel et al., 2017). The few data available suggest that the interaction with the organisms is NP-specific

regarding the diversity of effects of AgNPs and AuNPs on aquatic organisms and raise questions about clear

and defined mechanism of action.

NPs composition can play an important role on ROS generation, since some metals constituting the NP can

instigate Fenton and Weiss-type reactions releasing ROS in the intra or extracellular media. Evidently, and as

usually performed, the bioassays should contain a control group exposed to the salt form of the metal

constituting the NP, allowing the distinction between the effects provoked by the NP per se and/or by the NP

dissolution products.

Despite the large number of studies dealing with ecotoxicology of NPs, it is evident that is still not possible to

establish crucial predictive structure-activity relationships. This is mainly due to the fact that the majority of

the available studies have critical deficiencies on their experimental designs; a comprehensive physico-

chemical characterization of the particles under the exposure conditions is mostly miscarried or restricted to

a secondary task. Despite the scientific community is already aware about the importance of the NPs

physicochemical characterization prior and during the bioassay, this is still frequently neglected in the most

recent studies giving rise to more confusing and contradictory data. Clearly, this pushes back the possibility


47

to establish a proper environmental risk assessment plan for these current early generations on ENPs (1st and

2nd generation passive and active nanostructures, respectively), while advanced generations of ENPs (3rd and

4th generation nano-systems and molecular nano-systems) may not be far away, bringing additional

challenges that require further novel approaches.

The dynamic speciation of the NPs should be assessed in the same exposure media of the bioassay by

following key NPs transformations:

• Dissolution,

• Homo- and hetero-aggregation,

• Sedimentation.

Several analytical tools are nowadays available for the quantification of these physicochemical

transformations, each of them having their specific advantages and limitations being able to provide different

information on ENPs properties (see reviews (Domingos et al., 2009; Tiede et al., 2009). Several analytical

tools are nowadays available for the quantification of these physicochemical transformations, each of them

having their specific advantages and limitations being able to provide different information on ENPs

properties (see reviews Domingos et al., 2009; Tiede et al., 2009). This physicochemical characterization

approach allows:

• To assess the bioavailable NP-containing species to which the organisms will be exposed,

• To relate the biocompatible or bioadverse effects with the NP-containing species permitting a NP

categorization and function.

Nanotoxicology is indeed a multidisciplinary field where the study of the NPs physic, chemistry and biological

impacts is crucial for a complete toxicological assessment. Unfortunately, there is a lack of legislation

controlling the production, use and release of these materials to the environment, and new NPs are

commercialized every day without an appropriate assessment about their impacts in environment and

human health. The establishment of national and international laws regulating the production of these

materials is mandatory. Furthermore, it is also urgent to increase the number of comprehensive

nano(eco)toxicology studies under natural more environmentally-realistic conditions implying the co-

presence of ENPs (at low and environmentally-realistic doses) and environmental constituents such as natural

organic and inorganic dissolved and colloidal matter. Only with these approaches a comprehensive risk

assessment will be possible with production of environmentally safe-by-design ENPs.


48

Acknowledgements

All the authors acknowledge to the European COST action ES1205, Engineered Nanomaterials - From

Wastewater Treatment & Stormwater to Rivers (ENTER). G. Vale and R.F. Domingos acknowledges to

Fundação para a Ciência e Tecnologia (FCT, Portugal) the post - doctoral grant SFRH/BPD/73117/2010 and

the project PTDC/AAC-AMB/111998/2009, respectively. The contribution of K. Mehennaoui and S. Cambier

were in part possible within NanoGAM (AFP-PhD-9229040) and NANION (FNR/12/SR/4009651) respectively,

both founded by Fonds National de la Recherche Luxembourg.

REFERENCE

Adam, N., Vergauwen, L., Blust, R., Knapen, D., 2015. Gene transcription patterns and energy reserves in Daphnia

magna show no nanoparticle specific toxicity when exposed to ZnO and CuO nanoparticles. Environ. Res. 138, 82–

92. doi:10.1016/j.envres.2015.02.014

Ali, D., Yadav, P.G., Kumar, S., Ali, H., Alarifi, S., Harrath, A.H., 2014. Sensitivity of freshwater pulmonate snail

Lymnaea luteola L., to silver nanoparticles. Chemosphere. doi:10.1016/j.chemosphere.2013.10.081

Arce Funck, J., Danger, M., Gismondi, E., Cossu-Leguille, C., Guérold, F., Felten, V., 2013. Behavioural and

physiological responses of Gammarus fossarum (Crustacea Amphipoda) exposed to silver. Aquat. Toxicol. 142–

143, 73–84. doi:10.1016/j.aquatox.2013.07.012

AshaRani, P., Hande, M.P., Valiyaveettil, S., 2009. Anti-proliferative activity of silver nanoparticles. BMC Cell Biol.

10, 65. doi:10.1186/1471-2121-10-65

AshaRani, P.V., Low Kah Mun, G., Hande, M.P., Valiyaveettil, S., 2009. Cytotoxicity and Genotoxicity of Silver

Nanoparticles in Human Cells. ACS Nano 3, 279–290. doi:10.1021/nn800596w

Auffan, D.M., Santaella, D.C., Thiéry, P.A., Paillès, C., Rose, J., Achouak, D.W., Thill, D.A., Masion, A., Wiesner, M.,

Bottero, J.-Y., 2012. Ecotoxicity of Inorganic Nanoparticles: From Unicellular Organisms to Invertebrates, in:

Bhushan, P.B. (Ed.), Encyclopedia of Nanotechnology. Springer Netherlands, pp. 623–636.

Auffan, M., Rose, J., Wiesner, M., Bottero, J.-Y., 2009. Chemical stability of metallic nanoparticles : a parameter

controlling their potential cellular toxicity in vitro.

Bacchetta, C., Ale, A., Simoniello, M.F., Gervasio, S., Davico, C., Rossi, A.S., Desimone, M.F., Poletta, G., López, G.,

Monserrat, J.M., Cazenave, J., 2017. Genotoxicity and oxidative stress in fish after a short-term exposure to silver

nanoparticles. Ecol. Indic. 76, 230–239. doi:10.1016/j.ecolind.2017.01.018

Barata, C., Varo, I., Navarro, J.C., Arun, S., Porte, C., 2005. Antioxidant enzyme activities and lipid peroxidation in

the freshwater cladoceran Daphnia magna exposed to redox cycling compounds. Comp. Biochem. Physiol. Part C

Toxicol. Pharmacol. 140, 175–186. doi:10.1016/j.cca.2005.01.013

Baudrimont, M., Andrei, J., Mornet, S., Gonzalez, P., Mesmer-Dudons, N., Gourves, P.-Y., Jaffal, A., Dedourge-

Geffard, O., Geffard, A., Geffard, O., Garric, J., Feurtet-Mazel, A., 2017. Trophic transfer and effects of gold

nanoparticles (AuNPs) in Gammarus fossarum from contaminated periphytic biofilm. Environ. Sci. Pollut. Res.

doi:10.1007/s11356-017-8400-3

BCC RESEARCH, 2017. NAN021G_Report Overview.pdf [WWW Document]. URL

https://cdn2.hubspot.net/hubfs/308401/NAN%20Report%20Overviews/NAN021G_Report%20Overview.pdf?t=1

507039495677&utm_campaign=NAN021G&utm_source=hs_automation&utm_medium=email&utm_content=3


49

9598398&_hsenc=p2ANqtz-8aMzzG8s1ejWU5vfx62L2v9e7nraO-

2Zo5IxMvgn1QGf168euhj5LTcihQ1hQNtJhW680aqJqccZGQGLNnRuMjC9fE2g&_hsmi=39598398 (accessed

10.3.17).

Blaser, S.A., Scheringer, M., MacLeod, M., Hungerbühler, K., 2008. Estimation of cumulative aquatic exposure and

risk due to silver: Contribution of nano-functionalized plastics and textiles. Sci. Total Environ. 390, 396–409.

doi:10.1016/j.scitotenv.2007.10.010

Böhme, S., Baccaro, M., Schmidt, M., Potthoff, A., Stärk, H.-J., Reemtsma, T., Kühnel, D., 2017. Metal uptake and

distribution in the zebrafish (Danio rerio) embryo: differences between nanoparticles and metal ions. Environ. Sci.

Nano 4, 1005–1015. doi:10.1039/C6EN00440G

Botha, T.L., Boodhia, K., Wepener, V., 2016. Adsorption, uptake and distribution of gold nanoparticles in Daphnia

magna following long term exposure. Aquat. Toxicol. 170, 104–111. doi:10.1016/j.aquatox.2015.11.022

Bour, A., Mouchet, F., Silvestre, J., Gauthier, L., Pinelli, E., 2015. Environmentally relevant approaches to assess

nanoparticles ecotoxicity: a review. J. Hazard. Mater. 283, 764–777. doi:10.1016/j.jhazmat.2014.10.021

Bowman, C.R., Bailey, F.C., Elrod-Erickson, M., Neigh, A.M., Otter, R.R., 2012. Effects of silver nanoparticles on

zebrafish (Danio rerio) and Escherichia coli (ATCC 25922): a comparison of toxicity based on total surface area

versus mass concentration of particles in a model eukaryotic and prokaryotic system. Environ. Toxicol. Chem. 31,

1793–1800. doi:10.1002/etc.1881

Bozich, J.S., Lohse, S.E., Torelli, M.D., Murphy, C.J., Hamers, R.J., Klaper, R.D., 2014. Surface chemistry, charge and

ligand type impact the toxicity of gold nanoparticles to Daphnia magna. Environ. Sci. Nano 1, 260–270.

doi:10.1039/C4EN00006D

Brigelius-Flohé, R., Maiorino, M., 2013. Glutathione peroxidases. Biochim. Biophys. Acta BBA - Gen. Subj., Cellular

functions of glutathione 1830, 3289–3303. doi:10.1016/j.bbagen.2012.11.020

Bruneau, A., Turcotte, P., Pilote, M., Gagné, F., Gagnon, C., 2016. Fate of silver nanoparticles in wastewater and

immunotoxic effects on rainbow trout. Aquat. Toxicol. 174, 70–81. doi:10.1016/j.aquatox.2016.02.013

Burić, P., Jakšić, Ž., Štajner, L., Dutour Sikirić, M., Jurašin, D., Cascio, C., Calzolai, L., Lyons, D.M., 2015. Effect of

silver nanoparticles on Mediterranean sea urchin embryonal development is species specific and depends on

moment of first exposure. Mar. Environ. Res., Particles in the Oceans: Implication for a safe marine environment

111, 50–59. doi:10.1016/j.marenvres.2015.06.015

Canesi, L., Balbi, T., Fabbri, R., Salis, A., Damonte, G., Volland, M., Blasco, J., 2017. Biomolecular coronas in

invertebrate species: Implications in the environmental impact of nanoparticles. NanoImpact.

doi:10.1016/j.impact.2017.08.001

Choi, J.E., Kim, S., Ahn, J.H., Youn, P., Kang, J.S., Park, K., Yi, J., Ryu, D.-Y., 2010. Induction of oxidative stress and

apoptosis by silver nanoparticles in the liver of adult zebrafish. Aquat. Toxicol. 100, 151–159.

doi:10.1016/j.aquatox.2009.12.012

Clemente, Z., Castro, V.L.S.S., Moura, M.A.M., Jonsson, C.M., Fraceto, L.F., 2014. Toxicity assessment of TiO2

nanoparticles in zebrafish embryos under different exposure conditions. Aquat. Toxicol. 147, 129–139.

doi:10.1016/j.aquatox.2013.12.024

Dale, A.L., Lowry, G.V., Casman, E.A., 2013. Modeling Nanosilver Transformations in Freshwater Sediments.

Environ. Sci. Technol. 47, 12920–12928. doi:10.1021/es402341t

Dayal, N., Thakur, M., Patil, P., Singh, D., Vanage, G., Joshi, D.S., 2016. Histological and genotoxic evaluation of gold


50

nanoparticles in ovarian cells of zebrafish (Danio rerio). J. Nanoparticle Res. 18, 291. doi:10.1007/s11051-016-

3549-0

Dedeh, A., Ciutat, A., Treguer-Delapierre, M., Bourdineaud, J.-P., 2015. Impact of gold nanoparticles on zebrafish

exposed to a spiked sediment. Nanotoxicology 9, 71–80. doi:10.3109/17435390.2014.889238

Deponte, M., 2013. Glutathione catalysis and the reaction mechanisms of glutathione-dependent enzymes.

Biochim. Biophys. Acta BBA - Gen. Subj., Cellular functions of glutathione 1830, 3217–3266.

doi:10.1016/j.bbagen.2012.09.018

Domingos, R.F., Baalousha, M.A., Ju-Nam, Y., Reid, M.M., Tufenkji, N., Lead, J.R., Leppard, G.G., Wilkinson, K.J.,

2009. Characterizing Manufactured Nanoparticles in the Environment: Multimethod Determination of Particle

Sizes. Environ. Sci. Technol. 43, 7277–7284. doi:10.1021/es900249m

Domingos, R.F., Franco, C., Pinheiro, J.P., 2015a. The role of charged polymer coatings of nanoparticles on the

speciation and fate of metal ions in the environment. Environ. Sci. Pollut. Res. Int. 22, 2900–2906.

doi:10.1007/s11356-014-3546-8

Domingos, R.F., Gélabert, A., Carreira, S., Cordeiro, A., Sivry, Y., Benedetti, M.F., 2015b. Metals in the Aquatic

Environment—Interactions and Implications for the Speciation and Bioavailability: A Critical Overview. Aquat.

Geochem. 21, 231–257. doi:10.1007/s10498-014-9251-x

Domingos, R.F., Peyrot, C., Wilkinson, K.J., 2010. Aggregation of titanium dioxide nanoparticles: role of calcium

and phosphate. Environ. Chem.

Domingos, R.F., Rafiei, Z., Monteiro, C.E., Khan, M.A.K., Wilkinson, K.J., 2013. Agglomeration and dissolution of

zinc oxide nanoparticles: role of pH, ionic strength and fulvic acid. Environ. Chem. 10, 306–312.

doi:10.1071/EN12202

Fabrega, Samuel N. Luoma, Charles R. Tyler, Tamara S. Galloway, Jamie R. Lead, 2011. Silver nanoparticles:

Behaviour and effects in the aquatic environment. Environ. Int. 37, 517–531. doi:10.1016/j.envint.2010.10.012

Fu, P.P., Xia, Q., Hwang, H.-M., Ray, P.C., Yu, H., 2014. Mechanisms of nanotoxicity: Generation of reactive oxygen

species. J. Food Drug Anal., Nanomaterials - Toxicology and Medical Applications 22, 64–75.

doi:10.1016/j.jfda.2014.01.005

Gagné, F., André, C., Skirrow, R., Gélinas, M., Auclair, J., van Aggelen, G., Turcotte, P., Gagnon, C., 2012. Toxicity of

silver nanoparticles to rainbow trout: A toxicogenomic approach. Chemosphere 89, 615–622.

doi:10.1016/j.chemosphere.2012.05.063

Georgantzopoulou, A., Balachandran, Y.L., Rosenkranz, P., Dusinska, M., Lankoff, A., Wojewodzka, M., Kruszewski,

M., Guignard, C., Audinot, J.-N., Girija, S., Hoffmann, L., Gutleb, A.C., 2013. Ag nanoparticles: size- and surface-

dependent effects on model aquatic organisms and uptake evaluation with NanoSIMS. Nanotoxicology 1–11.

doi:10.3109/17435390.2012.715312

Gismondi, E., Cossu-Leguille, C., Beisel, J.-N., 2012a. Acanthocephalan parasites: help or burden in gammarid

amphipods exposed to cadmium? Ecotoxicology 21, 1188–1193. doi:10.1007/s10646-012-0873-8

Gismondi, E., Rigaud, T., Beisel, J.-N., Cossu-Leguille, C., 2012b. Microsporidia parasites disrupt the responses to

cadmium exposure in a gammarid. Environ. Pollut. 160, 17–23. doi:10.1016/j.envpol.2011.09.021

Gomes, T., Araújo, O., Pereira, R., Almeida, A.C., Cravo, A., Bebianno, M.J., 2013a. Genotoxicity of copper oxide

and silver nanoparticles in the mussel Mytilus galloprovincialis. Mar. Environ. Res. 84, 51–59.

doi:10.1016/j.marenvres.2012.11.009


51

Gomes, T., Chora, S., Pereira, C.G., Cardoso, C., Bebianno, M.J., 2014. Proteomic response of mussels Mytilus

galloprovincialis exposed to CuO NPs and Cu2+: An exploratory biomarker discovery. Aquat. Toxicol. 155, 327–

336. doi:10.1016/j.aquatox.2014.07.015

Gomes, T., Pereira, C.G., Cardoso, C., Bebianno, M.J., 2013b. Differential protein expression in mussels Mytilus

galloprovincialis exposed to nano and ionic Ag. Aquat. Toxicol. 136–137, 79–90.

doi:10.1016/j.aquatox.2013.03.021

Gomes, T., Pereira, C.G., Cardoso, C., Pinheiro, J.P., Cancio, I., Bebianno, M.J., 2012. Accumulation and toxicity of

copper oxide nanoparticles in the digestive gland of Mytilus galloprovincialis. Aquat. Toxicol. 118–119, 72–79.

doi:10.1016/j.aquatox.2012.03.017

Gottschalk, F., Sonderer, T., Scholz, R.W., Nowack, B., 2009. Modeled Environmental Concentrations of Engineered

Nanomaterials (TiO2, ZnO, Ag, CNT, Fullerenes) for Different Regions. Environ. Sci. Technol. 43, 9216–9222.

doi:10.1021/es9015553

Gottschalk, F., Sun, T., Nowack, B., 2013. Environmental concentrations of engineered nanomaterials: Review of

modeling and analytical studies. Environ. Pollut. 181, 287–300. doi:10.1016/j.envpol.2013.06.003

Griffitt, R.J., Hyndman, K., Denslow, N.D., Barber, D.S., 2009. Comparison of Molecular and Histological Changes in

Zebrafish Gills Exposed to Metallic Nanoparticles. Toxicol. Sci. 107, 404–415. doi:10.1093/toxsci/kfn256

Gündel, U., Benndorf, D., von Bergen, M., Altenburger, R., Küster, E., 2007. Vitellogenin cleavage products as

indicators for toxic stress in zebra fish embryos: A proteomic approach. PROTEOMICS 7, 4541–4554.

doi:10.1002/pmic.200700381

Hansen, S.F., Roverskov Heggelund, L., Besora, P.R., Mackevica, A., Boldrin, A., Baun, A., 2016. Nanoproducts –

what is actually available to European consumers? Environ. Sci. Nano 3, 169–180. doi:10.1039/C5EN00182J

Heinlaan, M., Ivask, A., Blinova, I., Dubourguier, H.-C., Kahru, A., 2008. Toxicity of nanosized and bulk ZnO, CuO

and TiO2 to bacteria Vibrio fischeri and crustaceans Daphnia magna and Thamnocephalus platyurus. Chemosphere

71, 1308–1316. doi:10.1016/j.chemosphere.2007.11.047

Heinlaan, M., Kahru, A., Kasemets, K., Arbeille, B., Prensier, G., Dubourguier, H.-C., 2011. Changes in the Daphnia

magna midgut upon ingestion of copper oxide nanoparticles: a transmission electron microscopy study. Water

Res. 45, 179–190. doi:10.1016/j.watres.2010.08.026

Hood, E., 2004. Nanotechnology: Looking As We Leap. Environ. Health Perspect. 112, A740–A749.

Hotze, E.M., Bottero, J.-Y., Wiesner, M.R., 2010. Theoretical framework for nanoparticle reactivity as a function of

aggregation state. Langmuir 26, 11170–11175. doi:10.1021/1a9046963

Hu, W., Culloty, S., Darmody, G., Lynch, S., Davenport, J., Ramirez-Garcia, S., Dawson, K.A., Lynch, I., Blasco, J.,

Sheehan, D., 2014. Toxicity of copper oxide nanoparticles in the blue mussel, Mytilus edulis: A redox proteomic

investigation. Chemosphere 108, 289–299. doi:10.1016/j.chemosphere.2014.01.054

Ivask, A., ElBadawy, A., Kaweeteerawat, C., Boren, D., Fischer, H., Ji, Z., Chang, C.H., Liu, R., Tolaymat, T., Telesca,

D., Zink, J.I., Cohen, Y., Holden, P.A., Godwin, H.A., 2014. Toxicity Mechanisms in Escherichia coli Vary for Silver

Nanoparticles and Differ from Ionic Silver. ACS Nano 8, 374–386. doi:10.1021/nn4044047

Jovanović, B., Ji, T., Palić, D., 2011. Gene expression of zebrafish embryos exposed to titanium dioxide

nanoparticles and hydroxylated fullerenes. Ecotoxicol. Environ. Saf. 74, 1518–1525.

doi:10.1016/j.ecoenv.2011.04.012

Kaegi, R., Voegelin, A., Ort, C., Sinnet, B., Thalmann, B., Krismer, J., Hagendorfer, H., Elumelu, M., Mueller, E., 2013.


52

Fate and transformation of silver nanoparticles in urban wastewater systems. Water Res. 47, 3866–3877.

doi:10.1016/j.watres.2012.11.060

Keller, A.A., McFerran, S., Lazareva, A., Suh, S., 2013. Global life cycle releases of engineered nanomaterials. J.

Nanoparticle Res. 15, 1692. doi:10.1007/s11051-013-1692-4

Keller, A.A., Wang, H., Zhou, D., Lenihan, H.S., Cherr, G., Cardinale, B.J., Miller, R., Ji, Z., 2010. Stability and

Aggregation of Metal Oxide Nanoparticles in Natural Aqueous Matrices. Environ. Sci. Technol. 44, 1962–1967.

doi:10.1021/es902987d

Klaine, S.J., Alvarez, P.J.J., Batley, G.E., Fernandes, T.F., Handy, R.D., Lyon, D.Y., Mahendra, S., McLaughlin, M.J.,

Lead, J.R., 2008. Nanomaterials in the environment: Behavior, fate, bioavailability, and effects. Environ. Toxicol.

Chem. 27, 1825–1851. doi:10.1897/08-090.1

Kumar, D., Kumari, J., Pakrashi, S., Dalai, S., Raichur, A.M., Sastry, T.P., Mandal, A.B., Chandrasekaran, N.,

Mukherjee, A., 2014. Qualitative toxicity assessment of silver nanoparticles on the fresh water bacterial isolates

and consortium at low level of exposure concentration. Ecotoxicol. Environ. Saf. 108, 152–160.

doi:10.1016/j.ecoenv.2014.05.033

Kuznetsova, G.P., Larina, O.V., Petushkova, N.A., Kisrieva, Y.S., Samenkova, N.F., Trifonova, O.P., Karuzina, I.I.,

Ipatova, O.M., Zolotaryov, K.V., Romashova, Y.A., Lisitsa, A.V., 2014. Effects of Fullerene C60 on Proteomic Profile

of Danio Rerio Fish Embryos. Bull. Exp. Biol. Med. 156, 694–698. doi:10.1007/s10517-014-2427-y

Lacave, J.M., Fanjul, Á., Bilbao, E., Gutierrez, N., Barrio, I., Arostegui, I., Cajaraville, M.P., Orbea, A., 2017. Acute

toxicity, bioaccumulation and effects of dietary transfer of silver from brine shrimp exposed to PVP/PEI-coated

silver nanoparticles to zebrafish. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol.

doi:10.1016/j.cbpc.2017.03.008

Larkin, P., Folmar, L.C., Hemmer, M.J., Poston, A.J., Denslow, N.D., 2003. Expression profiling of estrogenic

compounds using a sheepshead minnow cDNA macroarray. EHP Toxicogenomics J. Natl. Inst. Environ. Health Sci.

111, 29–36.

Lee, B., Duong, C.N., Cho, J., Lee, J., Kim, K., Seo, Y., Kim, P., Choi, K., Yoon, J., 2012. Toxicity of Citrate-Capped

Silver Nanoparticles in Common Carp (Cyprinus carpio). J. Biomed. Biotechnol. 2012. doi:10.1155/2012/262670

Levard, C., Hotze, E.M., Lowry, G.V., Brown, G.E., 2012. Environmental Transformations of Silver Nanoparticles:

Impact on Stability and Toxicity. Environ. Sci. Technol. 46, 6900–6914. doi:10.1021/es2037405

Libralato, G., 2014. The case of Artemia spp. in nanoecotoxicology. Mar. Environ. Res. 101, 38–43.

doi:10.1016/j.marenvres.2014.08.002

Lipinski, B., 2011. Hydroxyl Radical and Its Scavengers in Health and Disease [WWW Document]. Oxid. Med. Cell.

Longev. doi:10.1155/2011/809696

Lovern, S.B., Owen, H.A., Klaper, R., 2008. Electron microscopy of gold nanoparticle intake in the gut of Daphnia

magna. Nanotoxicology 2, 43–48. doi:10.1080/17435390801935960

Lowry, G.V., Gregory, K.B., Apte, S.C., Lead, J.R., 2012. Transformations of Nanomaterials in the Environment.

Environ. Sci. Technol. 46, 6893–6899. doi:10.1021/es300839e

Luoma, S.N., Khan, F.R., Croteau, M.-N., 2014. Chapter 5 - Bioavailability and Bioaccumulation of Metal-Based

Engineered Nanomaterials in Aquatic Environments: Concepts and Processes, in: Jamie R. Lead and Eugenia

Valsami-Jones (Ed.), Frontiers of Nanoscience, Nanoscience and the Environment. Elsevier, pp. 157–193.

Ma, H., Williams, P.L., Diamond, S.A., 2013. Ecotoxicity of manufactured ZnO nanoparticles – A review. Environ.


53

Pollut. 172, 76–85. doi:10.1016/j.envpol.2012.08.011

Manke, A., Wang, L., Rojanasakul, Y., 2013. Mechanisms of Nanoparticle-Induced Oxidative Stress and Toxicity

[WWW Document]. BioMed Res. Int. doi:10.1155/2013/942916

Manzo, S., Miglietta, M.L., Rametta, G., Buono, S., Di Francia, G., 2013. Toxic effects of ZnO nanoparticles towards

marine algae Dunaliella tertiolecta. Sci. Total Environ. 445–446, 371–376. doi:10.1016/j.scitotenv.2012.12.051

McGillicuddy, E., Murray, I., Kavanagh, S., Morrison, L., Fogarty, A., Cormican, M., Dockery, P., Prendergast, M.,

Rowan, N., Morris, D., 2017. Silver nanoparticles in the environment: Sources, detection and ecotoxicology. Sci.

Total Environ. 575, 231–246. doi:10.1016/j.scitotenv.2016.10.041

Michalec, F.-G., Holzner, M., Barras, A., Lacoste, A.-S., Brunet, L., Lee, J.-S., Slomianny, C., Boukherroub, R., Souissi,

S., 2017. Short-term exposure to gold nanoparticle suspension impairs swimming behavior in a widespread

calanoid copepod. Environ. Pollut. 228, 102–110. doi:10.1016/j.envpol.2017.04.084

Moore, M.N., 2006. Do nanoparticles present ecotoxicological risks for the health of the aquatic environment?

Environ. Int., Environmental Risk Management - the State of the Art 32, 967–976.

doi:10.1016/j.envint.2006.06.014

Morgan, I.J., Henry, R.P., Wood, C.M., 1997. The mechanism of acute silver nitrate toxicity in freshwater rainbow

trout (Oncorhynchus mykiss) is inhibition of gill Na+ and Cl−1 transport. Aquat. Toxicol. 38, 145–163.

doi:10.1016/S0166-445X(96)00835-1

Mueller, N.C., Nowack, B., 2008. Exposure Modeling of Engineered Nanoparticles in the Environment. Environ. Sci.

Technol. 42, 4447–4453. doi:10.1021/es7029637

Nasser, F., Lynch, I., 2016. Secreted protein eco-corona mediates uptake and impacts of polystyrene nanoparticles

on Daphnia magna. J. Proteomics. doi:10.1016/j.jprot.2015.09.005

Nowack, B., Ranville, J.F., Diamond, S., Gallego-Urrea, J.A., Metcalfe, C., Rose, J., Horne, N., Koelmans, A.A., Klaine,

S.J., 2012. Potential scenarios for nanomaterial release and subsequent alteration in the environment. Environ.

Toxicol. Chem. 31, 50–59. doi:10.1002/etc.726

Oukarroum, A., Bras, S., Perreault, F., Popovic, R., 2012. Inhibitory effects of silver nanoparticles in two green algae,

Chlorella vulgaris and Dunaliella tertiolecta. Ecotoxicol. Environ. Saf. 78, 80–85. doi:10.1016/j.ecoenv.2011.11.012

Park, Woodhall, J., Ma, G., Veinot, J.G.., Boxall, A.B.A., 2015. Do particle size and surface functionality affect uptake

and depuration of gold nanoparticles by aquatic invertebrates? Environ. Toxicol. Chem. 34, 850–859.

doi:10.1002/etc.2868

Pham, C.H., Yi, J., Gu, M.B., 2012. Biomarker gene response in male Medaka (Oryzias latipes) chronically exposed

to silver nanoparticle. Ecotoxicol. Environ. Saf. 78, 239–245. doi:10.1016/j.ecoenv.2011.11.034

Piccinno, F., Gottschalk, F., Seeger, S., Nowack, B., 2012. Industrial production quantities and uses of ten

engineered nanomaterials in Europe and the world. J. Nanoparticle Res. 14, 1–11. doi:10.1007/s11051-012-1109-

9

Poynton, H.C., Lazorchak, J.M., Impellitteri, C.A., Blalock, B.J., Rogers, K., Allen, H.J., Loguinov, A., Heckman, J.L.,

Govindasmawy, S., 2012. Toxicogenomic Responses of Nanotoxicity in Daphnia magna Exposed to Silver Nitrate

and Coated Silver Nanoparticles. Environ. Sci. Technol. 46, 6288–6296. doi:10.1021/es3001618

Rainville, L.-C., Carolan, D., Varela, A.C., Doyle, H., Sheehan, D., 2014. Proteomic evaluation of citrate-coated silver

nanoparticles toxicity in Daphnia magna. Analyst 139, 1678–1686. doi:10.1039/C3AN02160B


54

Rauscher, H., Roebben, G., Boix Sanfeliu, A., Emmons, H., Gibson, N., Koeber, R., Linsinger, T., Rasmussen, K., Riego

Sintes, J., Sokull-Klüttgen, B., Stamm, H., 2015. Towards a review of the EC Recommendation for a definition of the

term “nanomaterial” Part 3 Scientific-technical evaluation of options to clarify the definition and to facilitate its

implementation. doi:10.2788/678452

Regoli, F., Gorbi, S., Frenzilli, G., Nigro, M., Corsi, I., Focardi, S., Winston, G.., 2002. Oxidative stress in

ecotoxicology: from the analysis of individual antioxidants to a more integrated approach. Mar. Environ. Res. 54,

419–423. doi:10.1016/S0141-1136(02)00146-0

Renault, S., Baudrimont, M., Mesmer-Dudons, N., Gonzalez, P., Mornet, S., Brisson, A., 2008. Impacts of gold

nanoparticle exposure on two freshwater species: a phytoplanktonic alga (Scenedesmus subspicatus) and a

benthic bivalve (Corbicula fluminea). Gold Bull. 41, 116–126. doi:10.1007/BF03216589

Revel, M., Châtel, A., Mouneyrac, C., 2017. Omics tools: New challenges in aquatic nanotoxicology? Aquat. Toxicol.

doi:10.1016/j.aquatox.2017.10.005

Savolainen, K., Backman, U., Brouwer, D., Fadeel, B., Fernandes, T., Kuhlbusch, T.A.J., Landsiedel, R., Lynch, I.,

Pylkkanen, L., Ahluwalia, A., Jukka, A., Aitken, R., Alenius, H., Asbach, C., Athersuch, T., Bergamaschi, E., Berges,

M., Bessems, J., Bleeker, E.A.J., Özgüz, V., 2013. Nanosafety in Europe 2015-2025: Towards Safe and Sustainable

Nanomaterials and Nanotechnology Innovations. doi:10.13140/2.1.3084.8969

Schultz, A.G., Boyle, D., Chamot, D., Ong, K.J., Wilkinson, K.J., McGeer, J.C., Sunahara, G., Goss, G.G., 2014. Aquatic

toxicity of manufactured nanomaterials: challenges and recommendations for future toxicity testing. Environ.

Chem. 11, 207–226. doi:10.1071/EN13221

Schultz, A.G., Ong, K.J., MacCormack, T., Ma, G., Veinot, J.G.C., Goss, G.G., 2012. Silver Nanoparticles Inhibit

Sodium Uptake in Juvenile Rainbow Trout (Oncorhynchus mykiss). Environ. Sci. Technol. 46, 10295–10301.

doi:10.1021/es3017717

Scown, T.M., Santos, E.M., Johnston, B.D., Gaiser, B., Baalousha, M., Mitov, S., Lead, J.R., Stone, V., Fernandes,

T.F., Jepson, M., Aerle, R. van, Tyler, C.R., 2010. Effects of Aqueous Exposure to Silver Nanoparticles of Different

Sizes in Rainbow Trout. Toxicol. Sci. 115, 521–534. doi:10.1093/toxsci/kfq076

Sharma, N., Rather, M.A., Ajima, M.N.O., Gireesh-Babu, P., Kumar, K., Sharma, R., 2016. Assessment of DNA

damage and molecular responses in Labeo rohita (Hamilton, 1822) following short-term exposure to silver

nanoparticles. Food Chem. Toxicol. 96, 122–132. doi:10.1016/j.fct.2016.06.020

Shepard, J.L., Olsson, B., Tedengren, M., Bradley, B.P., 2000. Protein expression signatures identified in Mytilus

edulis exposed to PCBs, copper and salinity stress. Mar. Environ. Res. 50, 337–340. doi:10.1016/S0141-

1136(00)00065-9

Shi, J., Xu, B., Sun, X., Ma, C., Yu, C., Zhang, H., 2013. Light induced toxicity reduction of silver nanoparticles to

Tetrahymena Pyriformis: Effect of particle size. Aquat. Toxicol. 132–133, 53–60.

doi:10.1016/j.aquatox.2013.02.001

Skjolding, L.M., Ašmonaitė, G., Jølck, R.I., Andresen, T.L., Selck, H., Baun, A., Sturve, J., 2017. An assessment of the

importance of exposure routes to the uptake and internal localisation of fluorescent nanoparticles in zebrafish

(Danio rerio), using light sheet microscopy. Nanotoxicology 11, 351–359. doi:10.1080/17435390.2017.1306128

Skjolding, L.M., Kern, K., Hjorth, R., Hartmann, N., Overgaard, S., Ma, G., Veinot, J.G.C., Baun, A., 2014. Uptake and

depuration of gold nanoparticles in Daphnia magna. Ecotoxicology 23, 1172–1183. doi:10.1007/s10646-014-1259-

x

Sornom, P., Felten, V., Médoc, V., Sroda, S., Rousselle, P., Beisel, J.-N., 2010. Effect of gender on physiological and


55

behavioural responses of Gammarus roeseli (Crustacea Amphipoda) to salinity and temperature. Environ. Pollut.

158, 1288–1295. doi:10.1016/j.envpol.2010.01.022

Sroda, S., Cossu-Leguille, C., 2011. Seasonal variability of antioxidant biomarkers and energy reserves in the

freshwater gammarid Gammarus roeseli. Chemosphere 83, 538–544. doi:10.1016/j.chemosphere.2010.12.023

Tedesco, S., Doyle, H., Blasco, J., Redmond, G., Sheehan, D., 2010. Oxidative stress and toxicity of gold

nanoparticles in Mytilus edulis. Aquat. Toxicol. 100, 178–186. doi:10.1016/j.aquatox.2010.03.001

The European Commision, 2011. Commission Recommendation of 18 October 2011 on the definition of

nanomaterialText with EEA relevance - commission-recommendation-on-the-definition-of-nanomater-

18102011_en.pdf [WWW Document]. URL

https://ec.europa.eu/research/industrial_technologies/pdf/policy/commission-recommendation-on-the-

definition-of-nanomater-18102011_en.pdf (accessed 8.30.17).

Thompson, L.B., Carfagno, G.L.F., Andresen, K., Sitton, A.J., Bury, T., Lee, L.L., Lerner, K.T., Fong, P.P., 2017.

Differential uptake of gold nanoparticles by 2 species of tadpole, the wood frog (Lithobates sylvaticus) and the

bullfrog (Lithobates catesbeianus). Environ. Toxicol. Chem. n/a-n/a. doi:10.1002/etc.3909

Tiede, K., Hassellöv, M., Breitbarth, E., Chaudhry, Q., Boxall, A.B.A., 2009. Considerations for environmental fate

and ecotoxicity testing to support environmental risk assessments for engineered nanoparticles. J. Chromatogr. A,

Tools for the REACH Programme - analytical methods for the evaluation of industrial contaminants 1216, 503–509.

doi:10.1016/j.chroma.2008.09.008

US Geological Survey, 2016. Mineral Commodity Summaries 2016 - mcs-2016-silve.pdf [WWW Document]. URL

http://minerals.usgs.gov/minerals/pubs/commodity/silver/mcs-2016-silve.pdf (accessed 3.7.16).

Vale, G., Franco, C., Diniz, M.S., Santos, M.M.C. dos, Domingos, R.F., 2014. Bioavailability of cadmium and

biochemical responses on the freshwater bivalve Corbicula fluminea – the role of TiO2 nanoparticles. Ecotoxicol.

Environ. Saf. 109, 161–168. doi:10.1016/j.ecoenv.2014.07.035

Vance, M.E., Kuiken, T., Vejerano, E.P., McGinnis, S.P., Hochella, M.F., Rejeski, D., Hull, M.S., 2015. Nanotechnology

in the real world: Redeveloping the nanomaterial consumer products inventory. Beilstein J. Nanotechnol. 6, 1769–

1780. doi:10.3762/bjnano.6.181

Vasseur, P., Leguille, C., 2004. Defense systems of benthic invertebrates in response to environmental stressors.

Environ. Toxicol. 19, 433–436. doi:10.1002/tox.20024

Vioque-Fernández, A., Alves de Almeida, E., López-Barea, J., 2009. Assessment of Doñana National Park

contamination in Procambarus clarkii: Integration of conventional biomarkers and proteomic approaches. Sci.


Völker, C., Kämpken, I., Boedicker, C., Oehlmann, J., Oetken, M., 2015. Toxicity of silver nanoparticles and ionic

silver: Comparison of adverse effects and potential toxicity mechanisms in the freshwater clam Sphaerium

corneum. Nanotoxicology 9, 677–685. doi:10.3109/17435390.2014.963723

Volland, M., Hampel, M., Martos-Sitcha, J.A., Trombini, C., Martínez-Rodríguez, G., Blasco, J., 2015. Citrate gold

nanoparticle exposure in the marine bivalve Ruditapes philippinarum: uptake, elimination and oxidative stress

response. Environ. Sci. Pollut. Res. 22, 17414–17424. doi:10.1007/s11356-015-4718-x

Wang, L.-F., Habibul, N., He, D.-Q., Li, W.-W., Zhang, X., Jiang, H., Yu, H.-Q., 2015. Copper release from copper

nanoparticles in the presence of natural organic matter. Water Res. 68, 12–23. doi:10.1016/j.watres.2014.09.031

Wang, Z., von dem Bussche, A., Kabadi, P.K., Kane, A.B., Hurt, R.H., 2013. Biological and environmental


56

transformations of copper-based nanomaterials. ACS Nano 7, 8715–8727. doi:10.1021/nn403080y

Wepener, V., van Dyk, C., Bervoets, L., O’Brien, G., Covaci, A., Cloete, Y., 2011. An assessment of the influence of

multiple stressors on the Vaal River, South Africa. Phys. Chem. Earth Parts ABC, 11th WaterNet/WARFSA/GWP-SA

Symposium: IWRM for National and Regional Integration through Science, Policy and Practice 36, 949–962.

doi:10.1016/j.pce.2011.07.075

Wray, A.T., Klaine, S.J., 2015. Modeling the influence of physicochemical properties on gold nanoparticle uptake

and elimination by Daphnia magna. Environ. Toxicol. Chem. 34, 860–872. doi:10.1002/etc.2881

Yang, Y., Long, C.-L., Li, H.-P., Wang, Q., Yang, Z.-G., 2016. Analysis of silver and gold nanoparticles in environmental

water using single particle-inductively coupled plasma-mass spectrometry. Sci. Total Environ.

doi:10.1016/j.scitotenv.2015.12.150

Zhang, C., Hu, Z., Deng, B., 2016. Silver nanoparticles in aquatic environments: Physiochemical behavior and

antimicrobial mechanisms. Water Res. 88, 403–427. doi:10.1016/j.watres.2015.10.025

Zhang, P., Selck, H., Tangaa, S.R., Pang, C., Zhao, B., 2017. Bioaccumulation and effects of sediment-associated

gold- and graphene oxide nanoparticles on Tubifex tubifex. J. Environ. Sci. 51, 138–145.

doi:10.1016/j.jes.2016.08.015

Chapter 1 Part 2: G. fossarum as a model organism

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CHAPTER 1

Part 2: G. fossarum as a model organism in nanotoxicology

CHAPTER 1

Part 2: G. fossarum as a model organism in nanotoxicology


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1. SYSTEMATIC AND IDENTIFICATION OF Gammarus fossarum

1.1. Systematic position

More than 4500 species belong to the crustacean super-family Gammaridea and represent more than 85%

of amphipods (Bousfield, 1973; Kunz et al., 2010). Gammarids are one of the most widespread group

geographically, with more than 300 species composing the Gammaridae family (Väinölä et al., 2008) (Figure

P2. 1 A, B). They are found throughout a range of marine, freshwater and terrestrial habitats (Bousfield, 1973;

Kunz et al., 2010) whereas the three other amphipod families (Hyperiida, Ingolfiellidea and Caprellida) are

highly specialized and ecologically restricted (Kunz et al., 2010).

Figure P2. 1. (A) Tree of life of the order Amphipoda (Yellow arrow) and its constituents’ suborders Gammaridea,

Senticaudata and Hyperiidea. (B) Tree of life of Gammarus fossarum (Yellow arrow) (Lifemap)

The genus Gammarus, the most important component of the Gammaridae family, is one of the most frequent

in freshwater ecosystems (Barnard and Barnard, 1983; Karaman, 1977). The genus Gammarus counts more

than 100 known species (MacNeil et al., 1997). The classification of Gammarus fossarum, the model organism

used in the present work, is presented below:

AB


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Table P2. 1. Classification of the species Gammarus fossarum

Gammarus are epigean species widely distributed all over the northern hemisphere (Karaman, 1977; Kunz et

al., 2010; Živić and Marković, 2007) (Figure P2.2 A). The most frequent species in Europe are: Gammarus

fossarum (Koch, 1835), Gammarus pulex (Linnaeus, 1758) and Gammarus roeseli (Gervais, 1835) with G.

fossarum being one with the most common and the most diverse species complex in Europe (Weiss et al.,

2014). A species complex represents a group of species being morphologically close but very distinct from a

molecular biology point of view (Brown et al., 1995). The highest diversity of G. fossarum complex is located

in the southern Carpathian and the Balkan peninsula (Copilaş-Ciocianu et al., 2017). G. fossarum is also the

most abundant species in Luxembourg (Figure P2.2 B).

Figure P2. 2. Geographical distribution of Gammarus fossarum in (A) Europe and (B) Luxembourg (Dohet et al.,

2008; Fauna Europaea).

Kingdom Metazoa

Phylum Arthropoda

Subphylum Crustacea

Class Malacostraca

sub-class Eumalacostraca

Super-order Peracarida

Order Amphipoda

Sub-order Senticaudata

Infra-order Gammarida

Parv-order Gammaridira

Super-family Gammaroidea

Family Gammaridae

Genus Gammarus

Species fossarum

FaunaEuropea

A B


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1.2. Identification of Gammarus fossarum

Figure P2. 3. Determination key [in French] of Gammarus fossarum (Felten, 2003).


61

The identification of the Gammarus species in question is generally easy and is performed on mature adult

males as they present morphological characteristics more easily discernible compared to females (Piscart and

Bollache, 2012). The identification key (Figure P2. 3, [in French]) describes the species mainly found in France

and Luxembourg. G. fossarum are characterised by the absence of dorsal carinae (hull) and strips. G. fossarum

are described to have few spines and bristles on the antennae and the extremity of urosoma. The internal

oar of the third uropod is 1/3 the size of the external one. This parameter allows morphological distinction

between G. fossarum and G. pulex (Felten 2003, Figure P2. 3). However, the recent advances in molecular

biology techniques allow a better identification of Gammarus species. Indeed, different cryptic species have

been identified within the G. fossarum complex (Müller, 2000).

The concept of cryptic species is known for more than 300 years (Bickford et al., 2007; Lagrue et al., 2014). It

describes species that are morphologically similar but with a significant genetic differentiation that led to

their wrong classification under the same name (Bickford et al., 2007). The genetic differentiation between

cryptic species is linked to a reproductive isolation that occurred potentially already millions years ago (Webb

and Bartlein, 1992). Three G. fossarum cryptic species were genetically identified, namely Type A, B and C

(Müller, 2000). This split occurred during or before the ice age of the Pleistocene (Webb and Bartlein, 1992)

leading to biological (drift in mitochondrial DNA) rather that morphological differentiation (Feckler et al.,

2012; Müller, 1998; Westram et al., 2010). G. fossarum cryptic species have been reported to have different

geographical distributions (Figure P2. 4). Type A and Type B have been observed in central Europe

(Scheepmaker and van Dalfsen, 1989) and can coexist (Müller, 1998). Type C, in addition to Type B have been

found in Luxembourg (Müller, 1998) with type B being the most abundant one (Annexe 2).

An accurate identification of the cryptic species is crucial in ecotoxicological studies (Feckler et al., 2014;

Lagrue et al., 2014; Sattler et al., 2007; Westram et al., 2010). Differences in sensitivity between species have

already been reported. For instance, G. fossarum were found to be more sensitive to the fungicide

Tebuconazole than G. pulex (Adam et al., 2009). Therefore, it can be assumed that different cryptic species

have different sensitivities to chemicals and deviations among cryptic lineages regarding their physiological

and behavioural characteristics are possible and have been reported (Bickford et al., 2007). A higher

sensitivity to pesticides was observed in G. fossarum Type A compared to Type B (Feckler et al., 2012).

Different sensitivities to parasitism was also highlighted as a higher infection rate by acanthocephalan

parasites was observed in Type B compared to Type A (Westram et al., 2011). All this could explain the

observed species differences in sensitivity to chemicals (Gismondi et al., 2012a, 2012b).


62

Figure P2. 4. Distribution of Gammarus fossarum cryptic species in Europe (Adapted from Westram et al., 2011).

Grey shaded zone indicates the contact zone between Type A and B.

2. MORPHOLOGY

Gammarus are Crustacea malacostraca characterized by a segmented body. They present a bilateral

symmetry, a laterally flattened body, sessile eyes, a pair of antennae with homogenous size and a total length

of 20 mm maximum (Tachet, 2000). Usually, females are smaller than males at the same developmental stage

(Gagné et al., 2005; Tachet, 2000). Compared to shrimps, Gammarus are characterized by the absence of a

carapace which is replaced by a cuticle composed of chitin. The first thoracic segment is fused to the head

and their body is divided in four different parts: prosoma, mesosoma, metasoma and urosoma (Chevreux et

al., 1970, Figure P2. 5). Gammarids usually have large coxal plates and a large abdomen with six pairs of

appendages (Figure P2. 5).


63

Figure P2. 5. General morphology of Gammarus sp. Adapted from Felten 2003.

2.1. Prosoma

Prosoma represents the cephalic region of Gammarus sp. Antennae (2 pairs of sensorial appendages), eyes,

mouth (4 pairs of masticatory appendages), and cephalon compose this region. The eyes are kidney-shaped

and sessile and the antennae have similar sizes and present a flagellum allowing distinction of Gammarus

from Niphargidae (Tachet, 2000). The mouth present mandibles located lateral to the mouth, in conjunction

with the upper and lower lips and surround the mouth opening (MacNeil et al., 1997). They consist of a strong

chitinized incisor, a small accessory plate (the lacinia mobilis), a large medial molar and a spine row between

the molar and the lacinia. In Gammarus, the molar is tough and ridged for crushing and grinding (Lincoln,

1979). Hence, these characteristics allow a wide variety of food to be used by Gammarus sp. (MacNeil et al.,

1997).

2.2. Mesosoma

The mesosoma represents the thoracic region. Seven pereon plates compose this region on which coxal

plates are sealed. Each coxal plate carry a pair of periopods. The first pair represent the gnatopods and they

are used for handling and catching females and for locomotor activity. Males usually present bigger

gnatopods than females. Moreover, these appendages are used for feeding, grooming, burrowing and for

agonistic encounters between males (Borowsky, 1984; MacNeil et al., 1997). Gills and genital organs are also

present in this region fixed under the coxal plates and are in direct contact with the external environment.

Antennae

Peduncle

eyes

Head

GnathopodsPereopods

Uropods

Telson

Pereopods

Pleopods

Pe: Pereo PC: Coxae Pl: Pleon Ur: Urosome


64

2.3. Metasoma

The metasoma represents the abdominal region of a Gammarus body. It carries three pairs of pleopods. In

addition to locomotion, these appendages are used for renewal of water flux on the gills surface. Their

continuous movement is directly linked to oxygen absorption. Therefore, the pleopod beats frequency is

increasingly used as a behavioural biomarker in ecotoxicology studies. This response indicates the

oxygenation rate (Andreï et al., 2016; Arce Funck et al., 2013; Felten et al., 2008; Sornom et al., 2010; Vellinger

et al., 2013).

2.4. Urosoma

The urosoma presents three different segments called uromeres. Each of them carries a uropod pair allowing

the animal to move. Each uropod presents a peduncle, an endopod and an expopod. The shape and size of

the third uropod is different from the two others. This parameter is used for the species determination and

is used as a characteristic for discriminating Gammarus from Echinogammarus. The last segment of the

urosoma present a telson at its extremity.

3. ANATOMY OF Gammarus fossarum

3.1. Nervous system and c irculatory organs

The nervous system consists of a protocerebrum or brain with a long nerval chain covering the entire body

of G. fossarum (Figure P2.6). There are ganglions in the ventral part of the body all the way to the urosoma

and they are all linked with intersegmental nerve connections. All appendages are connected to nervous nets

(Chevreux et al., 1970; Felten, 2003; Charron, 2014). The eyes communicate directly with the ganglia in the

head by large optic nerves. The antenna are sensory and large nerve run out to them as well (Shimek, 2008).

Haemolymph circulation is based on the heart located close to the head (Figure P2.6). This organ has his two

extremities open allowing circulation of haemolymph in all the body. The heart is surrounded by an

heamocoele located on the dorsal side of G. fossarum body and is found in the mesosoma (Felten, 2003;

Charron 2014).

All the organs are in direct contact with haemolymph. Haemolymph plays an important role is transporting

nutrients, hormones, excretions products, ions and oxygen (Nebeker et al., 1992). Haemolymph has an

osmolality around 300 mOsmol allowing a good equilibrium of water and ions in the cells (Charron, 2014).


65

Figure P2. 6. Lateral view of the anatomy of Gammarus sp. Illustrating the principal organs (Schmitz, 1992).

aa: anterior aorta, an: anus, ap: posterior aorta, ba: arterial bulb, cc: circumoesophagial connexion, cd: digestives caeca, cda: anterior digestive caeca, chc: heart chamber, ci: intersegmental connexion, co: heart, cp: posterior caeca, cpy: pyloric chamber, gpe: pereionic ganglion gpl: pereionic ganglion 1, im: midgut, ip: intestinal tract, lds: suspended dorsal ligament, mu: urosomic mass, ms: suboesophagial mass, nan: antennule nerve, nat: antennae nerve, oe: oesophagus, pr: protocerebrum

3.2. Intestina l caeca and digestion system

In Gammarids the mouth is located near the base of the head. It leads to a short oesophagus located before

the stomach where the food is grinded by chitinized plates (Shimek, 2008; Figure P2.6). Behind the stomach

is the midgut located which is as long as G. fossarum body. The midgut is surrounded by two pairs of pouches

or intestinal caeca. Two caeca are located on the side and the two other are located at the bottom of the

midgut (Shimek, 2008). These organs are the central site of metabolism were both digestion and secretion of

digestive enzymes occur. Intestinal caeca or hepatopancreatic caeca are the central organs of digestions in

amphipods. The caeca play a role in nutrient uptake, excretion, moulting cycle, storage of inorganic reserves

and lipid and carbohydrates metabolism (Chevreux et al., 1970; Correia et al., 2002; Grassé, 1961; Schmitz

and Scherrey, 1983; Charron, 2014). They present a tubular structure surrounded with epithelial cells and

muscles covered with connective tissues (Schmitz and Scherrey, 1983,

Figure P2. 7 B). There are four different types of cells forming the epithelial tissues in amphipods (cells E, F,

R and B). The epithelial cells form a barrier between the lumen of the gut and the haemolymph (Kutlu et al.,

2002). The plant material and the fungi are digested in the midgut using cellulase (Chamier and Willoughby,

1986; MacNeil et al., 1997).


66

3.3. Gammarus g il ls and respiration

Gammarids have 6 pairs of gills in direct contact with the external environment. They are present under the

thoracic appendages (Felten and Guérold, 2006; Sutcliffe, 1984). Gills are thin flattened oval plates called

lamellae formed from the epipodits of the pereopods (Sutcliffe, 1984, Figure P2. 7 A, C).

Crustacean gills are lined by a single layer of epithelial cells whose basement membrane at the serosal side is

directly bathed by the haemolymph (Figure P2. 7 C). The mucosal side is covered by a chitinous cuticle and

faces the external medium. The thickness of the epithelium varies depending on its physiological and

biochemical function (Dunel-Erb et al., 1997; Martinez et al., 2005). Thinner regions of the epithelium (1-5

µm) are associated with gaseous exchanges and passive diffusion of ions while thicker epithelium (10-20 µm)

are mainly concerned with ions transport mechanisms such as sodium potassium-adenosine triphosphate

(Na+K+-ATPase) (Henry et al., 2012). It has been described that the thick hydrophobic cuticle on the surface

body limit the gaseous exchanges and electrolytes. They play a crucial role in gaseous exchanges, respiration,

ions absorption, osmoregulation, acid-base balance, calcium homeostasis, extraction of ammonia and

intracellular pH regulation (Mantel and Farmer, 1983; Pequeux, 1995).

In addition to the gills, the pleopods present in the metasoma area are in continuous movement for the

renewal of the water influx on the surface of the gills leading to the improvement of oxygen absorption

(Sutcliffe, 1984). The diffusion of the oxygen is therefore dependent on the surface and thickness of the gills,

the thickness of the water layer around the gills and the difference in the internal and external pressure on

oxygen (Sutcliffe, 1984). Once the oxygen is taken up, it is trapped by hemocyanin which allow its transport

from the haemolymph to the different organs.

Figure P2. 7. Cross section of (A) Gammarus fossarum illustrating the main organs. (B) midgut and intestinal

caeca and (B) gills of G. fossarum observed with optical microscope at 60x magnification. MG: midgut, IC:

intestinal caeca (Pictures: Chauvière A. and Mehennaoui K.)


67

3.4. Reproduction organs, life cyc le and development

3.4.1. Anatomy of the reproduction organs

Two long tubules compose the reproduction apparel in males and are located above the midgut. They are

divided in three different parts: testicle, seminal vesicle and the different canal. Each canal lead to the

periomere 7 where the genital papillae are located (Figure P2. 8). The female reproductive organs consist of

two ovaries and are also located above the midgut. In periomere 5, an oviduct stands out from each ovary

and reach the ventral side of the G. fossarum body. Each oviduct leads to the osteitis where the embryo

development until hatching of eggs takes place (Figure P2. 8) (Charron, 2014; Felten 2003, Trapp 2015).

Figure P2. 8. Reproduction organs of a male

Gammarus and a female (Trapp, 2015). cmu: mucus

cells, GA: androgen gland, ci: incubation chamber,

mt: non-differentiated mesenchymal tissue, ov1:

primer vitellogenese oocyte, ov2: secondary

vitellogenese oocyte, ovd: oviduct, ovd vst: vestigial

oviduct, spc: spermatocyte, spg: spermatogonium,

spz: spermatozoid, vd: spermiduct

3.4.2. Reproduction and l i fe cycle

Gammarus sp. have a complex life cycle, which is of value in ecotoxicological studies (Kunz et al., 2010).

Generally, Gammarids reproduce throughout most of the year with a peak in spring and early summer (Pöckl

et al., 2003). Females are available for mating only during brief periods directly after moulting while males

are available for mating during almost the entire moult cycle (Sutcliffe, 1993). This results in a male-biased

operational sex-ratio. Therefore, males are usually engaged in precopula mate guarding when meeting a

female close to the moult stage (Ridely 1983). When the females are sexually mature, they release

pheromones in the environment. Males detect these pheromones with the chemoreceptors present on their

antennae (Sutcliffe, 1992). Males grab different females before choosing the one that is likely to produce

more eggs than the others (Kunz et al., 2010). Pairing in Gammarus is size-dependent as bigger males grab

bigger females (Kunz et al., 2010). Males will hold the female under and parallel to their body using the first


68

pair of gnatopodes (Borowsky, 1984; Chevreux et al., 1970). Hence, males present bigger gnatopodes

compared to females. This characteristic is used for gender differentiation in sorting (Figure P2. 9). Female

may produce 6 to 10 broods of juveniles and three generations may be reproducing at the same time in

midsummer (Pöckl et al., 2003; Sutcliffe, 1992). Once the female sheds her skin, male can mate with her and

release her. Female carries the developing eggs in her brooding pouch. Juveniles hatches after 1 to 3 weeks

and remains in the brooding pouch until the next moulting stage. After 4 to 6 weeks juveniles swim out of

the brooding pouch. They feed first by coprophagy and then their diet starts to include conditioned leaves

when they reach 1 month age (McCahon and Pascoe, 1988). Gammarids can live for 1 to 2.5 years (Kunz et

al., 2010).

Figure P2. 9. Gammarus fossarum male and female forming a prepopulate pair (Picture: Untereiner B. and Mehennaoui K.)

3.4.3. Development

Gammarids growing is effective through a successive moult process as the presence of the cuticle does not

allow a continuous growth (Sutcliffe, 1992). A cycle of moulting and inter-moulting steps can take place for

many weeks (~4 weeks) and is dependent on the water temperature (12 °C). One cycle is divided in six steps

• Two pre-moult phases (1-3 days, each) lead to a thickening of the existing cuticle

• Two inter-moult phases (9 days each) lead to a total change of the existing cuticle and they are

characterised by an increase in Gammarus permeability for water (Lockwood and Inman, 1973)

• Two post-moult phases (3 to 5 days each) lead to the formation of the new cuticle

Whereas many crustaceans’ juvenile present larvae stages during their life cycle, Gammarus juvenile are

directly similar to adults. Genital papillae appear in males and osteitis appear in females after the fifth moult

stage. Then, the gnatopodes are differentiated and finally the reproduction organs are well differentiated

0.5mm


69

after 9 to 10 moult stages (Gross et al., 2001). Generally, males have a longer life time than females. Females

can leave to up to 23 months while male can live for up to 30 months (Leroy et al., 2010; Pöckl et al., 2003).

Moulting is a very critical process during Gammarus lifetime. Indeed, cuticle is described to be impermeable

to water and pollutants (Rainbow, 1997, 1995), the inter-moult stage may represent a very critical moment

for the animals. Hence, it had been suggested that Gammarids use more energy for osmoregulation processes

and if needed for defence mechanisms (Lockwood and Inman, 1973).

4. Gammarus sp. ECOLOGY

Abiotic factors such as temperature, salinity, oxygen, acidity and pollution play an important role in the

distribution of Gammarus species (Kunz et al., 2010; Meijering, 1991; Whitehurst and Lindsey, 1990). As

benthic macroinvertebrates and shredders, gammarids are usually found in high abundance under rocks,

gravels, in coarse substrates and among living and dead vegetation (Kunz et al., 2010). These substrates offer

shelter from predators and a supply of organic detritus leading to a high presence of amphipods species,

making them one of the most important component of the aquatic macroinvertebrate assemblage in terms

of density and biomass (Kunz et al., 2010; MacNeil et al., 1997). Gammarus fossarum are usually found in

central and eastern areas in Europe headwaters. They are therefore considered as a typical woodland-brook

element in respect to their feeding habits and their resistance to currents (Meijering, 1991). Hence, it had

been described that G. fossarum could be the first species to be impacted by souring water and pollution as

breeding conditions in running waters can be impacted (Meijering, 1991).

5. FORAGING PLASTICITY

Gammarus play an important role as shredders in aquatic ecosystems and present a very diversified trophic

repertoire. They have been described as herbivores, detritivores and predators (Dangles and Guérold, 2001;

Kelly et al., 2002). Their diet is composed of stream conditioned leaves, biofilms that grow on the leaves,

dead chironomids, live juvenile isopods and even juvenile and wounded/trapped fish (Fielding et al., 2003;

Kelly et al., 2002; MacNeil et al., 1997). Gammarus have also been described as cannibals (MacNeil et al.,

1997). Their success in colonizing very diverse habitats is linked to their foraging plasticity making them of

high ecological value (Kunz et al., 2010; MacNeil et al., 1997). Generally, Gammarus feed of conditioned alder,

elm or maple leaves. The presence of microorganisms will influence the palatability of leaves (MacNeil et al.,

1997). Gammarus will prefer fresh conditioned leaves than sterile old ones in addition to leaves with low

level of tannin and lignin (MacNeil et al., 1997). The fresh conditioned leaves with high presence of fungi

leads to an increase of 30% of digestibility as leaves are partially degraded by microorganisms (Chamier and

Willoughby, 1986). Moreover, Gammarus were described to differentiate conditioned leaves regarding the

species of fungi present on their surface (Graça et al., 1994).


70

6. Gammarus fossarum AS A MODEL ORGANISM IN ECOTOXICOLOGY

Bioindicators have been defined as species that are sensitive to different kind of stressors, give significant

biochemical and physiological responses to anthropogenic pollutants and allow the predictions of the effects

of contaminants on higher level of biological organisations (population, community, ecosystems) (Vellinger,

2012). Gammarids fulfil all these criteria and are therefore increasingly used as bioindicators of the quality of

aquatic ecosystems (Farkas et al., 2003). Additionally, Gammarids have a well-known ecology and sensitivity

to different pollutants (Kunz et al., 2010) and are increasingly used as model organisms in ecotoxicology

studies (Andreï et al., 2016; Arce Funck et al., 2013; Felten et al., 2006; Gismondi et al., 2012a; Vellinger et

al., 2013).

As described above, G. fossarum is considered as an ubiquitous species with a wide distribution all over

Europe (Janetzky, 1994; Karaman, 1977; Živić and Marković, 2007). They are found in high abundance and

high density in many freshwaters. Their wide foraging plasticity offer them a central position in the food web

chain. G. fossarum play also a major functional role in litter breakdown process and nutrient cycling (Forrow

and Maltby, 2000; Kelly et al., 2002; Lacaze et al., 2010; MacNeil et al., 1997). They also play an important

structural role as they represent an important food source for other aquatic species like fish, amphibians and

other invertebrates (Kelly et al., 2002; MacNeil et al., 1997) and they can also be prey for birds like ducks.

Many individuals are parasitized with acanthocephalans which modify their behavior and make them swim

close to the water surface and be more available for the birds the final host of the parasite (Gismondi et al.,

2012a).

In contrast with other macroinvertebrates, Gammarids have a long life cycle, as it could reach 2.5 years.

Therefore, they are able to integrate the different contamination fluxes that occur in the aquatic ecosystems.

Gammarids have also been described as good accumulators of metal contaminations as the internal

concentrations measured within the animal were usually proportional and dependent on the concentrations

present in the environment (Arce Funck et al., 2013; Kunz et al., 2010; Lebrun et al., 2012; Vellinger et al.,

2012).

Gammarids are easy to collect, handle and manipulate in the laboratory and it is possible to collect them in

the field all year long. They are very easy to identify and genders are easy to differentiate. They can be used

for laboratory experiment as well as in situ experiments (Andreï et al., 2016; Besse et al., 2013; Coulaud et

al., 2011; Dedourge-Geffard et al., 2009). G. fossarum is described as a sensitive species to contaminants.

Thus, they are frequently used as bioindicators as their presence or absence in aquatic ecosystems, their

density, population dynamic are good indicators of the quality of the aquatic environment. Furthermore, they

are considered as a good model for integrative studies by enhancing the link between responses observed at


71

the molecular level (omics) to responses observed at the individual and functional level (Andreï et al., 2016;

Lagadic et al., 1994).

REFERENCES

Adam, O., Badot, P.-M., Degiorgi, F., Crini, G., 2009. Mixture toxicity assessment of wood preservative pesticides

in the freshwater amphipod Gammarus pulex (L.). Ecotoxicol. Environ. Saf. 72, 441–449.

doi:10.1016/j.ecoenv.2008.07.017

Andreï, J., Pain-Devin, S., Felten, V., Devin, S., Giambérini, L., Mehennaoui, K., Cambier, S., Gutleb, A.C., Guérold,

F., 2016. Silver nanoparticles impact the functional role of Gammarus roeseli (Crustacea Amphipoda). Environ.

Pollut. 208, Part B, 608–618. doi:10.1016/j.envpol.2015.10.036



143, 73–84. doi:10.1016/j.aquatox.2013.07.012

Barnard, J.L., Barnard, C.M., 1983. Freshwater amphipoda of the world / J.L. & C.M. Barnard. - Version details

[WWW Document]. Mt Vernon Va Hayfield Assoc. URL http://trove.nla.gov.au/version/26012490 (accessed

4.22.16).

Besse, J.-P., Coquery, M., Lopes, C., Chaumot, A., Budzinski, H., Labadie, P., Geffard, O., 2013. Caged Gammarus

fossarum (Crustacea) as a robust tool for the characterization of bioavailable contamination levels in continental

waters: Towards the determination of threshold values. Water Res. 47, 650–660.

doi:10.1016/j.watres.2012.10.024

Bickford, D., Lohman, D.J., Sodhi, N.S., Ng, P.K.L., Meier, R., Winker, K., Ingram, K.K., Das, I., 2007. Cryptic species

as a window on diversity and conservation. Trends Ecol. Evol. 22, 148–155. doi:10.1016/j.tree.2006.11.004

Borowsky, B., 1984. The Use of the Males' Gnathopods During Precopulation in Some Gammaridean

Amphipods. Crustaceana 47, 245–250. doi:10.1163/156854084X00504

Bousfield, E.L., 1973. shallow-water-gammaridean-amphipoda-of-new-england-handbooks-of-american.pdf

[WWW Document]. URL http://www.thebookishblog.com/shallow-water-gammaridean-amphipoda-of-new-

england-handbooks-of-american.pdf (accessed 8.20.17).

Brown, J.K., Frohlich, D.R., Rosell, R.C., 1995. The Sweetpotato or Silverleaf Whiteflies: Biotypes of Bemisia tabaci

or a Species Complex? Annu. Rev. Entomol. 40, 511–534. doi:10.1146/annurev.en.40.010195.002455

Chamier, A.-C., Willoughby, L.G., 1986. The role of fungi in the diet of the amphipod Gammarus pulex (L.): an

enzymatic study. Freshw. Biol. 16, 197–208. doi:10.1111/j.1365-2427.1986.tb00964.x

Chevreux, É., Fage, L., Faunistique, F. des S.F. de S.N.O.C. de, 1970. Amphipodes. Kraus.

Copilaş-Ciocianu, D., Rutová, T., Pařil, P., Petrusek, A., 2017. Epigean gammarids survived millions of years of severe

climatic fluctuations in high latitude refugia throughout the Western Carpathians. Mol. Phylogenet. Evol. 112, 218–

229. doi:10.1016/j.ympev.2017.04.027

Correia, A.D., Pereira, A.L., Costa, M.H., Carrapi??o, F., 2002. Functional anatomy of the midgut gland of Gammarus

locusta (Crustacea: Amphipoda). J. Mar. Biol. Assoc. UK 82, 201–204.

Coulaud, R., Geffard, O., Xuereb, B., Lacaze, E., Quéau, H., Garric, J., Charles, S., Chaumot, A., 2011. In situ feeding

assay with Gammarus fossarum (Crustacea): Modelling the influence of confounding factors to improve water


72

quality biomonitoring. Water Res. 45, 6417–6429. doi:10.1016/j.watres.2011.09.035

Dangles, O., Guérold, F., 2001. Linking Shredders and Leaf Litter Processing: Insights from an Acidic Stream Study.

Int. Rev. Hydrobiol. 86, 395–406. doi:10.1002/1522-2632(200107)86:4/5<395::AID-IROH395>3.0.CO;2-1

Dedourge-Geffard, O., Palais, F., Biagianti-Risbourg, S., Geffard, O., Geffard, A., 2009. Effects of metals on feeding

rate and digestive enzymes in Gammarus fossarum: An in situ experiment. Chemosphere 77, 1569–1576.


Dohet, A., Ector, L., Cauchie, H.-M., Hoffmann, L., 2008. Identification of benthic invertebrate and diatom indicator

taxa that distinguish different stream types as well as degraded from reference conditions in Luxembourg. Anim.

Biol. 58, 419–472. doi:10.1163/157075608X383719

Dunel-Erb, S., Barradas, C., Lignon, J., 1997. Morphological Evidence for the Existence of Two Distinct Types of

Mitochondria Rich Cells in the Gill of the Crayfish Astacus leptodactylusEschscholtz. Acta Zool. 78, 195–203.

doi:10.1111/j.1463-6395.1997.tb01006.x

Farkas, A., Salánki, J., Varanka, I., 2003. Crustaceans as biological indicators of heavy metal pollution in Lake

Balaton (Hungary). Hydrobiologia 506–509, 359–364. doi:10.1023/B:HYDR.0000008615.86330.1d

Fauna Europaea : Distribution [WWW Document], n.d. URL http://www.faunaeur.org/distribution.php (accessed

2.17.14).

Feckler, A., Thielsch, A., Schwenk, K., Schulz, R., Bundschuh, M., 2012. Differences in the sensitivity among cryptic

lineages of the Gammarus fossarum complex. Sci. Total Environ. 439, 158–164.

doi:10.1016/j.scitotenv.2012.09.003

Feckler, A., Zubrod, J.P., Thielsch, A., Schwenk, K., Schulz, R., Bundschuh, M., 2014. Cryptic species diversity: an

overlooked factor in environmental management? J. Appl. Ecol. 51, 958–967. doi:10.1111/1365-2664.12246

Felten, V., Baudoin, J.M., Guérold, F., 2006. Physiological recovery from episodic acid stress does not mean

population recovery of Gammarus fossarum. Chemosphere 65, 988–998. doi:10.1016/j.chemosphere.2006.03.059

Felten, V., Charmantier, G., Mons, R., Geffard, A., Rousselle, P., Coquery, M., Garric, J., Geffard, O., 2008.

Physiological and behavioural responses of Gammarus pulex (Crustacea: Amphipoda) exposed to cadmium. Aquat.

Toxicol. 86, 413–425. doi:10.1016/j.aquatox.2007.12.002

Felten, V., Guérold, F., 2006. Short-term physiological responses to a severe acid stress in three macroinvertebrate

species: A comparative study. Chemosphere 63, 1427–1435. doi:10.1016/j.chemosphere.2005.10.003

Fielding, N.J., MacNeil, C., Dick, J.T.A., Elwood, R.W., Riddell, G.E., Dunn, A.M., 2003. Effects of the

acanthocephalan parasite Echinorhynchus truttae on the feeding ecology of Gammarus pulex (Crustacea:

Amphipoda). J. Zool. 261, 321–325. doi:10.1017/S0952836903004230

Forrow, D.M., Maltby, L., 2000. Toward a mechanistic understanding of contaminant-induced changes in detritus

processing in streams: Direct and indirect effects on detritivore feeding. Environ. Toxicol. Chem. 19, 2100–2106.

doi:10.1002/etc.5620190820

Gagné, F., Blaise, C., Pellerin, J., 2005. Altered exoskeleton composition and vitellogenesis in the crustacean

Gammarus sp. collected at polluted sites in the Saguenay Fjord, Quebec, Canada. Environ. Res. 98, 89–99.

doi:10.1016/j.envres.2004.09.008

Gismondi, E., Cossu-Leguille, C., Beisel, J.-N., 2012a. Acanthocephalan parasites: help or burden in gammarid



73

Gismondi, E., Rigaud, T., Beisel, J.-N., Cossu-Leguille, C., 2012b. Microsporidia parasites disrupt the responses to


Graça, M. a. S., Maltby, L., Calow, P., 1994. Comparative ecology of Gammarus pulex (L.) and Asellus aquaticus (L.)

I: population dynamics and microdistribution. Hydrobiologia 281, 155–162. doi:10.1007/BF00028678

Grassé, P.-P., 1961. Traité de zoologie: anatomie, systématique, biologie. Tome IV, Premier fascicule Tome IV,

Premier fascicule. Masson, Paris.

Gross, M.Y., Maycock, D.S., Thorndyke, M.C., Morritt, D., Crane, M., 2001. Abnormalities in sexual development of

the amphipod Gammarus pulex (L.) found below sewage treatment works. Environ. Toxicol. Chem. 20, 1792–1797.

doi:10.1002/etc.5620200824

Henry, R.P., Lucu, Č., Onken, H., Weihrauch, D., 2012. Multiple functions of the crustacean gill: osmotic/ionic

regulation, acid-base balance, ammonia excretion, and bioaccumulation of toxic metals. Front. Physiol. 3.

doi:10.3389/fphys.2012.00431

Janetzky, W., 1994. Distribution of the genus Gammarus (Amphipoda: Gammaridae) in the River Hunte and its

tributaries (Lower Saxony, northern Germany). Hydrobiologia 294, 23–34. doi:10.1007/BF00017622

Karaman, G., 1977. freshwater Gammarus species from europe, North Africa and adjacent regions of Asia.

Gammarus pulex group and related species.pdf [WWW Document]. URL

http://www.nativefishlab.net/library/textpdf/17884.pdf (accessed 2.27.14).

Kelly, D.W., Dick, J.T.A., Montgomery, W.I., 2002. The functional role of Gammarus(Crustacea, Amphipoda):

shredders, predators, or both? Hydrobiologia 485, 199–203. doi:10.1023/A:1021370405349

Kunz, P.Y., Kienle, C., Gerhardt, A., 2010. Gammarus spp. in Aquatic Ecotoxicology and Water Quality Assessment:

Toward Integrated Multilevel Tests, in: Whitacre, D.M. (Ed.), Reviews of Environmental Contamination and

Toxicology Volume 205, Reviews of Environmental Contamination and Toxicology. Springer New York, pp. 1–76.

Kutlu, M., Düzen, A., Bayçu, C., Özata, A., 2002. A transmission electron microscope investigation of the effect of

lead acetate on the hepatopancreatic ceca of Gammarus pulex. Environ. Toxicol. Pharmacol. 12, 181–187.

doi:10.1016/S1382-6689(02)00036-4

Lacaze, E., Geffard, O., Bony, S., Devaux, A., 2010. Genotoxicity assessment in the amphipod Gammarus fossarum

by use of the alkaline Comet assay. Mutat. Res. Toxicol. Environ. Mutagen. 700, 32–38.

doi:10.1016/j.mrgentox.2010.04.025

Lagadic, L., Caquet, T., Ramade, F., 1994. The role of biomarkers in environmental assessment (5). Invertebrate

populations and communities. Ecotoxicology 3, 193–208. doi:10.1007/BF00117084

Lagrue, C., Wattier, R., Galipaud, M., Gauthey, Z., Rullmann, J.-P., Dubreuil, C., Rigaud, T., Bollache, L., 2014.

Confrontation of cryptic diversity and mate discrimination within Gammarus pulex and Gammarus fossarum

species complexes. Freshw. Biol. 59, 2555–2570. doi:10.1111/fwb.12453

Lebrun, J.D., Perret, M., Geffard, A., Gourlay-Francé, C., 2012. Modelling copper bioaccumulation in Gammarus

pulex and alterations of digestive metabolism. Ecotoxicology 21, 2022–2030. doi:10.1007/s10646-012-0955-7

Leroy, D., Haubruge, E., De Pauw, E., Thomé, J.P., Francis, F., 2010. Development of ecotoxicoproteomics on the

freshwater amphipod Gammarus pulex: identification of PCB biomarkers in glycolysis and glutamate pathways.

Ecotoxicol. Environ. Saf. 73, 343–352. doi:10.1016/j.ecoenv.2009.11.006

Lincoln, R.J., 1979. British Marine Amphipoda: Gammaridea. - -With 290 figs., 658 pp. London: British Museum

(Natural History) 1979. ISBN 0 565 00818 8. £ 50.00. Int. Rev. Gesamten Hydrobiol. Hydrogr. 66, 446–447.


74

doi:10.1002/iroh.19810660336

Lockwood, A.P.M., Inman, C.B.E., 1973. The blood volume of some amphipod crustaceans in relation to the salinity

of the environment they inhabit. Comp. Biochem. Physiol. A Physiol. 44, 935–941. doi:10.1016/0300-

9629(73)90158-8

MacNeil, C., Dick, J.T.A., Elwood, R.W., 1997. The Trophic Ecology of Freshwater Gammarus Spp.

(crustacea:amphipoda): Problems and Perspectives Concerning the Functional Feeding Group Concept. Biol. Rev.

72, 349–364. doi:10.1111/j.1469-185X.1997.tb00017.x

Mantel, L.H., Farmer, L.L., 1983. 2 - Osmotic and Ionic Regulation, in: Linda Mantel (Ed.), Internal Anatomy and

Physiological Regulation. Academic Press, pp. 53–161.

Martinez, A.-S., Charmantier, G., Compère, P., Charmantier-Daures, M., 2005. Branchial chamber tissues in two

caridean shrimps: the epibenthic Palaemon adspersus and the deep-sea hydrothermal Rimicaris exoculata. Tissue

Cell 37, 153–165. doi:10.1016/j.tice.2004.12.004

McCahon, C.P., Pascoe, D., 1988. Use of Gammarus pulex (L.) in safety evaluation tests: Culture and selection of a

sensitive life stage. Ecotoxicol. Environ. Saf. 15, 245–252. doi:10.1016/0147-6513(88)90078-4

Meijering, M.P.D., 1991. Lack of oxygen and low pH as limiting factors for Gammarus in Hessian brooks and rivers.

Hydrobiologia 223, 159–169. doi:10.1007/BF00047637

Müller, J., 2000. Mitochondrial DNA Variation and the Evolutionary History of Cryptic Gammarus fossarum Types.

Mol. Phylogenet. Evol. 15, 260–268. doi:10.1006/mpev.1999.0740

Müller, J., 1998. Genetic population structure of two cryptic Gammarus fossarum types across a contact zone. J.

Evol. Biol. 11, 79–101. doi:10.1046/j.1420-9101.1998.11010079.x

Nebeker, A.V., Dominguez, S.E., Chapman, G.A., Onjukka, S.T., Stevens, D.G., 1992. Effects of low dissolved oxygen

on survival, growth and reproduction of Daphnia, Hyalella and Gammarus. Environ. Toxicol. Chem. 11, 373–379.

doi:10.1002/etc.5620110311

Pequeux, A., 1995. OSMOTIC REGULATION IN CRUSTACEANS. J. Crustac. Biol. 15, 1–60.

doi:10.1163/193724095X00578

Piscart, C., Bollache, L., 2012. Crustacés amphipodes de surface : gammares d’eau douce., Introduction pratique à

la systématique des organismes des eaux continentales de France. Association Française de Limnologie.

Pöckl, M., Webb, B.W., Sutcliffe, D.W., 2003. Life history and reproductive capacity of Gammarus fossarum and G.

roeseli (Crustacea: Amphipoda) under naturally fluctuating water temperatures: a simulation study. Freshw. Biol.

48, 53–66. doi:10.1046/j.1365-2427.2003.00967.x

Rainbow, P.S., 1997. Ecophysiology of Trace Metal Uptake in Crustaceans. Estuar. Coast. Shelf Sci. 44, 169–175.

doi:10.1006/ecss.1996.0208

Rainbow, P.S., 1995. Physiology, physicochemistry and metal uptake—A crustacean perspective. Mar. Pollut. Bull.

31, 55–59. doi:10.1016/0025-326X(95)00005-8

Sattler, T., Bontadina, F., Hirzel, A.H., Arlettaz, R., 2007. Ecological niche modelling of two cryptic bat species calls

for a reassessment of their conservation status. J. Appl. Ecol. 44, 1188–1199. doi:10.1111/j.1365-

2664.2007.01328.x

Scheepmaker, M., van Dalfsen, 1989. Genetic differentiation in Gammarus fossarum and G. caparti (Crustacea,

Amphipoda) with reference to G. pulex pulex in northwestern Europe. Bijdr. Tot Dierkd. 59, 127–139.


75

Schmitz, E.H., Scherrey, P.M., 1983. Digestive anatomy of Halella azteca (Crustacea, Amphipoda). J. Morphol. 175,

91–100. doi:10.1002/jmor.1051750109

Shimek, R.L., 2008. Amphipods [WWW Document]. URL http://reefkeeping.com/issues/2004-09/rs/index.php

(accessed 8.25.17).



158, 1288–1295. doi:10.1016/j.envpol.2010.01.022

Sutcliffe, D.W., 1992. Reproduction in Gammarus (Crustacea, Amphipoda): basic processes. Freshw. Forum 2, 102–

128.

Sutcliffe, D.W., 1984. Quantitative aspects of oxygen uptake by Gammarus (Crustacea, Amphipoda): a critical

review. Freshw. Biol. 14, 443–489. doi:10.1111/j.1365-2427.1984.tb00168.x

Tachet, H., 2000. Invertébrés d’eau douce: systématique, biologie, écologie. CNRS Editions.

Väinölä, R., Witt, J.D.S., Grabowski, M., Bradbury, J.H., Jazdzewski, K., Sket, B., 2008. Global diversity of amphipods

(Amphipoda; Crustacea) in freshwater. SpringerLink 241–255. doi:10.1007/978-1-4020-8259-7_27

Vellinger, C., Felten, V., Sornom, P., Rousselle, P., Beisel, J.-N., Usseglio-Polatera, P., 2012. Behavioural and

Physiological Responses of Gammarus pulex Exposed to Cadmium and Arsenate at Three Temperatures: Individual

and Combined Effects. PLoS ONE 7, e39153. doi:10.1371/journal.pone.0039153

Vellinger, C., Gismondi, E., Felten, V., Rousselle, P., Mehennaoui, K., Parant, M., Usseglio-Polatera, P., 2013. Single

and combined effects of cadmium and arsenate in Gammarus pulex (Crustacea, Amphipoda): Understanding the

links between physiological and behavioural responses. Aquat. Toxicol. 140–141, 106–116.

doi:10.1016/j.aquatox.2013.05.010

Webb, Bartlein, and P.J., 1992. Global Changes During the Last 3 Million Years: Climatic Controls and Biotic

Responses. Annu. Rev. Ecol. Syst. 23, 141–173. doi:10.1146/annurev.es.23.110192.001041

Weiss, M., Macher, J.N., Seefeldt, M.A., Leese, F., 2014. Molecular evidence for further overlooked species within

the Gammarus fossarum complex (Crustacea: Amphipoda). Hydrobiologia 721, 165–184. doi:10.1007/s10750-013-

1658-7

Westram, A.M., Jokela, J., Baumgartner, C., Keller, I., 2011. Spatial Distribution of Cryptic Species Diversity in

European Freshwater Amphipods (Gammarus fossarum) as Revealed by Pyrosequencing. PLoS ONE 6, e23879.

doi:10.1371/journal.pone.0023879

Westram, A.M., Jokela, J., Keller, I., 2010. Isolation and characterization of ten polymorphic microsatellite markers

for three cryptic Gammarus fossarum (Amphipoda) species. Conserv. Genet. Resour. 2, 401–404.

doi:10.1007/s12686-010-9287-1

Whitehurst, I.T., Lindsey, B.I., 1990. The impact of organic enrichment on the benthic macroinvertebrate

communities of a lowland river. Water Res. 24, 625–630. doi:10.1016/0043-1354(90)90195-C

Živić, I., Marković, Z., 2007. Distribution of the Species Gammarus balcanicus and Gammarus fossarum on the

Territory of Serbia (Central Part of the Balkan Peninsula). Crustaceana 80, 67–76. doi:10.2307/20107783

Chapter 2 G. fossarum as a model organism for nano-ecotoxicology

77

CHAPTER 2

Gammarus fossarum (Crustacea Amphipoda) as a model organism for

nano-ecotoxicology

CHAPTER 2

Gammarus fossarum (Crustacea Amphipoda)

as a model organism for nano-ecotoxicology


78

KEY FINDINGS

Aim of the study

G. fossarum was selected to assess effects of well-characterised and well-studied set of AgNPs. Synthetically

produced AgNPs of different sizes (20 and 200 nm) as well as AgNPs synthetized by a biological method (using

plant leaf extracts of Ocinum sanctum and Azadirachta indica, AgNPs 23 and 27 nm, respectively) were used

in order to elucidate the relation between size, synthesis method, NPs surface properties, ions dissolution

and toxicity. A multibiomarker approach was used to investigate the acute effects of these AgNPs on

physiological and behavioural responses of G. fossarum.

Experimental design

Key findings

• Size-dependent effects

o AgNPs 20 nm > AgNPs 200 nm

• Coating-dependent effects

o AgNPs 23 nm > AgNP 27 nm

• High ion release from AgNPs 23 nm compared to AgNPs 20, 27 and 200 nm

• Presence of Ag in gills of G. fossarum exposed AgNPs 23 nm and AgNPs 27 nm

• Significant Ag uptake from AgNO3 and AgNPs 23 nm

• AgNO3 and AgNPs 23 nm led to an osmoregulation impairment and a decrease in G. fossarum

locomotion

• The observed effects seemed to be linked to Ag ions released from AgNPs 23 nm


79

ARTICLE 1

Science of the Total Environment 566, 1649-1659.

Gammarus fossarum (Crustacea, Amphipoda) as a model organism to study silver nanoparticles effects

Kahina Mehennaoui a,b, Anastasia Georgantzopoulou a,d, Vincent Felten b, Jennifer Andreï b, Maël Garaud b,

Sébastien Cambier a, Tommaso Serchi a, Sandrine Pain-Devin b, François Guérold b, Jean-Nicolas Audinot c,

Laure Giambérini b, Arno C. Gutleb a *

a Environmental Research and Innovation (ERIN) Department, Luxembourg Institute of Science and

Technology (LIST), 5 avenue des Hauts-Forneaux, Esch-sur-Alzette, Luxembourg.

b Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360, Université de

Lorraine – Metz, France.

c Materials Research and Technology (MRT) Department, Luxembourg Institute of Science and Technology,

(LIST), 5 avenue des Hauts-Forneaux, Esch-sur-Alzette, Luxembourg.

d Norwegian Institute for Water Research (NIVA), Gaustadalléen 21, NO-0349 Oslo, Norway.

We wish to dedicate this manuscript to the late Stephen J. Klaine who inspired the work of our group for many

years.

Gammarus fossarum (Crustacea, Amphipoda) as a model organism tostudy the effects of silver nanoparticles☆

Kahina Mehennaoui a,b, Anastasia Georgantzopoulou a,d, Vincent Felten b, Jennifer Andreï b, Maël Garaud b,Sébastien Cambier a, Tommaso Serchi a, Sandrine Pain-Devin b, François Guérold b, Jean-Nicolas Audinot c,Laure Giambérini b, Arno C. Gutleb a,⁎

a Environmental Research and Innovation (ERIN) Department, Luxembourg Institute of Science and Technology (LIST), 5 avenue des Hauts-Fourneaux, Esch-sur-Alzette, Luxembourgb Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360, Université de Lorraine, Metz, Francec Materials Research and Technology (MRT) Department, Luxembourg Institute of Science and Technology, (LIST), 5 avenue des Hauts-Fourneaux, Esch-sur-Alzette, Luxembourgd Norwegian Institute for Water Research (NIVA), Gaustadalléen 21, NO-0349 Oslo, Norway

H I G H L I G H T S

• Gammarus fossarum was used as modelorganism to study silver nanoparticleseffects.

• Four AgNPs were used to link ions re-lease and surface chemistry to the ef-fects.

• AgNPs sub-lethal effects were investi-gated using multibiomarker approach.

• Survival, heamolymph osmolality andlocomotor activity of G. fossarum wereimpacted.

• Ions released by AgNPs dissolutioncould be a main factor of toxicity.

G R A P H I C A L A B S T R A C T

a b s t r a c ta r t i c l e i n f o

Article history:

Received 9 March 2016Received in revised form 10 June 2016Accepted 10 June 2016Available online 18 June 2016

Editor: D. Barcelo

Amphipods are one of the most important components of freshwater ecosystems. Among them, gammarids arethemostwidespread group in Europe and are often used as bioindicators andmodel organisms in ecotoxicology.However, their use, especially of Gammarus fossarum for the study of the environmental impact of nanoparticles,has been rather limited so far.G. fossarum was selected to assess effects of well-characterized chemically synthesized silver nanoparticles(AgNPs 20 nm and 200 nm) and “green” laboratory synthetized (from plant leaf extracts) AgNPs (AgNPs23 nm and 27 nm). AgNO3 was used as a positive control to compare AgNPs effects and silver ions effects. Amultibiomarker approachwas used to investigate the sub-lethal effects of AgNPs on physiological and behaviour-al responses of G. fossarum.Two different experiments were carried out. In a preliminary experiment, two populations of G. fossarum (G.f1and G.f2) were tested for sensitivity differences and the most sensitive one was exposed, in a final experiment,to sub-lethal concentrations of AgNO3 and the most toxic AgNPs. AgNO3 and AgNPs 23 nm led to a significant

Keywords:

Gammarus fossarum

Silver nanoparticlesLocomotor activityOsmoregulationBiomarkers

Science of the Total Environment 566–567 (2016) 1649–1659

☆ We wish to dedicate this manuscript to the late Stephen J. Klaine who inspired the work of our group for many years.⁎ Corresponding author at: Environmental Research and Innovation (ERIN) Department, Luxembourg Institute of Science and Technology, 5, avenue des Hauts-Fourneaux, L-4362 Esch-

sur-Alzette, Luxembourg.E-mail address: [email protected] (A.C. Gutleb).

http://dx.doi.org/10.1016/j.scitotenv.2016.06.0680048-9697/© 2016 Elsevier B.V. All rights reserved.

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decrease in survival rates, osmoregulation and locomotor activity. Ag internalisation, performedwith SecondaryIon Mass Spectrometry (SIMS), showed the presence of silver in gills of G.f2 exposed to AgNPs 23 and 27 nm.This study highlighted the influence ofmethod of synthesis on ion release, uptake and toxic effects of AgNPs onG.fossarum. Osmoregulation appeared to be an effective biomarker indicating the physiological health status of G.fossarum. Locomotor activity, which was the most impacted response, reflects the potential effects of releasedions from AgNPs 23 nm at the population level as locomotion is necessary for foraging, finding mates and escap-ing from predators. Therefore, we propose G. fossarum as a suitable model for environmental nanotoxicology,providing information both at individual and population levels.

© 2016 Elsevier B.V. All rights reserved.

1. Introduction

The increasing production of nanomaterials and their ongoing use ina wide range of industries leads to their inevitable release into the envi-ronment (Bhatt and Tripathi, 2011; Klaine et al., 2008). According to theWoodrow Wilson Database, 1814 nanotech consumer products existand 435 of them contain silver nanoparticles (AgNPs) (Vance et al.,2015). AgNPs have distinctive chemical and physical properties suchas catalytic and electronic properties, thermal conductivity and non-lin-ear optical behaviour (Fabrega et al., 2011; Sharma et al., 2014), andthey are widely used as bactericides in a variety of products such as cos-metics, clothing, detergents, water filters, etc. The global annual silverproduction was estimated to be about 27,000 t (US Geological Survey,2016), and due to the increasing usage, large quantities of silver maybe potentially released into the environment and enter aquatic ecosys-tems where Ag and AgNPs persist and bioaccumulate (Fabrega et al.,2011). Sound data on the environmental concentration of AgNPs arestill missing and somemodels calculated predicted environmental con-centrations of AgNPs in water ranging from 0.6 ng·L−1 to 0.32 μg·L−1

(Batley et al., 2013; Sun et al., 2014; Gottschalk et al., 2013). AgNPswere reported to contribute to only 1–15% of the total Ag present inthe environment (Blaser et al., 2008), however, given their expandeduse, AgNPs concentrations are expected to increase in the future.

Despite the growing number of studies on the toxicity of AgNPs,their behaviour, their environmental fate in aquatic environments andthe consequent effects on living organisms are still not fully understood(Foldbjerg et al., 2015; Vale et al., 2016). A few studies investigated andclearly demonstrated that AgNPs are taken up in aquatic organisms suchas molluscs (Ali, 2014; Buffet et al., 2013; Canesi et al., 2012; Ringwoodet al., 2010), crustaceans (Andreï et al., 2016; Georgantzopoulou et al.,2013; Ulm et al., 2015) and fish (Ašmonaite et al., 2016; Griffitt et al.,2013; Jung et al., 2014; Kwok et al., 2012), and behave differently de-pending on their physico-chemical characteristics. In this way, size,coating and chemical composition were suggested as important con-tributing factors to the toxicity of AgNPs (Castranova, 2011; Vale et al.,2016). For instance, laboratory synthesised “coffee-coated” AgNPswere reported to be more toxic to Daphnia magna than commercialPVP-coated and non-coated AgNPs (Allen et al., 2010). Similarly, differ-ent kinds of colloidal AgNPs induced higher mortality in D.magna thanAgNPs powder dispersed by sonication, and their effects were size-de-pendent with the smaller particles being more toxic than the largerones (Asghari et al., 2012).

In freshwater ecosystems, amphipods are an important componentand can be the dominant part of benthic macro-invertebrate assem-blages (MacNeil et al., 1997). Gammarus fossarum is increasingly usedin ecotoxicology as test species due to its wide distribution (BarnardandBarnard, 1983) all over Europe and all over Luxembourg, high abun-dance (Felten et al., 2008a; Kunz et al., 2010), clear sexual dimorphism,easiness of collection and handling and its sensitivity to a large range oftoxicants (Issartel et al., 2005; Xuereb et al., 2011; Bundschuh et al.,2013; Arce Funck et al., 2013; Besse et al., 2013). G. fossarum plays amajor functional role in ecosystems in leaf litter breakdown processes,freshwater food chains and nutrient cycling (Forrow and Maltby,2000; Kelly et al., 2002; MacNeil et al., 1997). Consequently, G. fossarum

could provide valuable information, both in laboratory and in situ exper-iments (Besse et al., 2013; Coulaud et al., 2011, 2014), about the poten-tial effects of environmental contaminants including nanoparticles inaquatic ecosystems. To the best of our knowledge, at present, only onestudy used Gammarus to assess AgNPs effects (Andreï et al., 2016). Cit-rate coated AgNPs (10 nm, 0.5 μg·L−1) impacted the functional role ofG. roeseli after 72 h of exposure, disturbed the production of fine partic-ulate organic matter (FPOM) and significantly decreased locomotor ac-tivity of G. roeseli (Andreï et al., 2016), a species with a very restricteddistribution in Luxembourg. However, there is still a lack of informationabout mechanisms of actions underlying AgNPs effects on Gammarids.Therefore, multibiomarker approaches, including biochemical (e.g. de-fencemechanisms, antioxidant responses), physiological (e.g. osmoreg-ulation) and behavioural (e.g. locomotion) responses, are appropriatetools to investigate sub-lethal effects and determine mechanisms of ac-tions of AgNPs in aquatic invertebrates (Lagadic, 2002; Moore et al.,2004). These biomarkers are known to be early warning indicatorsused as a diagnostic or predictive tool for long-term effects that couldoccur at individual and/or population level (Garaud et al., 2015, 2016;Jemec et al., 2008; Lagadic, 2002; Ulm et al., 2015; Vasseur andLeguille, 2004).

The aim of this study was to investigate in a multibiomarker ap-proach whether G. fossarum could serve as a good model organism inthe field of environmental nanotoxicology, allowing the identificationof potential AgNPs effects from the molecular to the organism level.Chemically synthesised non-coated particles of different initial sizes(AgNPs 20 nm and AgNPs 200 nm) were used to better understandthe link between size and toxicity. In addition, AgNPs synthesised byOcimum sanctum (initial size: AgNPs 23 nm) and Azadirachta indica (ini-tial size: AgNPs 27 nm) plant leaf extract (Balachandran et al., 2012)were used to evaluate the influence of themethod of synthesis and sur-face chemistry on the toxicity of AgNPs. Out of two different populationsof G. fossarum exposed to the selected AgNPs in a first acute experiment,the most sensitive population was then used for further experimentswith lower concentrations. Effects on survival, bioaccumulation, osmo-regulation, antioxidant responses (catalase, glutathione peroxidase,total antioxidant activity), defencemechanisms (glutathione S-transfer-ase, acid phosphatase), cellular damage (lactate dehydrogenase, lipidperoxidation, apoptosis), energy reserves (cholesterol, triglycerides,proteins) and behaviour (locomotor activity and ventilation) wereinvestigated.

2. Organisms, material and methods

2.1. Particles and chemicals

Non-coated AgNPs (20 and 200 nm) were obtained fromPlasmaChem GmbH (Berlin, Germany) and were characterised(Lankoff et al., 2012). A. indica (AgNPs 27 nm) and O. sanctum (AgNPs23 nm) plant leaf extract synthesized AgNPs were provided andcharacterised by the department of Biotechnology, Bharathiar Universi-ty, India (Balachandran et al., 2012, 2013). Silver nitrate (AgNO3,) waspurchased fromVWR (Leuven, Belgium), osmium tetroxide (OsO4), glu-taraldehyde and the epoxy resin embedding kit from Sigma Aldrich

1650 K. Mehennaoui et al. / Science of the Total Environment 566–567 (2016) 1649–1659

(Bornem, Belgium). Phosphate buffer saline (PBS) was obtained fromInvitrogen (Merelbeke, Belgium).

2.2. Particle dispersion and characterisation in Volvic® water

2.2.1. AgNPs 20 and 200 nm

Two milligrams (2 mg) of particles were weighed and dispersed in1 mL of a solution containing 5% (v/v) dimethyl sulfoxide (DMSO) inMilli-Q water. The solution was sonicated (cycle 0.5; amplitude 30%)on ice for 3 min using a UP200S probe ultra sonicator with a low diam-eter tip of 2 mm (Hielscher, Germany) resulting in 6.56 J/s of deliveredacoustic power (See Supplementary material for calculations details).Stocks were always prepared freshly before each experiment(Georgantzopoulou et al., 2013; Lankoff et al., 2012). Total dilutions ofDMSO in exposure medium were always in excess of 100.000, leadingto minute DMSO exposure.

2.2.2. AgNPs 23 nm and 27 nm

The NPs were provided as stocks in Milli-Q water and stored at 4 °Cin glass vials. Before use, the vials were gently vortexed. No further son-ication steps were included (Georgantzopoulou et al., 2013).

2.2.3. AgNPs characterisation

AgNPs 20 and 200 nm were characterised using Scanning electronmicroscopy (SEM) type DSM942 (Zeiss, Germany) in the secondaryelectron (SE) mode and transmission electron microscopy (TEM)(JEOL 1200 EXII,JEOL, Japan) (Lankoff et al., 2012).

AgNPs 23 and 27 nmwere characterised usingHigh Resolution Scan-ning Electron Microscope (HRSEM). Images were obtained using SirionHRSEM (FEI Company). Additionally, characterisation and size distribu-tion using TEM were provided (Balachandran et al., 2012). The surfacechemistry of AgNPs 23 and 27 nm were characterised using ShimadzuFTIR-8400S (Fourier Transform Infrared) spectrometer (Balachandranet al., 2012). Results were summarized in our previous study(Georgantzopoulou et al., 2013).

Particle size distribution of the AgNPs in the exposure medium(commercialmineralwater Volvic®, France)was evaluatedusingNano-particle Tracking Analysis (NanoSIGHT, Malvern Instrument Ltd., UK).

2.3. Organism sampling and acclimation

Two different G. fossarum populations were sampled at two riversconsidered to be unpolluted (Arce Funck et al., 2013; Dohet et al.,2008). The first set of animals (G.f1) was collected in June 2013

(experiment 1) at La Maix (48°28′55.7″ N and 07°04′20.1 E,Vexaincourt, Vosges Mountains, North-East France). The second set ofanimals (G.f2)was collected in July 2013 (experiment 1) and September2013 (experiment 2) at Schwaarzbaach (49°48′24.9″N and 06°04′53.2″E, Attert River, Colmar-berg, Luxembourg). Specimens were collectedusing a hand net and they were quickly transported in plastic tankswith river water to the laboratory, where they were kept at 12 °C untilsorting. In order to avoid potential gender and between-life stage influ-ences on the organism sensitivity to the treatments, only adult maleswere selected for the experiments (Arce Funck et al., 2013; Sornom etal., 2010; Sroda and Cossu-Leguille, 2011; Vellinger et al., 2013). Malesfrom precopula pairs were directly identified (males are bigger than fe-males). Other individuals were sexed based on sexual dimorphism suchas gnathopode sizes (males present bigger gnathopodes compared tofemales). As parasitism is known to interferewith the studied responses(Gismondi et al., 2012a, 2012b, 2012c), Acanthocephala sp. parasitizedorganisms (parasites appear as orange spots visible through the cuticle)were excluded from sampling. After selection, non-parasitized maleswere acclimatised for at least 10 days in commercial mineral water(Volvic®, France) under controlled conditions (13.1 ± 0.2 °C with aphotoperiod of 16 h light and 8 h darkness). Volvic® water was gradu-ally added to river water for the first three days of acclimation. Gamma-ridswere then kept for at least 10 days in 100%Volvic®water (Andreï etal., 2016). Individuals were fed ad libitum with alder leaves (Alnus sp.)up to 24 h before experiment. Volvic® water in the tanks was aeratedand changed every 24 h during the acclimation period to avoid organicmatter accumulation and potential increase of ammonium, nitrite andnitrate. All relevant characterization parameters such as mineral con-tent, pH, conductivity, dissolved oxygen, hardness and alkalinity ofVolvic® water were controlled and are presented in Supplementarymaterials (Table S1).

2.4. Experiment 1: acute toxicity test

After the acclimation period, 5 males per treatment group weretransferred into plastic tanks (250 mL polypropylene tanks) containing70 mL of exposure medium (Volvic® water with or without AgNPs orAgNO3). A piece of mesh was added in each tank in order to provide aresting surface for the organisms and to reduce losses due to cannibal-ism. Males were exposed to various treatments: AgNO3 and AgNPs23 nm (0.5–1–2–4–8 μg·L−1), AgNPs 27 nm (1–3–10–30–100 μg·L−1), AgNPs 20 nm and AgNPs 200 nm (10–30–100–300 -1000 μg·L−1), and a control for 72 h at 12 °C with a photoperiod of16 h light and 8 h darkness in the absence of feeding (Fig. 1). The

Fig. 1. Experimental design (adapted from Arce Funck et al., 2013). CAT: Catalase; GPx: Glutathione Peroxidase; TAC: Total Antioxidant Activity; ACP: Acid Phosphatase; GST: GlutathioneS-Transferase; CHOL: Cholesterol; TRIG: Triglyceride: PROT: Proteins: ETS: Electron Transport System; LDH: Lactate dehydrogenase; LPO: Lipid peroxidation; CASP-3: Caspase 3.

1651K. Mehennaoui et al. / Science of the Total Environment 566–567 (2016) 1649–1659

different tested range of concentrations were selected based on previ-ous results obtained after exposing D. magna to the same AgNPs(Georgantzopoulou et al., 2013). AgNO3 and AgNPs stock solutionswere diluted to the desired nominal concentrations in mineral water(Volvic®, France). Exposure medium was renewed daily (Volvic®water with or without AgNPs or AgNO3 at the same concentrationslisted above; no aeration). Dead animals were removed and survivorswere counted. Each treatmentwas performed in 4 replicates. Acute tox-icity of each AgNPs was assessed using LC50 values.

2.4.1. Total and dissolved Ag release

At the end of the exposure period, 5 mL of exposure medium(Volvic® water + AgNPs at the lowest and highest working concentra-tions where at least one animal survived)were kept for total silver con-centrations measurements.

In order to assess the ion release for each class of particles, 4 mL ofexposure medium (Volvic® water + AgNPs at the lowest and highestworking concentrations where at least one animal survived) were cen-trifuged at the end of the 72 h exposure period at 4000 g for 40 minusing centrifugal filter devices (Amicon ultra-4, Milli-pore, Ireland)with a 3 kDa cut-off level. Total (exposure medium) and dissolved(lower fraction) Ag were quantified using Inductively Coupled PlasmaMass Spectrometry (ICP-MS) (Elan DRC-e, Perkin-Elmer, Waltham,MA, USA) as previously described (Boscher et al., 2010;Georgantzopoulou et al., 2013). All measurements were performed inthe absence of animals. Percentages of dissolution were calculatedbased on total silver measured concentrations. The quantification limitof ICP-MS was around 0.051 μg·L−1.

2.4.2. Particle uptake in gills of G. fossarum

Gills of G. fossarum are one of the first organs that are directly ex-posed to contaminants in the water (Felten et al., 2008b; Henry andWheatly, 1992; Issartel et al., 2010; Lignot et al., 2000; Pequeux, 1995)and were therefore selected for AgNPs uptake evaluation.

At the end of the 72 h exposure period, gammarids' gills were har-vested and fixed overnight with 5% glutaraldehyde in Phosphate BufferSaline (PBS) at 4 °C. Glutaraldehyde was then removed and sampleswere washed in PBS. After that, gills were embedded in small 1% agarcubes (adapted from Georgantzopoulou et al., 2013). The gills werepost-fixed with 1% osmium tetroxide in Milli-Q water for 1 h at roomtemperature. After an additional washing step with PBS, samples weredehydrated with five increasing acetone concentrations (30, 50, 70and 100% v/v in Milli-Q water) and were embedded in epoxy resin(Epon 812 substitute). Samples were cut to 300 nm semi-thin sections(Leica ultracut UCT, LePeq Cedex, France) and finally placed on siliconwafers (Siltronix, Archamps, France) for Secondary Ion Mass Spectrom-etry (SIMS) analysis (Georgantzopoulou et al., 2013).

Sampleswere analysedwith a NanoSIMS 50 (Cameca, Gennevilliers,France) using Cs+ as primary ion source (8 KeV) sputtering the surfaceof the sample (−8 KeV) with a raster of 40 × 40 μm2 to generate sec-ondary negative ions. Images were recorded in a pixel format of256 × 256 image points with a counting time of 30ms per pixel. The in-strument was tuned for a mass resolution (M/ΔM) up to 5000 for theelimination of atomic or molecular isobar interference. The mass cali-bration of the silver ion was carried out using a silver foil (Goodfelow,Huntingdon, UK). The isotopic ratio between 107Ag− (m = 106.9051,51.8%) and 109Ag− (m = 108.9048, 48.2%) was measured and verified(ratio = 1074). The 12C14N− cluster was simultaneously detectedwith the silver ions to allow the recognition of the essential anatomicalfeatures (Eybe et al., 2009; Georgantzopoulou et al., 2013).

2.5. Experiment 2: physiological and behavioural responses

The G.f2 population, which was slightly more sensitive to AgNPs,was used for the second experiment in which the sub-lethal effects of

AgNPs that had showed a toxic effect in the first experiment (AgNPs20 nm, AgNPs 23 nm and AgNPs 27 nm) were assessed (Fig. 1).

Groups consisting of 15 individuals were housed in plastic tanks(one 500 mL polypropylene tank for each group) containing 210 mLof exposure medium (Volvic®, France). The same parameters used forexperiment 1 (exposure time, temperature and photoperiod) were ap-plied for the setup of experiment 2. Two concentrations of AgNO3 andfor each AgNPs (1 and 3 μg·L−1) were used. The group treated withonly Volvic® water represented the negative control group. The choiceof 1 and 3 μg·L−1 was based on the results obtained during the first ex-periment as they were lower than the LC5 of AgNPs 23 nm, which re-sulted as the most toxic AgNPs among those analysed in this work.Every 24 h, dead animals were removed from each tank and livingones were counted. Each treatment was performed in 3 replicates.

2.5.1. Ag bioconcentration

After 72 h of exposure, a pool of 4 gammarids per condition wererinsed with Milli-Q water and gently dried on filter paper. Animalswere weighed and stored at −20 °C until analysis. 3 mL of mass gradeMilli-Q water supplemented with 30% (v/v) of HNO3 and 14% (v/v) ofH2O2were added to each pool followed bymineralisation under a max-imum pressure of 35 bars andmaximum temperature of 200 °C in ami-crowave (Anton Paar Multiwave Pro). After mineralisation, total Agcontentwas quantified using ICP-MS following the procedure describedelsewhere (Georgantzopoulou et al., 2013). Bioconcentration factors(BCF) were calculated based on the internal Ag concentrations in gam-marids andmeasured total Ag concentration in the exposuremediumatthe end of exposure period (BCF= [Ag]organism in μg·kg−1 - [Ag]control inμg·kg−1/[Ag]water in μg·L−1).

2.5.2. Locomotor activity

For each treatment, measurements of locomotor activity, ventilationand haemolymph osmolalitywere performed on the same pool of 10 in-dividuals. At the end of the exposure period, locomotor activity wasfirstly assessed by counting the number of animals in movement in a80 mL glass tank containing 10 organisms with a piece of net added toprovide a resting surface (Felten et al., 2008b). Measurements weredone after 5 min of acclimation at the same time of day with similarlight conditions and in a quiet environment. Moving G.f2were countedfor a period of 2 s and this process was repeated 40 times.

2.5.3. Ventilatory activity

Ventilatory activity was recorded immediately after locomotor ac-tivity measurements on the same animals (G.f2), by measuring the fre-quency of pleopod beats (ventilator appendages of malacostraceancrustaceans). Ten gammarids from each treatment group were placedindividually in a glass tube containing Volvic® water and left for a 30 sacclimation period. Then, pleopod beats were visually counted threetimes for 10 s using a manual cell counter only when animals were atrest. Measurements were performed at the same period of the day toavoid possible effects of a circadian rhythm on respiration (Rosas etal., 1992).

2.5.4. Haemolymph osmolality

At the end of the exposure period and immediately after the mea-surement of locomotor and ventilator activities, the organisms wereused for the measurement of haemolymph osmolality. Prior to thehaemolymph extraction, animals (G.f2) were gently dried betweentwo pieces of filter paper to remove the excess of exposure media.Haemolymph samples were collected from the telson using a modifiedmicrocapillary (Felten et al., 2008b). A drop of haemolymph (≈20 nL)was transferred into mineral oil to avoid evaporation. Osmolality wasmeasured using an Otago nanolitre osmometer (Otago OsmometersLtd., Dunedin, New Zealand) using a 300 mOsm·kg−1 standard(Felten et al., 2008b).


2.5.5. Energy reserves, detoxification and lipid peroxidation (LPO) level

measurements

Five pools of five gammarids each (G.f2) exposed to 1 and 3 μg·L−1

of AgNO3 and AgNPs, as well as the non-treated negative control, wereweighed and frozen in liquid nitrogen and stored at−80 °Cuntil furtheranalysis. Each pool was homogenised as described previously (Srodaand Cossu-Leguille, 2011). Briefly, each pool was grindedwith amanualpotter Elvejheim tissue grinder at 4 °C in homogenisation buffer(50 mM phosphate buffer) (pH 7.6) supplemented with 1 mM L-ser-ine-borate mixture and 1 mM phenylmethylsulfonyl fluoride (PMSF)as protease inhibitors, adjusted to the weight of the pooled tissues(ratio 4/1, v/w). Homogenates were centrifuged at 250 g for 5min to re-move cuticle fragments and 112 μL of the resulting supernatant (S1)were kept for biomarker measurements. The remaining supernatantwas centrifuged at 1000 g for 20min at 4 °C. The supernatantwas recov-ered and centrifuged again at 20,000 g for 50 min at 4 °C. The final su-pernatant, corresponding to the cytosolic fraction (S2), was used forenzyme activities measurement.

Acid phosphatase (ACP), lipid peroxidation (LOOH), lactate dehy-drogenase (LDH), electron transport system (ETS) and energy reserveswere measured on the S1 fractions, while total antioxidant capacity(TAC), catalase (CAT), glutathione peroxidase (GPxtot), and glutathioneS-transferase (GST), caspase-3 (CASP-3) weremeasured on the cytosol-ic fraction (S2) by an automated colorimetricmethods developed on theKonelab 20-XTi (Thermo Scientific) (Garaud et al., 2015, 2016). All tech-nical procedures are described in detail in the Supplementary material.

2.6. Statistical analysis

LC50 values were calculated on nominal concentrations based on alogistic curve-fitting procedure applying the method described earlier(Isnard et al., 2001; Vindimian et al., 1999). The Excel macro REGTOXallowed us to estimate both LC50 values using Hill model (Vindimianet al., 1983) and their 95% confidence intervals (Vellinger et al., 2012).95% confidence intervals were considered for identifying significant dif-ferences existing between Gammarus populations.

All other results are reported as mean ± SD. Total and dissolved Agconcentrations in exposure medium were compared using two-wayANOVA, with exposure conditions and Gammarus population as factors,followed by Tukey HSD post hoc test for multiple comparisons. Thecomparison of each biomarker response in tested conditions was per-formed using one-way ANOVA (if normality and homoscedasticitywere identified using Shapiro-Wilk and Levene tests, respectively)followed by appropriate post hoc test formultiple comparisons. If signif-icant heterogeneity among tested conditions (P b 0.05) had been iden-tified, non-parametric Kruskal-Wallis test for multiple independentgroups followed by Mann-Whitney U test were performed. An arcsinesquare root transformationwas doneon responses expressed in % (mor-tality and locomotion) before applying statistical tests to normalise thedata. All statistical analyses were performed using Statistica 7.0 soft-ware (Statsoft Inc).

3. Results

3.1. Particle characterisation

Nanoparticles tracking analysis (NTA) showed that the studiedAgNPs of the nominal size of around 20 nm agglomerated in Volvic®water (Table 1, Fig. S1). Zeta (ζ) potential ranged from −3.5 to −4.5,suggesting unstable dispersion of AgNPs in Volvic® water (Table 1).

3.2. Experiment 1: acute toxicity test and particle uptake

In general, AgNO3 and AgNPs 23 nm were the most toxic for bothpopulations with AgNO3 having the lowest LC50 values followed byAgNPs 23 nm, AgNPs 27 nm and AgNPs 20 nm (Table 2). No LC50

value could be recorded for AgNPs 200 nm within the tested range ofconcentrations (10–1000 μg·L−1). The LC50 calculated for G.f2was sig-nificantly lower than the LC50 determined for G.f1 (no overlap in 95%confidence intervals) when they were exposed to AgNO3, AgNPs23 nm and AgNPs 27 nm. G.f2 was thus considered to be the slightlymore sensitive population and was therefore selected for furtherexperiments.

Uptake study, performed by NanoSIMS 50 analysis, showed no pres-ence of silver in the gill tissues of control gammarids (Fig. 2D).Internalisation of Ag within gill tissues was observed in G. fossarum ex-posed to AgNPs 23 nmand AgNPs 27 nm (fig. 2A and B). AgNO3 seem tobe present only in the external membrane of G. fossarum gills (Fig. 2C).

Dissolution of ions from AgNPs was below the limit of quantificationat the lowest exposure concentrations tested, therefore only concentra-tions from the high exposure can be reported.

The dissolved Ag concentrations measured in Volvic® water at theend of exposure period for AgNPs 23 nm were at the same level as forAgNO3 (~0.5 μg·L−1). The percentage of dissolution of AgNPs 23 nmwas about 30% (G.f2), while lower levels were found for all the otherAgNPs (between 0.3 and 4.4%) (Table 3).


3.3.1. Survival

After 72 h of exposure, survival rate in controls remained high andabove 95% (Fig. 3). Survival rates of G.f2 were significantly reduced(one-wayANOVA, F=9.470, 8 df, P b 0.0001)when organismswere ex-posed to the highest concentration (3 μg·L−1) of AgNO3, reaching 28±30% compared to the untreated control. No significant effects on surviv-al were recorded for G. fossarum exposed to AgNPs 23 nm, AgNPs 27 nmand AgNPs 20 nm (P N 0.05).

3.3.2. Ag bioconcentration

ICP-MS analysis of internal Ag+ concentrations in control gamma-rids revealed that non-treated animals contained already0.274 μg·g−1 of Ag in their body (Table. 4). 72 h of direct exposure to1 μg·L−1 of AgNO3 led to a 4-fold increase of Ag concentration in thewhole body of gammarids compared to controls.

Internal Ag concentration in G. fossarum exposed to AgNPs 20 nmand 1 μg·L−1 of AgNPs 23 nm were not different from those measuredin negative control animals. Internal Ag concentration of gammarids ex-posed to 3 μg·L−1 of AgNPs 23 nm was increased by 24 and 12-fold

Table 1

Size distribution of particles (mode± SD, 3 replicates) and ζ potential of AgNPs in Volvic®water.

Particle Size (nm) Zeta potential (mV)

AgNPs 23 nm 56 ± 3.2 −3.49AgNPs 27 nm 63 ± 3.2 −3.55AgNPs 20 nm 69 ± 8.4 −3.49AgNPs 200 nm 135 ± 18.7 −4.55

Measurements were performed in Volvic® water at 12 °C, pH 7.7. Particle concentrationused was 100 μg·L−1 (detection limit of NTA).

Table 2

LC50 values with 95% confidence intervals for G.f1 and G.f2 exposed to AgNO3 and AgNPs20 nm, AgNPs 23 nm, AgNPs 27 nm and AgNPs 200 nm for 72 h.

Compound G.f1 G.f2

LC50-72 h (μg·L−1) LC50-72 h (μg·L−1)

AgNO3 3.9 (3.3–4.3) 2.3 (2.0–2.6)AgNPs 20 nm 835 (590–988) N1000AgNPs 23 nm 7.7 (6.7–8.6) 4.9 (4.2–5.5)AgNPs 27 nm N100 5.5 (3.9–8.2)AgNPs 200 nm N1000 N1000


compared to control and animals exposed to 1 μg·L−1 of AgNPs 23, re-spectively. Ag concentration of gammarids exposed to 3 μg·L−1 ofAgNPs 27 nm were increased by 28 and 2.7-fold compared to controlanimals and those exposed to 1 μg·L−1 of AgNPs 27 nm, respectively.AgNPs 23 nm had the highest bioconcentration factor (BCF = 4238)in G.f2 followed by AgNPs 27 nm (BCF = 2382), AgNO3 (BCF = 2277)and AgNPs 20 nm (BCF = 147) (Table 4).


The basal level of haemolymph osmolality in controls G.f2 reached295 ± 13 mOsm·kg−1 (Fig. 4). AgNO3 (1 and 3 μg·L−1) and AgNPs23 nm (3 μg·L−1) significantly decreased haemolymph osmolality ofG.f2 (one-way ANOVA, F = 3.466, 8 df, P b 0.001) reaching 258 ±31mOsm·kg−1, 261 ± 24 mOsm·kg−1 and 275 ± 53 mOsm·kg−1 re-spectively. Osmolality in G. fossarum exposed to AgNPs 20 nm, 27 nmand 1 μg·L−1 of AgNPs 23 nm was not affected.

3.3.4. Energy reserves, antioxidants, defence mechanisms, cellular damage

and behavioural responses

Due to the high mortality rate recorded for G. fossarum exposed to3 μg.L−1 of AgNO3, antioxidant responses, defencemechanisms, cellulardamage and energy reserve measurements could not be performed forthis condition.

Antioxidant responses, defence mechanisms, cellular damage, ener-gy reserves (Table 5) and ventilatory activity (Fig. 5a) were not impact-ed by 1 μg·L−1 AgNO3 and 1 and 3 μg·L−1 AgNPs treatments (one-wayANOVA, F = 0.921, 8 df, P N 0.05).

Locomotion was significantly decreased by 20.3% and 33.9% (one-way ANOVA, F = 18.705, 8 df, P b 0.0001) when gammarids were ex-posed to 1 μg·L−1 and 3 μg·L−1 of AgNO3, respectively and by 17.7%at 3 μg·L−1 of AgNPs 23 nm. Locomotor activity of G. fossarum exposedto AgNPs 20 nm and AgNPs 27 nm was not affected (Fig. 5b).

4. Discussion

4.1. Experiment 1: acute toxicity test

The observed mortality rates in G. fossarum exposed to AgNO3 andAgNPs indicates that the observed effects are concentration-dependent.AgNO3 appeared to be themost toxic form of silver for both G. fossarumpopulations. The recorded LC50-72 h values were in the same rangethan those previously reported for Gammarus pulex and G. fossarum(Bury et al., 2002; Arce Funck et al., 2013). The G. fossarum populationused in this study was the same one used in Arce Funck et al. (2013)study (collected at la Maix).

Generally, the order of toxicity of all AgNPs was similar for both G.fossarum populations with AgNPs 23 nm being the most toxic onefollowed by AgNPs 27 nmand AgNPs 20 nm. Nomortality was observedfor both G. fossarum populations exposed to AgNPs 200 nm. The sameorder of toxicity of AgNPs for Vibrio fischeri, Desmodesmus subspicatus,D. magna (Georgantzopoulou et al., 2013) and in-vitro co-culturemodel for gastrointestinal epithelium (Georgantzopoulou et al., 2016)was reported. Ag 200 nm appeared the be the least toxic particle forall themodels tested andmultivariate analysis had shown that this par-ticle always clusters togetherwith control groups (Georgantzopoulou etal., 2016).

G.f2 appeared to be the slightly more sensitive of the two selected G.fossarum populations. The different LC50s obtained between the twopopulations could be linked to the presence of different cryptic species(Feckler et al., 2014;Weiss et al., 2014), particular life-history traits spe-cific to each population (e.g. size, reproduction, feeding habits, etc.)

Table 3

Total and dissolved Ag concentrations (mean± SD) and dissolution rates (mean± SD) ofAgNO3 and AgNPs after 72 h of exposure in Volvic®water.

Compound Nominal[Ag]

(μg·L−1)

Gammarus

populationTotal [Ag](μg·L−1)

Dissolved[Ag]

(μg·L−1)

Dissolution(%)

Control 0 G.f1 0.08 0 0 a

G.f2 0 0 0 a

AgNO3 2 G.f1 0.6 ± 0.2 0.5 ± 0.1 DNAG.f2 1.1 ± 0.1 0.4 ± 0.1 DNA

AgNPs 23nm

4 G.f1 1.2 ± 0.1 0.3 ± 0.1 25.1 ± 2.6b

G.f2 1.8 ± 0.2 0.5 ± 0 29.1 ± 1.0b

AgNPs 27nm

100 G.f1 61.3 ± 1.2 2.7 ± 0.1 4.4 ± 0.2a

G.f2 Na Na Na

AgNPs 20nm

300 G.f1 42.2 ± 4.1 1.8 ± 0.1 3.2 ± 1.9a

G.f2 20.4 ± 2.5 0.6 ± 0.1 0.3 ± 0.5a

AgNPs 200nm

1000 G.f1 26.7 ± 3.7 3.7 ± 0.1 1.9 ± 0.1a

G.f2 20.4 ± 2.2 0.8 ± 0.1 3.9 ± 0.4a

Different letters illustrate significant differences between different treatments (Two-wayANOVA + Tukey HSD post hoc test at P b 0.05 level of significance, n = 3), Na: NotAnalysed due to a high mortality rate, DNA= Do not apply.

Fig. 2. Elemental distribution of 12C14N− cluster and 109Ag− ion in 300 nm cuts of gills. G.fossarum were exposed to AgNPs (A) 23 nm, (B) 27 nm, (C) AgNO3 and (D) Control(Volvic® water). Scale bar is 5 μm.


(Lambrechts et al., 1997; Usseglio-Polatera et al., 2000) as well as phys-iological parameters (e.g. life cycle) (McCahon andPascoe, 1988; Alonsoet al., 2010; Vellinger et al., 2012; Arce Funck et al., 2013), samplingdates (June vs July), water quality in the field and food availability(Heugens et al., 2006).

The differences in toxicity observed for the different AgNPs (20 nmand 200 nm obtained by chemical synthesis and AgNPs 23 nm and27 nm obtained by “green” synthesis) could be partially explained bytheir different dissolution in exposuremedium. Indeed, ICP-MS analysisrevealed that AgNPs 23 nm dissolved more in exposure medium thanAgNPs 27 nm, AgNPs 20 nm and AgNPs 200 nm. These results are in ac-cordance with those reported previously after exposingD.magna to thesame particles (Georgantzopoulou, 2015). In this previous study theamount of dissolved Ag from AgNPs 23 nm was about 30% which fitsour results exactly. However, the observed effects could not only belinked to the effects of Ag ions as the observed effects for AgNPs23 nmwere lower than those observed for AgNO3with comparable dis-solved Ag concentrations in Volvic® water. The differences in effectscould be explained by different factors such as the size, surface chargeand specific molecules present on the surface of AgNPs. As previouslydescribed, AgNPs 23 nm and AgNPs 27 nm are synthetized using plantleaf extract. Therefore, bio-molecules (e.g proteins, flavonones, terpe-noids and sugars) are present on their surface (Balachandran et al.,2013; Georgantzopoulou et al., 2013). However, an additional com-pound was detected at the surface of AgNPs 27 nm (germinal methylor symmetric stretching of carbonyl group) making them more stablein exposuremedium (Georgantzopoulou et al., 2013) thereby potential-ly reducing their toxicity (Kroll et al., 2014). Uptake evaluation per-formed with NanoSIMS 50 showed the presence of Ag inside the gillsof gammarids exposed to AgNPs 23 nm and AgNPs 27 nm, whereas Agin AgNO3 exposed animals was only present in the external membraneof the gammarids' gills. Thus, it can be hypothesised that AgNPs 23 nmand AgNPs 27 nm have a different or an additional uptake route com-pared to AgNO3 and AgNPs 23 nm may readily release the ions in thecells after uptake (Georgantzopoulou, 2015).


4.2.1. Survival rates and Ag bioconcentration

This experiment showed that exposing G. fossarum to AgNPs and toAgNO3 led to a significant uptake of Ag. Increased internal concentra-tions of Ag are associated to decrease of survival rates of G.f2 withAgNO3 being the most toxic, followed by AgNPs 23 nm, AgNPs 27 nm

and AgNPs 20 nm. Additionally, AgNPs 23 nm also showed the highestpotential for bioaccumulationwith the highest bioconcentration factors,followed by AgNPs 27 nm and AgNPs 20 nm. These results are in accor-dance with previous studies that reported an accumulation of water-borne Ag in crustaceans, mainly in hepatopancreas and gills (Bury etal., 2002; Grosell et al., 2002; Arce Funck et al., 2013), which increasedthe mortality rate. However, despite a high amount of Agbioaccumulated in G. fossarum exposed to AgNPs 27 nm, this did notcause an increase inmortality rate or any other disturbances. The differ-ences in toxicity observed between the similar sized AgNPs 23 nm andAgNPs 27 nm can thus be linked to AgNPs dissolution and silver ions re-lease from AgNPs 23 nm (Georgantzopoulou, 2015). It could also besuggested that proteins released by G. fossarum in exposure mediummight increase the toxicity of AgNPs 23 nm by the formation of aneco-corona (Nasser and Lynch, 2016) due to observed differences inthe surface molecules of the native Ag particles (Georgantzopoulou etal., 2013). Indeed, it had been shown that the presence of proteins ofD.magna in exposure medium increased AgNPs uptake by D.magna ne-onates thus leading to a higher retaining dose of AgNPs and thus highertoxicity (Nasser and Lynch, 2016).

Fig. 3.G.f2 survival rates (Mean±SD) after 72 h of exposure to Ag, AgNPs 23nm, AgNPs 27nm, AgNPs 20nm. Datawere arcsine square root transformed. Letters (a–c) illustrate significantdifferences (One-way ANOVA + Fisher LSD post hoc test at P b 0.05 level of significance, n = 15).

Table 4

G.f2AgBioconcentration (Mean±SD) after 72 h exposure to AgNO3, AgNPs 20 nm, AgNPs23 nm, AgNPs 27 nm.a

Conditions Nominal[Ag]

(μg·L−1)

[Ag] inexposuremedium(μg·L−1)

Gammarusinternal [Ag]

(μg·ggammarids−1)

BCF (μg·kggammarids−1/μg·L−1)

Control 0 bLOQ 0.27 ± 0.02a NDAgNO3 1 0.36 ± 0.03 1.09 ± 0.010a 2277

AgNPs 20nm

1 0.23 ± 0.03 0.70 ± 0.003c 18693 0.68 ± 0.03 0.37 ± 0.001a 147

AgNPs 23nm

1 0.44 ± 0.05 0.54 ± 0.005a 6133 1.47 ± 0.01 6.50 ± 0.003b 4238

AgNPs 27nm

1 0.96 ± 0.04 2.83 ± 0.003a 26663 3.16 ± 0.10 7.80 ± 0.009b 2382

Different letters illustrate significant differences between different treatments (Kruskal-Wallis ANOVA + Mann-Whitney U test at P b 0.05 level of significance, n = 3). LOQ:limit of quantification.

a Due to an unexpectedmortality during sorting and acclimation period in addition to ahigh mortality rate during exposure, Ag bioconcentration in Gammarus exposed to 3μg·L−1 of AgNO3 could not be measured.



Freshwater organisms are dependent on osmoregulation. Absorptionand excretion of ions is needed to maintain their ion homeostasis. Inthis study, heamolymph osmolality of G.f2 exposed for 72 h to 1 and3 μg·L−1 of AgNO3 and 3 μg·L−1 of AgNPs 23 nm was significantly re-duced in addition to a significant higher mortality rate. This result wasin accordance with a previous study, in which a significant decrease ofheamolymph [Na+] and [Cl−], due to gill iono/osmoregulation failure,was observed in G. fossarum exposed for 96 h to 0.5 μg·L−1 of AgNO3

(Arce Funck et al., 2013). Ag+was described to act by disturbing the bran-chial ion transport pathways infish, leading to an ion influx inhibition anda significant loss of Na+ and Cl− across the gill (Morgan et al., 1997). Con-sequently, osmoregulation failure observed in gammarids exposed toAgNO3 and AgNPs 23 nm might be explained by the loss of Na+ andCl− ions through the gills. In crustaceans, Ag+was described as a compet-itor of Na+ for the same transport site, attributable to a similar radius be-tween the two ions (Grosell et al., 2002; Bianchini and Wood, 2003;Völker et al., 2013). Therefore, it could be assumed that the observedAg+ toxicity is related to the inhibition of Na+ uptake.

4.2.3. Defence mechanisms and energy reserves

In general, one of the most widely accepted toxicity mechanismsproposed for AgNPs is through the generation of ROS (Klaine et al.,2008; Vale et al., 2016), which can lead to lipid peroxidation inducing,finally, oxidative stress (Ulm et al., 2015). In this study, despite a

significant bioconcentration of Ag in gammarids exposed to AgNO3,AgNPs 23 nm and AgNPs 27 nm, and their presence within G. fossarumgills tissue, no significant alterations in antioxidant responses, defencemechanisms, cellular damage and energy reserves could be detected.These results are consistent with those reported by Arce Funck et al.(2013) who observed no effects at 0.5 and 1 μg·L−1 of AgNO3 on cata-lase and selenium-dependent GPx activities of G. fossarum. However,unlike our study, significant lipid peroxidation was observed and wasdiscussed to potentially result from an overwhelming of the defencemechanisms activities (Arce Funck et al., 2013). In our study, the ab-sence of effect could be explained by the fact that the basal levels of en-zyme activities were maybe effective enough to cope with potentialincreased ROS levels generated by the AgNPs.

4.2.4. Behavioural responses

In the current study, we could observe a significant decrease in loco-motor activity after 72 h of exposure to AgNO3 and AgNPs 23 nm (1 and3 μg·L−1) while no observable effect on locomotor activity of G.fossarum treated with AgNPs 20 nm and AgNPs 27 nm could be detect-ed. These results were in accordance with a study which showed that0.5 μg·L−1 of citrate-coated AgNPs significantly decreased locomotoractivity of G. roeseli after 72 h of exposure (Andreï et al., 2016). The ef-fects were size-dependent with the 10 nm citrate coated AgNPs beingmore potent in inhibiting locomotor activity than 60 nm citrate coatedAgNPs (Andreï et al., 2016). In our study, the results indicate that theimpact of AgNPs on behavioural responses of G. fossarummay be linked

Fig. 4. G.f2 haemolymph osmolality (Mean ± SD) after 72 h exposure to AgNO3, AgNPs 20 nm, AgNPs 23 nm and AgNPs 27 nm. Different letters illustrate significant differences betweendifferent treatments (One-way ANOVA + LSD Fisher post hoc test at P b 0.05 level of significance, n = 15).

Table 5

Mean values (±SD) of biomarkers measured in Gf2 exposed for 72 h to AgNO3, AgNPs 20 nm, AgNPs 23 nm and AgNPs 27 nm.

Biomarkers Energy reserves Defence mechanisms Antioxidant responses Cellular damage

Conditions(μg·L−1)

Prota Chola Tria ETSb ACPc GSTd GPxe TACf CATg LDHh CSP3i LOOHj

Control 0 21.7 ± 3.6 0.6 ± 0.1 1.8 ± 0.4 1.3 ± 0.1 89.7 ± 15.2 190.1 ± 31.8 68.3 ± 6.3 36.5 ± 2.2 17.5 ± 7.9 1047.4 ± 411.1 1.5 ± 0.7 1511.9 ± 350.9AgNO3 1 24 ± 6.5 0.7 ± 0.2 2 ± 1.1 1.2 ± 0.2 85 ± 7.1 156.1 ± 51.4 62.3 ± 12.2 29.9 ± 6.9 15 ± 6.1 1093.3 ± 327.5 1.2 ± 0.3 1208.2 ± 445.1AgNPs20 nm

1 22.8 ± 5.9 0.7 ± 0.2 1.9 ± 0.7 1.3 ± 0.2 87.3 ± 10.5 154 ± 58.8 61.1 ± 12.5 32.6 ± 7.5 16.2 ± 8.4 1093.4 ± 276.1 1.4 ± 0.9 1577.1 ± 6203 24.4 ± 1.4 0.8 ± 0.1 2.1 ± 0.5 1.3 ± 0.1 89.4 ± 13.7 147.5 ± 36.8 74.9 ± 2.5 33.6 ± 4.7 19.1 ± 8.5 1238 ± 167.5 1.4 ± 0.4 1308.8 ± 225.8

AgNPs23 nm

1 24.6 ± 2.6 0.7 ± 0.1 1.2 ± 0.3 1.1 ± 0.1 85 ± 7.9 154.1 ± 28.9 66.6 ± 9.3 31.1 ± 6.5 20 ± 4.6 1090.5 ± 306.6 1.3 ± 0.1 1110.6 ± 178.33 23.6 ± 4.1 0.7 ± 0.2 1.7 ± 0.5 1.2 ± 0.1 84.7 ± 3.3 146.2 ± 39.8 59.1 ± 12.3 27.9 ± 3.6 14.7 ± 5.7 963.9 ± 278.4 1.1 ± 0.3 1513.3 ± 566.5

AgNPs27 nm

1 24.1 ± 2.8 0.7 ± 0.1 1.7 ± 0.7 1.2 ± 0.1 87.8 ± 6 146.8 ± 28.1 65.8 ± 14.8 34.6 ± 9.7 19.6 ± 9.4 1056.8 ± 213.3 1.3 ± 0.4 1199.2 ± 204.33 24.6 ± 3.3 0.7 ± 0.1 2.4 ± 1.1 1.24± 0.1 82.9 ± 11.4 170.6 ± 30.5 71.7 ± 8 31.7 ± 6.3 14.6 ± 3.2 1336.1 ± 200.7 1.3 ± 0.2 1241.2 ± 156.6

Biomarkers in italic were analysed using One-way ANOVA and Tukey post hoc test, whereas the others were analysed using Kruskal-Wallis ANOVA andMann-Whitney U test. No signif-icant differences were detected for any of the exposure conditions.a:mg·g freshweight−1; b: μmol O2·g proteins

−1·h−1; c: μmol p-nitrophenol·g protein−1·h−1; d: μmol CDNB·min−1·g proteins−1; e: μmol NADPH·g protein−1·min−1; f: mmol Troloxequivalent·g protein−1; g: mmol H2O2·g proteins−1·min−1; μmol NADH·g proteins−1·h−1; h: μmolpNA·g proteins−1·h−1; j: nmol TBH·g proteins−1.


to the surface chemistry of AgNPs and eco-corona (Nasser and Lynch,2016). Indeed, authors reported that the formation of eco-corona onAgNPs affected the feeding ability of D. magna and could thereforelead to multi-trophic disturbances in aquatic ecosystems (Nasser andLynch, 2016). Besides the potential direct effect of AgNPs, the observeddecrease in locomotor activity could also be linked to an energy reallo-cation or preservation in favour of maintenance mechanisms. Otherstudies showed that the impairment of iono/osmoregulation of animalstreated with Cd or under acidic stress, led to a reduction of energy allo-cation to locomotor activity of G. pulex (Felten et al., 2008a, 2008b).Therefore, osmoregulation impairment of G.f2 exposed to AgNO3 andAgNPs 23 nm could explain the observed lower locomotion activity.Measurement of this parameter could allow the use of G. fossarum not

only at individual level, but also at population level, thus allowing a bet-ter and more realistic evaluation of the potency of a compound, whichgoes beyond the mere evaluation of the EC50s. Indeed, movement is ahighly ecologically relevant biomarker as locomotion is necessary forforaging, finding a mate and escaping from predators. Consequently,by interfering with locomotion, AgNO3 and AgNPs 23 nm could reducethe fitness of organisms and have a potential effect at population level,resulting in “ecological death” (Scott and Sloman, 2004).

5. Conclusion

This study allowed the obtaining of first information on the toxicity ofAgNPs on G. fossarum. AgNPs 23 nm, which was more toxic than AgNPs

Fig. 5. Behavioural responses of G.f2 after 72 h of exposure to AgNO3, AgNPs 20 nm, AgNPs 23 nm, AgNPs 27 nm. (A) Ventilation (Mean pleopod beat frequency ± SD). No significantdifferences were detected for any of exposure conditions (One-way ANOVA + Fischer LSD post hoc test, P b 0.05, n = 10). (B) Locomotor activities (Mean percentage of movingG.f2± SD). Letters illustrate significant differences. (One-way ANOVA + Fisher LSD post hoc test at P b 0.05 level of significance, n = 10).


27nm, appeared to be the one that releasedmore Ag ions andwas readilytaken up byG. fossarum. AgNPs 23 nm led to significant iono/osmoregula-tion impairment, decrease in locomotor activity and survival rates.Haemolymphosmolality appeared to be aneffectivemarker as it indicatesthe physiological health status of G. fossarum. Altered locomotor activity,which was the most impacted response, may indicate potential effectsof released silver ions from AgNPs 23 nm at population level.

G. fossarum is proposed in this work as model organism fornanotoxicology as it allows studies at individual levels and at populationlevel. Our results indicate the potential usefulness of G. fossarum as amodel organism for future nanotoxicology studies at population level.

Acknowledgments

This work was in part supported by the Fonds National de laRecherche (NanoGAM AFR-PhD-9229040), the ResEAU LorLux projectand the NanEau II project (C10/SR/799842). The authors are gratefulto S. Contal andD. Collard for technical support, C. Guignard and J. Ziebelfor chemical analyses, P. Grysan andA. Chauvière for NanoSIMS analysis.V. Peardon and S. Pitt are thanked for thorough proofreading and lan-guage corrections. The contribution of YL. Balachandran in the develop-ment of plant-synthesised AgNPs in gratefully acknowledged. M.Kruszewski is thanked for providing chemically synthesised AgNPs.

Appendix A. Supplementary data

Supplementary data to this article can be found online at http://dx.doi.org/10.1016/j.scitotenv.2016.06.068.

References

Ali, D., 2014. Oxidative stress-mediated apoptosis and genotoxicity induced by silvernanoparticles in freshwater snail Lymnea luteola L. Biol. Trace Elem. Res. 162,333–341. http://dx.doi.org/10.1007/s12011-014-0158-6.

Allen, H.J., Impellitteri, C.A., Macke, D.A., Heckman, J.L., Poynton, H.C., Lazorchak, J.M.,Govindaswamy, S., Roose, D.L., Nadagouda,M.N., 2010. Effects from filtration, cappingagents, and presence/absence of food on the toxicity of silver nanoparticles to Daph-nia magna. Environ. Toxicol. Chem. 29, 2742–2750. http://dx.doi.org/10.1002/etc.329.

Alonso, Á., Lange, H.J.D., Peeters, E.T.H.M., 2010. Contrasting sensitivities to toxicants ofthe freshwater amphipods Gammarus pulex and G. fossarum. Ecotoxicology 19,133–140. http://dx.doi.org/10.1007/s10646-009-0398-y.

Andreï, J., Pain-Devin, S., Felten, V., Devin, S., Giambérini, L., Mehennaoui, K., Cambier, S.,Gutleb, A.C., Guérold, F., 2016. Silver nanoparticles impact the functional role ofGammarus roeseli (Crustacea Amphipoda). Environ. Pollut. 208, 608–618. http://dx.doi.org/10.1016/j.envpol.2015.10.036 (Part B).

Arce Funck, J., Danger, M., Gismondi, E., Cossu-Leguille, C., Guérold, F., Felten, V., 2013. Be-havioural and physiological responses of Gammarus fossarum (CrustaceaAmphipoda) exposed to silver. Aquat. Toxicol. 142–143, 73–84. http://dx.doi.org/10.1016/j.aquatox.2013.07.012.

Asghari, S., Johari, S.A., Lee, J.H., Kim, Y.S., Jeon, Y.B., Choi, H.J., Moon, M.C., Yu, I.J., 2012.Toxicity of various silver nanoparticles compared to silver ions in Daphnia magna.J. Nanobiotechnology 10, 1–11. http://dx.doi.org/10.1186/1477-3155-10-14.

Ašmonaite, G., Boyer, S., de, S., Wassmur, B., Sturve, J., 2016. Behavioural toxicity assessmentof silver ions and nanoparticles on zebrafish using a locomotion profiling approach.Aquat. Toxicol. 173, 143–153. http://dx.doi.org/10.1016/j.aquatox.2016.01.013.

Balachandran, Y.L., Girija, S., Selvakumar, R., Tongpim, S., Gutleb, A.C., Suriyanarayanan, S.,2013. Differently environment stable bio-silver nanoparticles: study on their opticalenhancing and antibacterial properties. PLoS ONE 8, e77043. http://dx.doi.org/10.1371/journal.pone.0077043.

Balachandran, Y.L., Peranantham, P., Selvakumar, R., Gutleb, A.C., Girija, S., 2012. Size-con-trolled green synthesis of silver nanoparticles using dual functional plant leaf extractat room temperature. Int. J. Green Nanotechnol. 4, 310–325. http://dx.doi.org/10.1080/19430892.2012.706183.

Barnard, J., Barnard, C., 1983. Freshwater amphipoda of the world/J.L. & C.M. Barnard. -Version details [WWW Document]. Mt Vernon Va Hayfield Assoc URL http://trove.nla.gov.au/version/26012490 (accessed 4.22.16).

Batley, G.E., Kirby, J.K., McLaughlin, M.J., 2013. Fate and risks of nanomaterials in aquaticand terrestrial environments. Acc. Chem. Res. 46, 854–862. http://dx.doi.org/10.1021/ar2003368.

Besse, J.-P., Coquery, M., Lopes, C., Chaumot, A., Budzinski, H., Labadie, P., Geffard, O., 2013.Caged Gammarus fossarum (Crustacea) as a robust tool for the characterization of bio-available contamination levels in continental waters: towards the determination ofthreshold values. Water Res. 47, 650–660. http://dx.doi.org/10.1016/j.watres.2012.10.024.

Bhatt, I., Tripathi, B.N., 2011. Interaction of engineered nanoparticles with various compo-nents of the environment and possible strategies for their risk assessment.Chemosphere 82, 308–317. http://dx.doi.org/10.1016/j.chemosphere.2010.10.011.

Bianchini, A., Wood, C.M., 2003. Mechanism of acute silver toxicity in Daphnia magna. En-viron. Toxicol. Chem. 22, 1361–1367. http://dx.doi.org/10.1002/etc.5620220624.

Blaser, S.A., Scheringer, M., MacLeod, M., Hungerbühler, K., 2008. Estimation of cumula-tive aquatic exposure and risk due to silver: contribution of nano-functionalized plas-tics and textiles. Sci. Total Environ. 390, 396–409. http://dx.doi.org/10.1016/j.scitotenv.2007.10.010.

Boscher, A., Gobert, S., Guignard, C., Ziebel, J., L'Hoste, L., Gutleb, A.C., Cauchie, H.-M.,Hoffmann, L., Schmidt, G., 2010. Chemical contaminants in fish species from riversin the North of Luxembourg: potential impact on the Eurasian otter (Lutra lutra).Chemosphere 78, 785–792. http://dx.doi.org/10.1016/j.chemosphere.2009.12.024.

Buffet, P.-E., Pan, J.-F., Poirier, L., Amiard-Triquet, C., Amiard, J.-C., Gaudin, P., Faverney,C.R., Guibbolini, M., Gilliland, D., Valsami-Jones, E., Mouneyrac, C., 2013. Biochemicaland behavioural responses of the endobenthic bivalve Scrobicularia plana to silvernanoparticles in seawater and microalgal food. Ecotoxicol. Environ. Saf. 89,117–124. http://dx.doi.org/10.1016/j.ecoenv.2012.11.019.

Bundschuh, M., Zubrod, J.P., Klemm, P., Elsaesser, D., Stang, C., Schulz, R., 2013. Effects ofpeak exposure scenarios on Gammarus fossarum using field relevant pesticide mix-tures. Ecotoxicol. Environ. Saf. 95, 137–143. http://dx.doi.org/10.1016/j.ecoenv.2013.05.025.

Bury, N.R., Shaw, J., Glover, C., Hogstrand, C., 2002. Derivation of a toxicity-based model topredict how water chemistry influences silver toxicity to invertebrates. Comp.Biochem. Physiol. Part C Toxicol. Pharmacol. 133, 259–270. http://dx.doi.org/10.1016/S1532-0456(02)00096-0.

Canesi, L., Ciacci, C., Fabbri, R., Marcomini, A., Pojana, G., Gallo, G., 2012. Bivalve molluscsas a unique target group for nanoparticle toxicity. Mar. Environ. Res. 76, 16–21.http://dx.doi.org/10.1016/j.marenvres.2011.06.005 (Emerging and persistent im-pacts on Marine Organisms: Detection methods and action mechanisms).

Castranova, V., 2011. Overview of current toxicological knowledge of engineered nano-particles. J. Occup. Environ. Med. 53, S14–S17. http://dx.doi.org/10.1097/JOM.0b013e31821b1e5a.

Coulaud, R., Geffard, O., Coquillat, A., Quéau, H., Charles, S., Chaumot, A., 2014. Ecologicalmodeling for the extrapolation of ecotoxicological effects measured during in situ as-says in gammarus. Environ. Sci. Technol. 48, 6428–6436. http://dx.doi.org/10.1021/es501126g.

Coulaud, R., Geffard, O., Xuereb, B., Lacaze, E., Quéau, H., Garric, J., Charles, S., Chaumot, A.,2011. In situ feeding assay with Gammarus fossarum (Crustacea): modelling the in-fluence of confounding factors to improve water quality biomonitoring. Water Res.45, 6417–6429. http://dx.doi.org/10.1016/j.watres.2011.09.035.

Dohet, A., Ector, L., Cauchie, H.-M., Hoffmann, L., 2008. Identification of benthic inverte-brate and diatom indicator taxa that distinguish different stream types as well as de-graded from reference conditions in Luxembourg. Anim. Biol. 58, 419–472. http://dx.doi.org/10.1163/157075608X383719.

Eybe, T., Bohn, T., Audinot, J.N., Udelhoven, T., Cauchie, H.M., Migeon, H.N., Hoffmann, L.,2009. Uptake visualization of deltamethrin by NanoSIMS and acute toxicity to thewater flea Daphnia magna. Chemosphere 76, 134–140. http://dx.doi.org/10.1016/j.chemosphere.2009.02.005.

Fabrega, Luoma, S.N., Tyler, C.R., Galloway, T.S., Lead, J.R., 2011. Silver nanoparticles: Be-haviour and effects in the aquatic environment. Environ. Int. 37, 517–531. http://dx.doi.org/10.1016/j.envint.2010.10.012.

Feckler, A., Zubrod, J.P., Thielsch, A., Schwenk, K., Schulz, R., Bundschuh, M., 2014. Crypticspecies diversity: an overlooked factor in environmental management? J. Appl. Ecol.51, 958–967. http://dx.doi.org/10.1111/1365-2664.12246.

Felten, V., Charmantier, G., Charmantier-Daures, M., Aujoulat, F., Garric, J., Geffard, O.,2008a. Physiological and behavioural responses of Gammarus pulex exposed to acidstress. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol. 147, 189–197. http://dx.doi.org/10.1016/j.cbpc.2007.09.006.

Felten, V., Charmantier, G., Mons, R., Geffard, A., Rousselle, P., Coquery, M., Garric, J.,Geffard, O., 2008b. Physiological and behavioural responses ofGammarus pulex (Crus-tacea: Amphipoda) exposed to cadmium. Aquat. Toxicol. 86, 413–425. http://dx.doi.org/10.1016/j.aquatox.2007.12.002.

Foldbjerg, R., Jiang, X., Miclăuş, T., Chen, C., Autrup, H., Beer, C., 2015. Silver nanoparticles– wolves in sheep's clothing? Toxicol. Res. 4, 563–575. http://dx.doi.org/10.1039/C4TX00110A.

Forrow, D.M., Maltby, L., 2000. Toward a mechanistic understanding of contaminant-in-duced changes in detritus processing in streams: Direct and indirect effects on detriti-vore feeding. Environ. Toxicol. Chem. 19, 2100–2106. http://dx.doi.org/10.1002/etc.5620190820.

Garaud, M., Auffan, M., Devin, S., Felten, V., Pagnout, C., Pain-Devin, S., Proux, O., Rodius, F.,Sohm, B., Giamberini, L., 2016. Integrated assessment of ceria nanoparticle impacts onthe freshwater bivalve Dreissena polymorpha. Nanotoxicology 10, 935–944. http://dx.doi.org/10.3109/17435390.2016.1146363.

Garaud, M., Trapp, J., Devin, S., Cossu-Leguille, C., Pain-Devin, S., Felten, V.,Giamberini, L., 2015. Multibiomarker assessment of cerium dioxide nanoparticle(nCeO2) sublethal effects on two freshwater invertebrates, Dreissenapolymorpha and Gammarus roeseli. Aquat. Toxicol. 158, 63–74. http://dx.doi.org/10.1016/j.aquatox.2014.11.004.

Georgantzopoulou, A., 2015. Effects of silver nanoparticles and ions and interaction withfirst line of defense. Wageningen University, Wageningen Netherlands.

Georgantzopoulou, A., Balachandran, Y.L., Rosenkranz, P., Dusinska, M., Lankoff, A.,Wojewodzka, M., Kruszewski, M., Guignard, C., Audinot, J.-N., Girija, S., Hoffmann,L., Gutleb, A.C., 2013. Ag nanoparticles: size- and surface-dependent effects onmodel aquatic organisms and uptake evaluation with NanoSIMS. Nanotoxicology1–11 http://dx.doi.org/10.3109/17435390.2012.715312.


doi:10.1016/j.scitotenv.2016.06.068

doi:10.1016/j.scitotenv.2016.06.068

http://dx.doi.org/10.1007/s12011-014-0158-6

http://dx.doi.org/10.1002/etc.329

http://dx.doi.org/10.1007/s10646-009-0398-y

http://dx.doi.org/10.1016/j.envpol.2015.10.036

http://dx.doi.org/10.1016/j.aquatox.2013.07.012

http://dx.doi.org/10.1186/1477-3155-10-14


http://dx.doi.org/10.1371/journal.pone.0077043


http://dx.doi.org/10.1080/19430892.2012.706183

http://dx.doi.org/10.1080/19430892.2012.706183

http://trove.nla.gov.au/version/26012490

http://trove.nla.gov.au/version/26012490

http://dx.doi.org/10.1021/ar2003368

http://dx.doi.org/10.1021/ar2003368

http://dx.doi.org/10.1016/j.watres.2012.10.024


http://dx.doi.org/10.1016/j.chemosphere.2010.10.011





http://dx.doi.org/10.1016/j.ecoenv.2012.11.019



http://dx.doi.org/10.1016/S1532-0456(02)00096-0

http://dx.doi.org/10.1016/S1532-0456(02)00096-0

http://dx.doi.org/10.1016/j.marenvres.2011.06.005

http://dx.doi.org/10.1097/JOM.0b013e31821b1e5a

http://dx.doi.org/10.1097/JOM.0b013e31821b1e5a

http://dx.doi.org/10.1021/es501126g

http://dx.doi.org/10.1021/es501126g


http://dx.doi.org/10.1163/157075608X383719



http://dx.doi.org/10.1016/j.envint.2010.10.012

http://dx.doi.org/10.1111/1365-2664.12246

http://dx.doi.org/10.1016/j.cbpc.2007.09.006


http://dx.doi.org/10.1039/C4TX00110A

http://dx.doi.org/10.1039/C4TX00110A



http://dx.doi.org/10.3109/17435390.2016.1146363


http://refhub.elsevier.com/S0048-9697(16)31241-4/rf0170


http://dx.doi.org/10.3109/17435390.2012.715312

Georgantzopoulou, A., Serchi, T., Cambier, S., Leclercq, C.C., Renaut, J., Shao, J., Kruszewski,M., Lentzen, E., Grysan, P., Eswara, S., Audinot, J.-N., Contal, S., Ziebel, J., Guignard, C.,Hoffmann, L., Murk, A.J., Gutleb, A.C., 2016. Effects of silver nanoparticles and ions ona co-culture model for the gastrointestinal epithelium. Part. Fibre Toxicol. 13, 9.http://dx.doi.org/10.1186/s12989-016-0117-s9.

Gismondi, E., Beisel, J.-N., Cossu-Leguille, C., 2012a. Influence of gender and season on re-duced glutathione concentration and energy reserves of Gammarus roeseli. Environ.Res. 118, 47–52. http://dx.doi.org/10.1016/j.envres.2012.06.004.

Gismondi, E., Cossu-Leguille, C., Beisel, J.-N., 2012b. Acanthocephalan parasites: help orburden in gammarid amphipods exposed to cadmium? Ecotoxicology 21,1188–1193. http://dx.doi.org/10.1007/s10646-012-0873-8.

Gismondi, E., Rigaud, T., Beisel, J.-N., Cossu-Leguille, C., 2012c. Microsporidia parasites dis-rupt the responses to cadmium exposure in a gammarid. Environ. Pollut. 160, 17–23.http://dx.doi.org/10.1016/j.envpol.2011.09.021.

Gottschalk, F., Sun, T., Nowack, B., 2013. Environmental concentrations of engineerednanomaterials: Review of modeling and analytical studies. Environ. Pollut. 181,287–300. http://dx.doi.org/10.1016/j.envpol.2013.06.003.

Griffitt, R.J., Lavelle, C.M., Kane, A.S., Denslow, N.D., Barber, D.S., 2013. Chronicnanoparticulate silver exposure results in tissue accumulation and transcriptomicchanges in zebrafish. Aquat. Toxicol. 130–131, 192–200. http://dx.doi.org/10.1016/j.aquatox.2013.01.010.

Grosell, M., Nielsen, C., Bianchini, A., 2002. Sodium turnover rate determines sensitivity toacute copper and silver exposure in freshwater animals. Comp. Biochem. Physiol. PartC Toxicol. Pharmacol. 133, 287–303. http://dx.doi.org/10.1016/S1532-0456(02)00085-6.

Henry, R.P., Wheatly, M.G., 1992. Interaction of respiration, ion regulation, and acid-basebalance in the everyday life of aquatic crustaceans. Integr. Comp. Biol. 32, 407–416.http://dx.doi.org/10.1093/icb/32.3.407.

Heugens, E.H.W., Tokkie, L.T.B., Kraak, M.H.S., Hendriks, A.J., van Straalen, N.M., Admiraal,W., 2006. Population growth of Daphnia magna under multiple stress conditions:joint effects of temperature, food, and cadmium. Environ. Toxicol. Chem. 25,1399–1407. http://dx.doi.org/10.1897/05-294R.1.

Isnard, P., Flammarion, P., Roman, G., Babut, M., Bastien, P., Bintein, S., Esserméant, L.,Férard, J.F., Gallotti-Schmitt, S., Saouter, E., Saroli, M., Thiébaud, H., Tomassone, R.,Vindimian, E., 2001. Statistical analysis of regulatory ecotoxicity tests. Chemosphere45, 659–669. http://dx.doi.org/10.1016/S0045-6535(00)00600-7.

Issartel, J., Boulo, V., Wallon, S., Geffard, O., Charmantier, G., 2010. Cellular and molecularosmoregulatory responses to cadmium exposure in Gammarus fossarum (Crustacea,Amphipoda). Chemosphere 81, 701–710. http://dx.doi.org/10.1016/j.chemosphere.2010.07.063.

Issartel, J., Hervant, F., Voituron, Y., Renault, D., Vernon, P., 2005. Behavioural, ventilatoryand respiratory responses of epigean and hypogean crustaceans to different temper-atures. Comp. Biochem. Physiol. A Mol. Integr. Physiol. 141, 1–7. http://dx.doi.org/10.1016/j.cbpb.2005.02.013.

Jemec, A., Tišler, T., Drobne, D., Sepčić, K., Jamnik, P., Roš, M., 2008. Biochemical bio-markers in chronically metal-stressed daphnids. Comp. Biochem. Physiol. Part CToxicol. Pharmacol. 147, 61–68. http://dx.doi.org/10.1016/j.cbpc.2007.07.006.

Jung, Y.-J., Kim, K.-T., Kim, J.Y., Yang, S.-Y., Lee, B.-G., Kim, S.D., 2014. Bioconcentration anddistribution of silver nanoparticles in Japanese medaka (Oryzias latipes). J. Hazard.Mater. 267, 206–213. http://dx.doi.org/10.1016/j.jhazmat.2013.12.061.

Kelly, D.W., Dick, J.T.A., Montgomery, W.I., 2002. The functional role ofGammarus(Crustacea, Amphipoda): shredders, predators, or both? Hydrobiologia485, 199–203. http://dx.doi.org/10.1023/A:1021370405349.

Klaine, S.J., Alvarez, P.J.J., Batley, G.E., Fernandes, T.F., Handy, R.D., Lyon, D.Y., Mahendra, S.,McLaughlin, M.J., Lead, J.R., 2008. Nanomaterials in the environment: behavior, fate,bioavailability, and effects. Environ. Toxicol. Chem. 27, 1825–1851. http://dx.doi.org/10.1897/08-090.1.

Kroll, A., Behra, R., Kaegi, R., Sigg, L., 2014. Extracellular Polymeric Substances (EPS) ofFreshwater Biofilms Stabilize and Modify CeO 2 and Ag Nanoparticles. PLoS ONE 9,e110709. http://dx.doi.org/10.1371/journal.pone.0110709.

Kunz, P.Y., Kienle, C., Gerhardt, A., 2010. Gammarus spp. in Aquatic Ecotoxicology andWater Quality Assessment: Toward Integrated Multilevel Tests. In: Whitacre,D.M. (Ed.), Reviews of Environmental Contamination and Toxicology Volume205. Reviews of Environmental Contamination and Toxicology, Springer NewYork, pp. 1–76.

Kwok, K.W.H., Auffan, M., Badireddy, A.R., Nelson, C.M., Wiesner, M.R., Chilkoti, A., Liu, J.,Marinakos, S.M., Hinton, D.E., 2012. Uptake of silver nanoparticles and toxicity toearly life stages of Japanese medaka (Oryzias latipes): Effect of coating materials.Aquat. Toxicol. 120-121, 59–66. http://dx.doi.org/10.1016/j.aquatox.2012.04.012.

Lagadic, L., 2002. Biomarkers: Useful tools for the monitoring of aquatic environments.Rev. Med. Vet. 153, 581–588.

Lambrechts, M.M., Blondel, J., Hurtrez-Bousses, S., Maistre, M., Perret, P., 1997. Adaptiveinter-population differences in blue tit life-history traits on Corsica. Evol. Ecol. 11,599–612. http://dx.doi.org/10.1007/s10682-997-1515-0.

Lankoff, A., Sandberg, W.J., Wegierek-Ciuk, A., Lisowska, H., Refsnes, M., Sartowska, B.,Schwarze, P.E., Meczynska-Wielgosz, S., Wojewodzka, M., Kruszewski, M., 2012.The effect of agglomeration state of silver and titanium dioxide nanoparticles on cel-lular response of HepG2, A549 and THP-1 cells. Toxicol. Lett. 208, 197–213. http://dx.doi.org/10.1016/j.toxlet.2011.11.006.

Lignot, J.-H., Spanings-Pierrot, C., Charmantier, G., 2000. Osmoregulatory capacity as a tool inmonitoring the physiological condition and the effect of stress in crustaceans. Aquacul-ture 191, 209–245. http://dx.doi.org/10.1016/S0044-8486(00)00429-4.

MacNeil, C., Dick, J.T.A., Elwood, R.W., 1997. The trophic ecology of freshwater GammarusSpp. (crustacea:amphipoda): problems and perspectives concerning the functionalfeeding group concept. Biol. Rev. 72, 349–364. http://dx.doi.org/10.1111/j.1469-185X.1997.tb00017.x.

McCahon, C.P., Pascoe, D., 1988. Increased sensitivity to cadmium of the freshwater am-phipod Gammarus pulex (L.) during the reproductive period. Aquat. Toxicol. 13,183–193. http://dx.doi.org/10.1016/0166-445X(88)90051-3.

Moore, M.N., Depledge, M.H., Readman, J.W., Paul Leonard, D.R., 2004. An integrated bio-marker-based strategy for ecotoxicological evaluation of risk in environmental man-agement. Mutat. Res. Mol. Mech. Mutagen. 552, 247–268. http://dx.doi.org/10.1016/j.mrfmmm.2004.06.028 (Genotoxicological Studies in Aquatic Organisms).

Morgan, I.J., Henry, R.P., Wood, C.M., 1997. The mechanism of acute silver nitrate toxicity infreshwater rainbow trout (Oncorhynchusmykiss) is inhibition of gill Na+ and Cl−1 trans-port. Aquat. Toxicol. 38, 145–163. http://dx.doi.org/10.1016/S0166-445X(96)00835-1.

Nasser, F., Lynch, I., 2016. Secreted protein eco-corona mediates uptake and impacts ofpolystyrene nanoparticles on Daphnia magna. J. Proteome http://dx.doi.org/10.1016/j.jprot.2015.09.005.

Pequeux, A., 1995. Osmotic regulation in crustaceans. J. Crustac. Biol. 15, 1–60. http://dx.doi.org/10.1163/193724095X00578.

Ringwood, A.H., McCarthy, M., Bates, T.C., Carroll, D.L., 2010. The effects of silver nanopar-ticles on oyster embryos. Mar. Environ. Res. Supplement 1, S49–S51. http://dx.doi.org/10.1016/j.marenvres.2009.10.011 (PRIMO 15 Fifteenth International Symposiumon Pollutant Responses in Marine Organisms (PRIMO15) 69).

Rosas, C., Sanchez, A., Escobar, E., Soto, L., Bolongaro-Crevenna, A., 1992. Daily variationsof oxygen consumption and glucose hemolymph level related tomorphophysiological and ecological adaptations of crustacea. Comp. Biochem. Phys-iol. A Physiol. 101, 323–328. http://dx.doi.org/10.1016/0300-9629(92)90540-7.

Scott, G.R., Sloman, K.A., 2004. The effects of environmental pollutants on complex fishbehaviour: integrating behavioural and physiological indicators of toxicity. Aquat.Toxicol. 68, 369–392. http://dx.doi.org/10.1016/j.aquatox.2004.03.016.

Sharma, V.K., Siskova, K.M., Zboril, R., Gardea-Torresdey, J.L., 2014. Organic-coated silvernanoparticles in biological and environmental conditions: fate, stability and toxicity.Adv. Colloid Interf. Sci. 204, 15–34. http://dx.doi.org/10.1016/j.cis.2013.12.002.

Sornom, P., Felten, V., Médoc, V., Sroda, S., Rousselle, P., Beisel, J.-N., 2010. Effect of genderon physiological and behavioural responses of Gammarus roeseli (CrustaceaAmphipoda) to salinity and temperature. Environ. Pollut. 158, 1288–1295. http://dx.doi.org/10.1016/j.envpol.2010.01.022.

Sroda, S., Cossu-Leguille, C., 2011. Effects of sublethal copper exposure on two gammaridspecies: which is the best competitor? Ecotoxicology 20, 264–273. http://dx.doi.org/10.1007/s10646-010-0578-9.

Sun, T.Y., Gottschalk, F., Hungerbühler, K., Nowack, B., 2014. Comprehensive probabilisticmodelling of environmental emissions of engineered nanomaterials. Environ. Pollut.185, 69–76. http://dx.doi.org/10.1016/j.envpol.2013.10.004.

Ulm, L., Krivohlavek, A., Jurašin, D., Ljubojević, M., Šinko, G., Crnković, T., Žuntar, I., Šikić, S.,Vrček, I.V., 2015. Response of biochemical biomarkers in the aquatic crustaceanDaph-nia magna exposed to silver nanoparticles. Environ. Sci. Pollut. Res. 1–10. http://dx.doi.org/10.1007/s11356-015-5201-4.

US Geological Survey, 2016. Mineral Commodity Summaries 2016 - mcs-2016-silve.pdf[WWW Document]. URL http://minerals.usgs.gov/minerals/pubs/commodity/silver/mcs-2016-silve.pdf (accessed 3.7.16).

Usseglio-Polatera, P., Bournaud, M., Richoux, P., Tachet, H., 2000. Biological and ecologicaltraits of benthic freshwater macroinvertebrates: relationships and definition ofgroups with similar traits. Freshw. Biol. 43, 175–205. http://dx.doi.org/10.1046/j.1365-2427.2000.00535.x.

Vale, G., Mehennaoui, K., Cambier, S., Libralato, G., Jomini, S., Domingos, R.F., 2016.Manufactured nanoparticles in the aquatic environment–Biochemical responses onfreshwater organisms: a critical overview. Aquat. Toxicol. http://dx.doi.org/10.1016/j.aquatox.2015.11.019.

Vance, M.E., Kuiken, T., Vejerano, E.P., McGinnis, S.P., Hochella, M.F., Rejeski, D., Hull, M.S.,2015. Nanotechnology in the real world: Redeveloping the nanomaterial consumerproducts inventory. Beilstein J. Nanotechnol. 6, 1769–1780. http://dx.doi.org/10.3762/bjnano.6.181.

Vasseur, P., Leguille, C., 2004. Defense systems of benthic invertebrates in response to envi-ronmental stressors. Environ. Toxicol. 19, 433–436. http://dx.doi.org/10.1002/tox.20024.

Vellinger, C., Gismondi, E., Felten, V., Rousselle, P., Mehennaoui, K., Parant, M., Usseglio-Polatera, P., 2013. Single and combined effects of cadmium and arsenate in Gammaruspulex (Crustacea, Amphipoda): understanding the links between physiological and be-havioural responses. Aquat. Toxicol. http://dx.doi.org/10.1016/j.aquatox.2013.05.010.

Vellinger, C., Parant, M., Rousselle, P., Immel, F., Wagner, P., Usseglio-Polatera, P., 2012.Comparison of arsenate and cadmium toxicity in a freshwater amphipod (Gammaruspulex). Environ. Pollut. 160, 66–73. http://dx.doi.org/10.1016/j.envpol.2011.09.002.

Vindimian, É., Garric, J., Flammarion, P., Thybaud, É., Babut, M., 1999. An index of effluentaquatic toxicity designed by partial least squares regression, using acute and chronictests and expert judgements. Environ. Toxicol. Chem. 18, 2386–2391. http://dx.doi.org/10.1002/etc.5620181037.

Vindimian, E., Robaut, C., Fillion, G., 1983. A method for cooperative or noncooperativebinding studies using nonlinear regression analysis on a microcomputer. J. Appl.Biochem. 5, 261–268.

Völker, C., Oetken, M., Oehlmann, J., 2013. The biological effects and possible modes of ac-tion of nanosilver. In: Whitacre, D.M. (Ed.), Reviews of Environmental Contaminationand Toxicology Volume 223. Springer, New York, pp. 81–106.

Weiss, M., Macher, J.N., Seefeldt, M.A., Leese, F., 2014. Molecular evidence for furtheroverlooked species within the Gammarus fossarum complex (Crustacea: Amphipoda).Hydrobiologia 721, 165–184. http://dx.doi.org/10.1007/s10750-013-1658-7.

Xuereb, B., Bezin, L., Chaumot, A., Budzinski, H., Augagneur, S., Tutundjian, R., Garric, J.,Geffard, O., 2011. Vitellogenin-like gene expression in freshwater amphipodGammarus fossarum (Koch, 1835): functional characterization in females and poten-tial for use as an endocrine disruption biomarker in males. Ecotoxicology 20,1286–1299. http://dx.doi.org/10.1007/s10646-011-0685-2.


http://dx.doi.org/10.1186/s12989-016-0117-s9

http://dx.doi.org/10.1016/j.envres.2012.06.004

http://dx.doi.org/10.1007/s10646-012-0873-8





http://dx.doi.org/10.1016/S1532-0456(02)00085-6

http://dx.doi.org/10.1093/icb/32.3.407

http://dx.doi.org/10.1897/05-294R.1

http://dx.doi.org/10.1016/S0045-6535(00)00600-7



http://dx.doi.org/10.1016/j.cbpb.2005.02.013

http://dx.doi.org/10.1016/j.cbpb.2005.02.013

http://dx.doi.org/10.1016/j.cbpc.2007.07.006

http://dx.doi.org/10.1016/j.jhazmat.2013.12.061

http://dx.doi.org/10.1023/A:1021370405349

http://dx.doi.org/10.1897/08-090.1










http://dx.doi.org/10.1007/s10682-997-1515-0

http://dx.doi.org/10.1016/j.toxlet.2011.11.006

http://dx.doi.org/10.1016/S0044-8486(00)00429-4

http://dx.doi.org/10.1111/j.1469-185X.1997.tb00017.x

http://dx.doi.org/10.1111/j.1469-185X.1997.tb00017.x

http://dx.doi.org/10.1016/0166-445X(88)90051-3

http://dx.doi.org/10.1016/j.mrfmmm.2004.06.028

http://dx.doi.org/10.1016/j.mrfmmm.2004.06.028

http://dx.doi.org/10.1016/j.jprot.2015.09.005

http://dx.doi.org/10.1016/j.jprot.2015.09.005

http://dx.doi.org/10.1163/193724095X00578

http://dx.doi.org/10.1016/j.marenvres.2009.10.011

http://dx.doi.org/10.1016/0300-9629(92)90540-7


http://dx.doi.org/10.1016/j.cis.2013.12.002


http://dx.doi.org/10.1007/s10646-010-0578-9


http://dx.doi.org/10.1007/s11356-015-5201-4

http://minerals.usgs.gov/minerals/pubs/commodity/silver/mcs-2016-silve.pdf

http://minerals.usgs.gov/minerals/pubs/commodity/silver/mcs-2016-silve.pdf

http://dx.doi.org/10.1046/j.1365-2427.2000.00535.x

http://dx.doi.org/10.1046/j.1365-2427.2000.00535.x



http://dx.doi.org/10.3762/bjnano.6.181

http://dx.doi.org/10.3762/bjnano.6.181

http://dx.doi.org/10.1002/tox.20024










http://dx.doi.org/10.1007/s10750-013-1658-7

http://dx.doi.org/10.1007/s10646-011-0685-2


91

COMPLEMENT 1

Science of the Total Environment 566, 1649-1659.

Gammarus fossarum (Crustacea, Amphipoda) as a model organism to study silver nanoparticles

effects

Kahina Mehennaoui a,b, Anastasia Georgantzopoulou a,d, Vincent Felten b, Jennifer Andreï b, Maël

Garaud b, Sébastien Cambier a, Tommaso Serchi a, Sandrine Pain-Devin b, François Guérold b, Jean-

Nicolas Audinot c, Laure Giambérini b, Arno C. Gutleb a *


Technology (LIST), 5 avenue des Hauts-Forneaux, Esch-sur-Alzette, Luxembourg.

b Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360,

Université de Lorraine – Metz, France.

c Materials Research and Technology (MRT) Department, Luxembourg Institute of Science and

Technology, (LIST), 5 avenue des Hauts-Forneaux, Esch-sur-Alzette, Luxembourg.

d Norwegian Institute for Water Research (NIVA), Gaustadalléen 21, NO-0349 Oslo, Norway.


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SUPPLEMENTARY MATERIAL

Particle dispersion and characterisation in Volvic® water

Sonication calorimetric calibration

The delivered acoustic power of the UP200S probe ultra sonicator was determined. Temperature (K°) of

medium during sonication was measured for 30 minutes and recorded as a function of time. Then, the

delivered acoustic power P (W= J/s) was calculated as the following:

P = (dT/dt)MCp

Where (dT/dt) is the slope of temperature (K) vs time (s), M is the mass of water and Cp is the specific heat of

water (4.186J/gK°).

Energy reserves, detoxification and LPO level measurements

Detoxification mechanisms and antioxidant responses

TAC was measured based on a method previously published (Erel, 2004). The loss of colour of the radical 2,2'-

azino-bis(3-ethylbenzothiazoline-6-sulphonic acid) (ABTS*+) upon reduction by antioxidants in the cytosolic

fraction was measured by measuring absorbance at 660nm after a 342s incubation time. TAC is calculated

against a calibration curve designed using Trolox (6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid)

as standard, and expressed in mmol Trolox equivalent. g protein-1 (mmolTeq.gprot-1).

GPxtot activity was measured based on a method (Paglia and Valentine, 1967) adapted for its automation on

the Konelab (Garaud et al., 2015). The cytosolic fraction was diluted to 1/32 (v/v) to a total volume of 160 µL

with a solution containing 50 mM phosphate buffer (pH = 7.6), reduced glutathione (GSH, 2 mM), NADPH

(0.36 mM), glutathione reductase (1 U/mL) and cumene hydroperoxyde (1.125 mM), being the last substrate

for the reduction. NADPH consumption was followed for 126s by monitoring absorbance at 340 nm and

GPxtot activities were expressed in µmol NADPH.g protein-1.min-1 (ε NADPH = 6220 M-1.cm-1).

GST activity was measured on the Konelab automat by monitoring the absorbance at 340 nm for 4 min. The

measurements were performed in a phosphate buffer (0.1 M; pH 6.5) supplemented with 1-chloro2,4-

dinitobenzene (CDNB; 1 mM), GSH (1 mM) and 13/200 (v/v) of diluted cytosolic fraction. GST activity is

expressed in µmol CDNB.min-1.g-1 proteins.

ACP activity was measured by measuring the hydrolysis of 4-nitrophenyl phosphate (1.67 g/L) by acid

phosphatase. Total homogenate was diluted (1/144 v/v) in a total volume of 90 µL of citrate buffer (41.7 mM,

pH=4.8) supplemented with 0.16% Triton X-100. Reaction was stopped and the p-nitrophenol resulting from


93

the hydrolysis of 4-nitrophenyl was revealed after 15 min by the addition of 125 µL NaOH (138.7 mM). A p-

nitrophenol standard calibration curve was constructed. ACP activity was expressed in µmol p-nitrophenol. g

protein-1.h-1 (Garaud et al., 2015).

Catalase activity was measured by spectrophotometry (SPECORD® 205; analytikjena) according to a published

method (Beers and Sizer, 1951). Catalase catalyses the decomposition of hydrogen peroxide into water and

oxygen. Briefly, cytosolic fraction (S2) was first diluted 1 to 4 in 50 mM phosphate buffer solution (pH=7.6).

Then, the diluted S2 was incubated for 30 seconds with H2O2 (30 mM) in the reactive mixture (1/10 v/v) and

the consumption of H2O2 was followed at 240 nm. Catalase enzymatic activity is expressed in mmol H2O2.g

proteins-1.min-1 (ε H2O2 = 40 M-1.cm-1).

Cellular damage and oxidative stress assessment

Lipid hydroperoxide (LOOH) concentration was measured using the Konelab following the automated method

previously described (Arab and Steghens, 2004). LOOH were quantified based on the oxidation of Fe2+ to Fe3+,

under acidic conditions, followed by a complexation of Fe3+ by xynelol orange whose absorbance is finally

measured at 620 nm after 20 min incubation time. The final xylenol orange (XO) concentration used was 240

mM as described (Garaud et al., 2015). A tert-butylhydroperoxide (TBH) standard calibration curve (0.125-4

µmol/L) was established and LOOH contents were expressed in nmol TBH. g proteins-1.

Lactate Dehydrogenase (LDH) is an enzyme involved in glycogen metabolism and is used as a biomarker of

metabolic disruption. LDH activity was measured using Thermo-Scientifc Konelab ready-to-use reagents and

methods (LDH IFCC) adapted to gammarids (Garaud et al., 2015). LDH activity was expressed µmol NADH. g

proteins-1.h-1.

Apoptosis was measured based on Caspase-3 (CASP-3) activity. CASP-3 was measured in the cytosolic fraction

following an automated method developed on the Konelab. Cytosolic fraction was diluted 1 to 7 (v/v) with a

solution containing Tris buffer (10 mM, pH=7.4), DTT (5 mM) and EDTA (2 mM). The absorbance of p-

Nitroaniline (pNA) released by the cleavage of caspase—3–substrate z-DEVD-pNA (25 µM) by CASP-3 was

followed at 405 nm for 1h. CASP-3 activity was expressed in µmolpNA. g proteins-1.h-1 (µmolpNa/gprot/h,

εpNA=10500 M-1.cm-1 at 405 nm) (Garaud et al., 2015).

Energy reserves

Protein, triglyceride and cholesterol concentrations were measured using Thermo-Scientifc Konelab ready-

to-use reagents (following manufacturer’s instructions) and methods adapted to gammarid (Garaud et al.,

2015). Scal bovine-based serum (Thermo-Scientific) was used as standard to create calibration curves and

energetic reserve contents were expressed in mg.g-1 fresh weight.


94

Total protein concentration in the cytosolic fraction was determined by the pyrogallol red colorimetric

method using ready-to-use U/CSF protein (following manufacturer’s instructions) reagents and Scal bovine-

based serum as reference (Thermo-Scientific).

Electron Transport System (ETS) mitochondrial activity was measured following the original method (Owens

and King, 1975) modified by (De Coen and Janssen, 1997) and automated on the Konelab (Garaud et al.,

2015). ETS activity was measured by monitoring for 12 min at 480 nm, the production of formazan by the

reduction of p-iodonitrotetrazolium (133 mg. L-1), which works as final electron acceptor of the mitochondrial

respiratory chain. This reaction was measured in 80 mM phosphate buffer (pH=8.5) containing MgSO4 (15

µM), Triton X-100 (0.16%), PVP (300 mg. L-1), NADH (1 mM) and NADPH (0.15 mM) with 1/35 (v/v) of diluted

whole homogenate. ETS activity was calculated according to (De Coen and Janssen, 1997) and expressed in

µmol O2. g proteins-1.h-1.

REFERENCES

Arab, K., Steghens, J.-P., 2004. Plasma lipid hydroperoxides measurement by an automated xylenol orange

method. Anal. Biochem. 325, 158–163. doi:10.1016/j.ab.2003.10.022

Beers, R.F., Sizer, I.W., 1951. A Spectrophotometric Method for Measuring the Breakdown of Hydrogen Peroxide

by Catalase. J. Biol. Chem. 195, 133–140.

De Coen, W.M.D., Janssen, C.R., 1997. The use of biomarkers in Daphnia magna toxicity testing. IV. Cellular Energy

Allocation: a new methodology to assess the energy budget of toxicant-stressed Daphnia populations. J. Aquat.

Ecosyst. Stress Recovery 6, 43–55. doi:10.1023/A:1008228517955

Erel, O., 2004. A novel automated direct measurement method for total antioxidant capacity using a new

generation, more stable ABTS radical cation. Clin. Biochem. 37, 277–285. doi:10.1016/j.clinbiochem.2003.11.015

Garaud, Auffan, M., Devin, S., Felten, V., Pagnout, C., Pain-Devin, S., Proux, O., Rodius, F., Sohm, B., Giambérini, L.,

2015. Fate and integrated assessment of ceria nanoparticle impacts on the freshwater bivalve Dreissena

polymorpha: a mesocosm approach. Nanotoxicology.

Owens, T.G., King, F.D., 1975. The measurement of respiratory electron-transport-system activity in marine

zooplankton. Mar. Biol. 30, 27–36. doi:10.1007/BF00393750

Paglia, D.E., Valentine, W.N., 1967. Studies on the quantitative and qualitative characterization of erythrocyte

glutathione peroxidase. J. Lab. Clin. Med. 70, 158–169.


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TABLE S 1. PHYSICO-CHEMICAL PARAMETERS OF VOLVIC® WATER

Mineral content and physic chemical Parameters Volvic water

Ca2+ (mg.L-1) 11.5

Mg2+ (mg.L-1) 8

Na+ (mg.L-1) 11.6

K+ (mg.L-1) 6.2

SO42- (mg.L-1) 8.1

HCO3-(mg.L-1) 71

NO3- (mg.L-1) 6.3

Cl- (mg.L-1) 13.5

SiO2 (mg.L-1) 31.7

F (mg.L-1) 0.22

pH 7.75 ± 0.07

Conductivity (µS.cm-1) 213.67 ± 1.53

Dissolved oxygen (mg.L-1) 8.31 ± 0.02

Alkalinity (meq.L-1) 1.214 ± 0.1

Hardness (mg CaCO3.L-1) 63.3


96

FIGURE S 1. SIZE DISTRIBUTION OF AG NPS 23 NM (A), 27 NM (B), 20 NM (C) AND 200 NM (D) IN VOLVIC® WATER

EXPRESSED AS PARTICLE CONCENTRATION X 106.ML-1. THE RED ERROR BARS INDICATE THE ± SD OF THE MEAN OF TRIPLICATE

MEASUREMENTS.

Chapter 3 Reference genes of G. fossarum

96

CHAPTER 3

G. fossarum molecular responses

CHAPTER 3

G. fossarum molecular responses


97

KEY FINDINGS

Aim of the study

The aim of the study is the determination of a suitable set of reference genes for data normalization of RT-

qPCR experiments using G. fossarum. Six common candidate reference genes, tested and validated with other

species, were selected and their reliability for RT-qPCR were tested in different exposure conditions using

AgNO3, silver nanoparticles (AgNPs) and gold nanoparticles (AuNPs) contamination.

Experimental design

Key findings

• Set of six reference genes was tested

o Actin, TUB, UB, GAPDH, SDH, Clathrin

• One target gene was used to validated the stability of reference genes

o HSP90

• Stability of reference genes:

o Clathrin > SDH > GAPDH > TUB > Actin > UB

• RT-qPCR data normalization

o SDH + clathrin: down-regulation of HSP90 in G. fossarum exposed to AuNPs 40 nm

o SDH + clathrin + GAPDH: same results when data were normalized only with the two most

stable genes

o All genes: up-regulation of HSP90 in G. fossarum exposed to AgNPs 40 nm

• These results highlight the importance of selection of reference genes to obtain robust RT-qPCR data

RNA

Ribonucleic acid

Nucleobases

Base pair

Guanine

Uracil

Guanine

Adeninehelix of

sugar-phosphates

Nucleobases

of RNA

DNAoxyribonucleic a

Nucleobases

Base pair

Cytosi

Guanine

Adenine

Thymine

elix ofosphates

RNAextraction

Reversetranscriptase

cDNAsynthesis

qPCR

Acclimation

D-10

Exposure

D-7 D-1

Temperature : 12 oC

Photoperiod : 16h/8h

Daily renewal of Volvic water

Stalling

D15D-30 D-20

Leave

conditioning

Temperature : 12 oC


Exposure medium renewal: every 3 days Addition of food: every 2 days

D0


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Article 2

Article submitted to Scientific Reports

Identification of reference genes for RT-qPCR data normalization in Gammarus fossarum (Crustacea

Amphipoda)

Kahina Mehennaoui 1,2, Sylvain Legay 1, Tommaso Serchi 1, François Guérold 2, Laure Giambérini 2, Arno C.

Gutleb 1, Sébastien Cambier 1*

1 Environmental Research and Innovation (ERIN) Department, Luxembourg Institute of Science and

Technology (LIST), 5, avenue des Hauts-Fourneaux, Esch-sur-Alzette, Luxembourg

2 Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360, Université de

Lorraine – Metz, France


99

Identification of reference genes for RT-qPCR data normalization in Gammarus fossarum

(Crustacea Amphipoda)

Kahina Mehennaoui 1,2, Sylvain Legay 1, Tommaso Serchi 1, François Guérold 2, Laure Giambérini

2, Arno C. Gutleb 1, Sébastien Cambier 1*

1 Environmental Research and Innovation (ERIN) Department, Luxembourg Institute of Science

and Technology (LIST), 5, avenue des Hauts-Fourneaux, Esch-sur-Alzette, Luxembourg;

[email protected] (K.M.); [email protected] (T.S.); [email protected] (A.C.G.)

2 Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360,

Université de Lorraine – Metz, France ; [email protected] (F.G.) ;

[email protected] (L.G.)

* Correspondence: [email protected]; Tel.: +352-275-888-5018


100

Abstract: Gene expression profiling via RT-qPCR is a robust technique increasingly used in ecotoxicology.

Determination and validation of optimal reference genes is a requirement for initiating RT-qPCR experiments.

To our best knowledge, this study is the first attempt of identifying a set of reference genes for the freshwater

crustacean Gammarus fossarum. Six candidate genes (actin, TUB, UB, SDH, clathrin and GAPDH) were tested

in order to determine the most stable ones in different stress conditions and to increase the robustness of

RT-qPCR data. SDH and clathrin appeared as the most stable ones. A validation was performed using G.

fossarum samples exposed for 15 days to AgNO3, AgNPs 40 nm and AuNPs 40 nm. Effects on HSP90 were

evaluated and data normalized using clathrin and SDH. HSP90 was down-regulated when G. fossarum were

exposed to both AuNPs 40 nm whereas no effects were observed when G. fossarum were exposed to AgNPs

40 nm. This study highlights the importance of the preliminary determination of suitable reference genes for

RT-qPCR experiments. Additionally, this study allowed the determination of a set of valuable genes that can

be used in other RT-qPCR studies using G. fossarum as model organism

Keywords: Gammarus fossarum; Reference genes; RT-qPCR; Gene expression; Environmental toxicology

1. INTRODUCTION

Understanding the mechanisms underlying the effects of stressors on organisms needs sensitive analytical

techniques that can cover and link responses observed at different biological levels (from molecular to

individual responses). One of the most reliable technique is the measurement of changes or alterations in

gene expression in response to an external stimuli [1]. Recent advances in “omics” and bioinformatics

methodologies applied in ecotoxicological studies provided a new-angle of study of non-model organisms,

opening new ways in determining new molecular biomarkers (genes) as an alteration of their regulation may

influence the fitness of organisms [2,3]. Reverse transcription quantitative polymerase chain reaction (RT-

qPCR) is currently described as one of the most reliable techniques to assess these changes due to its

effectiveness, sensitivity and reproducibility [1,2,4]. This method, which allows studying the expression of a

set of selected genes in an organism, requires multiple critical quality controls in order to obtain robust

results. This includes RNA purity and integrity control, genomic DNA contamination assessment, evaluation

of PCR primer efficiency and specificity and, in case of relative quantification of gene expression, the

identification of suitable reference genes for data normalization [1,5,6].

Reference genes are described to be stable regardless the exposure conditions making them suitable for data

normalization of genes of interest. Therefore, their determination is crucial for a good analyze of RT-qPCR

results [2,5,7,8]. Furthermore, combination of a set of the most stable reference genes is fundamental to

avoid misinterpretation of the results that may occur when using a unique reference gene [1]. Consequently,


101

a preliminary experimental determination of the optimal reference genes is necessary before each new

experimental design [5,8]. For this purpose, many software and tools have been developed in order to

determine the best reference genes. This selection is performed following a ranking method based on their

stability across different stress conditions [5,7,9]. Unfortunately, many ecotoxicological studies are still using

only one, or a set of reference genes selected from previous studies, applying different experimental designs,

making them sometimes improper for data normalization. Classically known housekeeping genes such as

actin, tubulin (TUB), glyceraldehyde-3-phosphate dehydrogenase (GAPDH), and 18S ribosomal RNA are used

as reference genes [2,4,6,8]. However, recent studies demonstrated that some of these genes might not be

suitable as they are not always stable depending on the experimental design, studied stress and organisms

leading sometimes to the misinterpretation of the results [8,10].

Among the aquatic macroinvertebrates assemblage, amphipods of the genus Gammarus, and more

specifically Gammarus fossarum, represent a major part of it [11,12]. They are used in different

ecotoxicological studies for their high ecological relevance linked to their wide distribution all over Europe

and their major functional role in litter breakdown process and nutrient cycling [13,14]. Their well-

documented sensitivity to different kind of pollutants make them good model organisms for ecotoxicological

studies [15–18]. However, to our knowledge, few studies investigated their responses to exogenous stress at

molecular level, such as the vitellogenin expression following an estrogenic stress (Xuereb et al., 2011), the

antioxidant responses via catalase and MnSOD gene expression following exposure to gold nanoparticles

(AuNPs) (Baudrimont et al., 2017) and the identification of proteins expression profiles during

spermatogenesis [19–21] . Nevertheless, there is still no reference genes properly characterized and

dedicated to the amphipod G. fossarum. Reference genes are mandatory for an accurate data normalization

of gene expression [1]. Therefore, the aim of the present study is to fill this gap and to determine a suitable

set of reference genes for data normalization of RT-qPCR experiments using G. fossarum. Six common

candidate reference genes, tested and validated with other species [4,8,9], were selected and their reliability

for RT-qPCR were tested in different exposure conditions using AgNO3, silver nanoparticles (AgNPs) and gold

nanoparticles (AuNPs) contamination.

2. RESULTS

2.1. Stability of the candidate reference genes in G. fossarum

To identify the most appropriate set of reference genes for G. fossarum, six candidates were tested. The

genes included some frequently used ones (actin, TUB and GAPDH) and less common ones, namely, succinate

dehydrogenase (SDH), ubiquitin (UB) and clathrin. Five different methods were compared in order to


102

determine the most stable gene sub-set: GeNormPlus [22], NormFinder [23], BestKeeper [7], RefFinder [24]

and the comparative delta-CT method [25].

The ranking of the studied genes varied dependently on the algorithms used (Table 3. 1). Interestingly, the

same ranking was obtained using the most commonly used softwares, GenormPlus and NormFinder, with

clathrin and SDH as the most stable genes and UB as the least stable one. In the same way, RefFinder and the

comparative delta-CT method identified clathrin and SDH as the most stable genes. GAPDH was the third

most stable gene according to GenormPlus, NormFinder, and RefFinder, whereas, the rankings were different

for the least stable ones (Table 3. 1) as Actin was assigned the highest score by BestKeeper, the comparative

delta-CT method and RefFinder (Table 3. 1).

As BestKeepper’s ranking is based on correlation factor (R, the closer to 1, the better) and standard deviation

(the bigger, the worse) [7, 42], the obtained results differed from the other algorithms as Clathrin and Actin appeared

as the most stable genes whereas GAPDH and TUB were the least stable ones. According to BestKeeper, SDH was the

third most stable gene (Table 3.1)

Table 3. 1.Ranking of candidate reference genes according to the five algorithms used

Ranking

GeNorm NormFinder BestKeeper Comparative delta

Ct RefFinder

Gene Stability

coeff. Gene

Stability

coeff. Gene

Stability

coeff. Gene

Stability

coeff. Gene

Stability

coeff.

1 Clathrin 0.433 Clathrin 0.105 Clathrin 0.670 Clathrin 1.035 Clathrin 1.000

2 SDH 0.470 SDH 0.198 Actin 0.624 SDH 1.067 SDH 1.682

3 GAPDH 0.550 GAPDH 0.265 SDH 0.571 TUB 1.188 GAPDH 3.464

4 Actin 0.593 Actin 0.292 UB 0.575 GAPDH 1.190 TUB 3.464

5 TUB 0.652 TUB 0.465 GAPDH 0.459 UB 1.232 UB 5.000

6 UB 0.703 UB 0.520 TUB 0.395 Actin 3.217 Actin 6.000

In order to determine the stability of each gene, a global ranking was generated by assigning a number (from

1 to 6 where 1 is the most stable gene) to each stability coefficient presented in Table 3. 1 and by averaging

them [1,9]. This allowed the confirmation of the high stability of clathrin and SDH (Figure 3. 1). UB appeared

as the least stable gene followed by actin and TUB (Figure 3. 1).


103

Figure 3. 1. Global ranking of candidate reference genes in G. fossarum. A number (from 1 to 6) was assigned

to each stability coefficient. A mean rank was generated and error bars

2.2. Optimal number of reference gene for data normalization in G. fossarum using GeNorm

In order to calculate the optimal number of reference genes for data normalization in G. fossarum,

GeNormplus was used to generate the pairwise variation (Vn/Vn+1) between two normalization factors

(NF/NFn+1). Indeed, only GenormPlus allows an estimation of the optimal number of reference genes to use

in a specific experimental design.

The analysis conducted on 31 samples of G. fossarum exposed to AgNO3, AgNPs 40 nm and AuNPs 40 nm,

showed that the optimal number of reference genes is 2 as the V value is below the cut-off threshold of 0.15

making the addition of a third gene unnecessary (Figure 3. 2). As previously described, GeNormPlus identified

clathrin and SDH as the best combination for data normalization. In the same way, NormFinder determined

clathrin/SDH as the best pair as these two genes showed the highest stability coefficient (Table 3. 1).

0

1

2

3

4

5

6

7

clathrin SDH GAPDH TUB Actin UB

Meanrankvalue

Candidates referencegenes


104

Figure 3. 2 Determination of the optimal number of reference gens for data normalization in G. fossarum

exposed to AgNO3, AgNPs 40 nm and AuNPs 40 nm. The pairwise variation (Vn/Vn+1) was calculated between normalization factors NF/NFn+1. The recommended cut-off threshold of 0.15 was applied in this study.

2.3. Validation of the selected reference genes for G. fossarum

In order to validate G. fossarum reference genes, the expression of a general stress-related gene, the heat

shock protein 90 (HSP90), was evaluated. HSP90 expression profiles were studied on G. fossarum exposed

for 15 days to 0.5 µg. L-1 of AgNO3, CIT-AgNPs 40 nm, PEG-AgNPs 40 nm, CIT-AuNPs 40 nm and PEG-AuNPs

40 nm. Data were analysed using the Biogazelle qbase+ software and normalized using clathrin and SDH. As

shown in Figure 3. 3, a significant decrease in HSP90 expression was observed when G. fossarum were

exposed to 0.5 µg. L-1 of CIT-AuNPs 40 nm and PEG-AuNPs 40 nm (One-way ANOVA, P <0.001) while none of

the tested AgNPs or AgNO3 impacted HSP90 expression (Figure 3. 3, One-way ANOVA, P>0.05). However,

when data were normalized using all the six reference genes, the statistical analysis showed a significant

induction of the expression of HSP90 after treatment with CIT-AgNPs 40 nm (Figure 3. 3, One-way ANOVA, P

<0.05). Data were also normalized using the three best reference genes clathrin, SDH and GAPDH. No

differences in HSP90 expression were observed (Figure 3. 3, One-way ANOVA, P <0.05) enhancing that adding

a third gene for data normalization is unnecessary for our experimental design (Figure 3. 3).

0

0.02

0.04

0.06

0.08

0.1

0.12

0.14

V2/3 V3/4 V4/5 V5/6

Pairwisevariation(V)


105

Figure 3. 3 HSP90 expression analysis using different normalization strategies. Error bars indicate the standard

errors of the means (n=4). Different letter (a-c) indicate significant differences at P < 0.05.

3. DISCUSSION

Despite the rapid advances in the “omics” field marked by the development of next generation sequencing

(NGS) methods, RT-qPCR remains one of the most accurate and reliable techniques for targeted gene

expression and NGS data validation. This method is recognized for its capacity to highlight sensitive changes

in gene transcription levels [1,26]. However, for an accurate interpretation of RT-qPCR results, in term of

relative gene expression, one of the most important criteria is the selection of suitable reference genes,

which, to the best of our knowledge, are lacking for G. fossarum species. Previous studies have already

investigated molecular responses in Gammarus sp., exposed to AuNPs, temperature and ammonia, using

actin, or GAPDH as reference genes [19,27] on the assumption that these genes were stable and without

prior experimental verifications. Such approaches might conduct to misinterpretation of RT-qPCR results

[5,28,29] since the expression of these genes might be influenced by biotic or abiotic stress as well as

developmental stage and tissue type [4,8,30]. It is well established that a single reference gene could not be

applied to all the experimental designs [1,5]. Therefore, a case-specific choice of the best reference genes for

RT-qPCR is mandatory. In the present study, six candidate genes have been tested in adult G. fossarum males,

regarding a contamination with AgNO3, AgNPs and AuNPs, for their appropriateness to be used as reference

genes in transcriptomic studies [4]. The efficiency and specificity of the designed primers have been checked.

All the tested primers showed correct PCR efficiency and specificity during the PCR ensuring robust and

precise results interpretation [5], and confirmed that our RT-qPCR experiments comply with the known

0.00

0.50

1.00

1.50

2.00

2.50

Citrate PEG Citrate PEG

AgNPs40nm AuNPs40nm AgNO3

Norm

alizedrelativeexpression

Exposureconditions

Clathrin+SDH allgenes clathrin+SDH+GAPDH

b b

c

ac

a aa

aa

ab

a


106

recommendations [4,5]. Five different algorithms were used to determine optimal reference genes to

normalize our data. The five used softwares identified clathrin and SDH as the most stable genes. These

results are in accordance with previous studies that described SDH as a suitable reference gene in Rhodnius

prolixus [10] and in red abalone Haliotis rufescens’ gonad and digestive tissues [31]. Clathrin was also

identified as one of the most stable genes in Lilium formolongi [32].

It is important to notice that our ranking of the candidate reference genes shows that the most commonly

used genes in transcriptomics studies are the least stable ones in this study. This observation is in agreement

with other recent studies which showed that GAPDH, 18s rRNA, actin and TUB were not good candidates as

reference genes [4,8,26,33]. GAPDH, commonly considered as a housekeeping gene, plays an important role

in energetic metabolism and its expression was described as significantly impacted in bivalves, like Mytilus

spp. or Crassostrea gigas, exposed to different environmental stress (harbor pollution), and in Haliotis discus

hannai under heavy metal stress conditions [4,33]. On the contrary, other studies showed that GAPDH was

one of the most stable gene in two flatworms species, Macrostomum lignano and Schmidtea mediterranea,

exposed to cadmium [26] and in R. prolixus in different physiological conditions and feeding status [10].These

observation indicate that GAPDH should not be considered as suitable for data normalization without prior

validation. Other studies demonstrated that selection of unstable genes like actin and 18S rRNA as reference

genes greatly altered interpretation of data for Cathepsin D in Ruditapes philippinarum exposed to copper

[8]. Similar observations were done for TUB and UB in Haliotis discus hannai [4]. These results are in

agreement with our observations in G. fossarum where classical reference genes like actin, TUB, UB and

GAPDH appeared among the least stable genes. The present study shows that it is crucial to experimentally

assess the stability of reference genes for each species tested in each experimental design applied before

they can be selected as housekeeping genes [8].

In addition, the stability of the present reference genes was verified by exposing G. fossarum to AgNO3, AgNPs

40 nm and AuNPs 40 nm for 15 days and studying the expression of HSP90. HSP90 is involved in the regulation

of proteostasis under both physiological and stress conditions. It plays many roles such as protein folding as

a chaperone, DNA repair and immune responses [34]. HSP90 is also known to play a role, with HSP70, in

keeping inactive the heat shock factor (HSF1) which induce the expression of HSP90 in stress condition

[34,35]. Therefore, it was used as a target gene in the present study. Data were normalized using, firstly

clathrin and SDH, which were indicated by GeNormPlus to be the most stable genes, then, clathrin, SDH and

GPDH and finally with the whole set of six candidate reference genes. When clathrin and SDH are used for

normalization, a significant decrease in HSP90 expression was observed in G. fossarum exposed to CIT-AuNPs

40 nm and PEG-AuNPs 40 nm. However, when data were normalized using clathrin, SDH and GAPDH, no

differences in HSP90 expression profile was observed making the addition of a third gene unnecessary.


107

Furthermore, data normalization using all the six-studied reference genes, led to the detection of an

upregulation of HSP90 in G. fossarum exposed to CIT-AgNPs 40 nm, which in this case is to be considered as

an overestimation of the response as “incorrect references” were used. This is in accordance with what has

been previously reported [26]. Normalization of HSP90 expression in M. lignano exposed to Cd using all the

nine reference genes tested led to a high variability between replicates. Authors stated that this observation

could be linked to an important variability between replicates that lower the resolution of detection of

differences between their different conditions [26]. However, when data were normalized using the three

most stable genes, no significant differences in HSP90 expression were observed [26]. Other study showed

that a non-optimal selection of the best combination of reference genes may lead to statistical

misinterpretation [10]. Authors showed that data normalization using an unstable gene such eiF-1a in R.

prolixus led to a clear but false increase in an olfactory gene, RproIR76b expression while no statistical

difference were observed when data were normalized using the most stable genes [10]. Moreover, other

study underlined the importance of selection of the appropriate reference genes with the highest stability

coefficient as data normalization of metallothionein expression in abalone exposed to copper let to an

underestimation or overestimation of the effects when data were normalized using unstable reference genes

[4]. These results highlight once more the importance of an experimental validation of reference gene in

addition to the selection of the optimal number and the appropriate genes for data normalization [5,10,26].

4. MATERIALS AND METHODS

4.1. Organisms sampling and acclimation

G. fossarum were collected at an unpolluted stream (49°48’24.9’’ N and 06°04’53.2’’ E, Schwaarzbaach,

Colmar-berg, Luxembourg) [18,36]. Animals were collected using a hand net and were sorted in the field.

They were immediately brought to the laboratory in river water, where they were kept at 12 °C. In order to

avoid influence of gender on the studied parameters, only adult males were kept for the experiment [16,37].

They were selected from precopula pairs or based on sexual dimorphism like gnatopode size [18]. Adult males

were then acclimated to laboratory conditions [18,38]. The acclimation was conducted on 2 steps. First,

Gammarids were acclimated for 72h to mineral water (Volvic, France) by progressively changing field water

to Volvic water (30% v/v, 50% v/v, 100% v/v) Then, a stalling period of 10 days was conducted in 100% Volvic

water [18,38]. The acclimation was performed under controlled conditions at 12 °C with a 16h light 8h dark

photoperiod. Volvic water was aerated and changed every 24h to avoid organic matter accumulation and

potential increase of ammonium, nitrite and nitrate. Gammarids were fed ad libitum with alder leaves (Alnus

glutinosa) up to 24h before experiment.


108

4.2. AgNO3, AgNPs and AuNPs contamination

At the end of the acclimation period, four pools of 3 males were exposed to 0.5 µg.L-1 of AgNO3, AgNPs 40

nm and AuNPs 40 nm, either stabilized with citrate (CIT-AgNPs 40 nm and CIT-AuNPs) or coated with

polyethylene-glycol (PEG-AgNPs 40 nm and PEG-AuNPs 40 nm), for 15 days at 12 ºC with a photoperiod of

16h light and 8h darkness [18]. Exposure medium (Volvic water) was changed every 72h. Food (Alnus

glutinosa disk leaves) was added every 48h. At the end of the exposure period, Gammarids were gently dried

and stored at -80 ºC in RLT buffer supplemented with 1% β-mercaptoethanol until RNA extraction.

4.3. Gene identification and qPCR primer design

4.3.1. DNA extraction

Pools of 50 eggs of G. fossarum were placed in ATL Buffer supplemented with 10% (v/v) of proteinase K,

placed on thermomixer at 56 ºC at 800 rpm for 1h. Genomic DNA was extracted using DNeasy blood and

tissue kit following the manufacturer’s instructions (Qiagen, Leusden, The Netherlands). Quantification and

purity were performed using Nanodrop ND-1000 (Thermo scientific, Waltham, MA, USA). The resulting gDNA

was stored at -20ºC until use.

4.3.2. Library preparation and sequencing

Libraries were prepared from 50 ng of genomic DNA using the Nextera DNA library preparations kit

according to the manufacturer’s instructions (Illumina, San Diego, CA, USA). After labelling of gDNA

fragments, clean up and PCR amplification, libraries were controlled using the Agilent 2100

bioanalyzer and DNA high sensitivity kit. Libraries were quantified using the Library quantification

kit – Illumina Low ROX (Kapa Biosystems, Boston, MA, USA) on a Viia7 96-well real-time-qPCR

instrument (Life Technologies). The libraries were sequenced using an Illumina Miseq by means of

3 consecutive runs (Illumina Miseq reagent V3) to generate 76 base-pairs paired-end reads.

4.3.3. De novo assembly

Miseq generated FASTQ files were imported in CLC genomics workbench v9 discarding reads with low quality

(<Q30). Illumina adapters and indexes were trimmed using the Nextera adaptor sequence and filtered

(%filtered). Reads with nucleotide ambiguity (N) and CLC quality index above 0.05 were then filtered from

the remaining sequences. Two trimming procedures were applied. First, the Nextera adaptor sequences were

filtered in order to avoid the presence of technical sequences (adaptors and indexes) in the reads. Conserved

sequences generated by the hybridization and amplification of random hexamers (commonly found in the


109

stranded libraries), were trimmed using an arbitrary “hard trim” of 16 base pairs at the 5’ end of the reads

and 5 base pairs at the 3’ end of the reads. These values were determined from the QC report analysis

obtained from CLC genomics. Finally, the remaining reads were assembled using a De novo assembly

following the upcoming criteria: minimum contig length of 200bp, mismatch, insertion and deletion costs of

3, length and similarity fraction of 0.8 and maximum distance of 1000 bp.

4.3.4. Gene identification

In order to identify and amplify putative genes from G. fossarum, an initial data mining was performed on

the raw reads previously sequenced by [20] in addition to the reads obtained as described above (see 4.3.3).

Reads were mapped to the sequences of Hyalella azteca (https://www.hgsc.bcm.edu/arthropods/hyalella-

azteca-genome-project), a closely related species for which the transcriptome is available, with the following

criteria: the mapped reads must have multiple hits lower than 10, a minimum of 80% identity and 80%

coverage with the reference. Mismatch costs was set at 2 (medium) and deletion/insertion cost at 3 (highest

stringency). A consensus sequence was generated from the mapped reads for each gene (Table 3. 2). In order

to verify whether the obtained sequences are coding for a protein, a translation of the obtained nucleotide

sequences was performed using ExPASy translation tool (http://web.expasy.org/translate/). Finally, a Blastx

search was performed against non-redundant protein databases from the National Centre for Biotechnology

(NCBI) to check the identity of the selected sequences. (Table 3. 2).


108

Table 3. 2 Identification of Gammarus fossarum gene sequences

Genes

Hyalella azteca G. fossarum consensus

sequences NCBI Blastx

Accession

number Accession number Homology Identification Accession number

Actin XM_018157137.1 MF940257 97% actin, partial

[Hoplolaimus galeatus] AEM45650.1

Tubulin XM_018153872.1 MF940258

96%

PREDICTED: tubulin alpha-8

chain-like isoform X1

[Serinus canaria]

XP_009098159.2

Ubiquitin XM_018170409.1 MF940259 96%

ubiquitin conjugating

enzyme-3

[Eriocheir sinensis]

ADF45343.1

GAPDH XM_018154227.1 MF940254 95%

putative glyceraldehyde-3-

phosphate dehydrogenase

[Gammarus locusta]

CAQ60115.1

SDH XM_018156499.1 MF940255 96%

PREDICTED: succinate

dehydrogenase

[ubiquinone] iron-sulfur

subunit, mitochondrial-like

[Hyalella azteca]

XP_018011988.1

Clathrin XM_018171236.1 Sequence provided in

supplementary material 93%

PREDICTED: clathrin light

chain-like isoform X1

[Hyalella azteca]

XP_018025977.1

HSP90 XM_018155941.1 MF04256 98%

PREDICTED: heat shock

protein HSP 90-alpha-like

[Hyalella azteca]

XP_018022683.1


109

Table 3. 3 List of primers of the candidate reference genes and target gene HSP90

Name Sequence (5'→ 3') Amplicon Length (bp) Amplicon Tm (°C) PCR Efficiency Regression

Coeff. (R2)

Actin_F CTCATGCTATCCTTCGTCTTGA 103 78 2.02 0.999

Actin_R CGTTCAGCGGTGGTTACAA

Tubulin_F CGGCTGTTGTTGAACCTTAC 93 81 2.09 0.999

Tubulin_R AGATGGCCTCATTGTCAACC

GAPDH_F GTCCGTCTCGCTAAGGAGTG 94 85 1.91 0.999

GAPDH_R TGTATCCGAGGTAGCCCTTG

SDH_F GGAAGAAGCTGGATGGTCTG 87 84 1.98 0.998

SDH_R ACTTGTCTCCGTTCCACCAG

Ubiquitin_F CCCACGATACTCCCTTTGAA 82 79 2.01 0.991

Ubiquitin_R ACAATCGGTGGCTTGTTAGG

Clathrin_F ATCGCCAAGCTTTGTGACTT 107 85 1.99 0.999

Clathrin_R GCTTTGATAGGCGGACTCTG

HSP90_F CTGGTTTCTTCTCCCTGCTG 135 85 1.99 0,995

HSP90_R GATCTCGAGGTGCTTCTTGG


110

4.3.5. Primer design

All the primers were designed using Primer3Plus (http://primer3.ut.ee) with the following criteria: primer

size between 18 and 25 base pairs, GC content between 40% and 60 %, amplicon size from 80 to 150 base

pairs, primer annealing temperatures in the 58-61 ºC range. Primers were checked using NetPrimer

(http://www.premierbiosoft.com/netprimer/) for secondary unexpected structures. PCR efficiency was

evaluated using decreasing five-fold dilutions from cDNA pool (from 25 ng to 0.04 ng and no template

control). A melting curve was performed at the end of each run, in order to assess the specificity of the

amplified products. All tested genes displayed one clear peak and were therefore retained for analyses.

Primers sequences, amplicon size, and melting temperature are described in Table 3. 3.

4.4. RNA extraction, cDNA and RT-qPCR

G. fossarum tissues were ground on ice using a pellet pestle tissue grinder. Homogenates were centrifuged

at 250g for 5 min at 4°C to remove cuticle fragments as described in [18]. Total RNA was extracted using

RNeasy mini kit (Qiagen, Leusden, The Netherlands) according to the manufacturer’s instructions for cells

and animal tissues (including DNase treatment) [39,40]. RNA concentrations and purity were assessed

measuring the absorbance at 230, 260 and 280 nm using Nanodrop ND-1000 (ThermoScientific, Villebon-sur-

Yvette, France). Finally, RNA integrity was checked using the RNA Nano 6000 assay (Agilent Technologies,

Diegem, Belgium) and 2100 Bioanalyzer (Agilent, Santa Clara, CA, USA)[39]. All RNA samples displayed no

degradation patterns (sharp peaks and clean baseline).

The extracted RNA were reverse-transcribed into cDNA using the Protoscript II reverse transcriptase (New

England Biolabs, Leiden, The Netherlands) and random primers, according to the manufacturer’s instructions.

cDNAs were further diluted to 0.8 ng. µL-1 and used for RT-qPCR analyses using a 384-well plate design. An

automated liquid handling robot (epMotion 5073, Eppendorf, Hambourg, Germany) was used to properly

prepare the 384-well plates. qPCR were performed using the Takyon Low ROX SYBR MasterMix dTTP Blue Kit

(Eurogentec, Liège, Belgium) on a ViiA 7 Real-Time PCR System (Thermo-Fisher, Waltham, MA, USA) in a 10

µL final volume [41]. All reactions were performed in technical triplicates and repeated on four biological

replicates. The PCR conditions consisted on an initial denaturation step at 95°C for 3 min, followed by 40

cycles of denaturation at 95 °C for 15 sec and annealing/extension steps at 60 °C for 60 sec.

4.5. Stability of the candidates’ reference genes

The stability of the selected genes was analysed using five different methodologies. GeNormplus performs a

pairwise comparison and generate the M-value which consist on a comparison of the variation of a gene

compared to all the remaining candidates [22] while NormFinder calculates both a single best gene (best


111

gene) and an optimal gene pair (best pair) as the best pair may compensate the expression in the different

experimental groups [1,23]. BestKeeper on its side is based on assigning correlation factor of each gene with

the geometric means of all genes [7,42] while RefFinder is an online very easy to use platform which compiles

the three most popular algorithms for reference gene validation based on an input of Cq values only [24,42].

Additionally, a simple delta-Ct comparison approach was applied [25].

4.6. Statistical analyses

Normal distribution of data-set was checked using a Shapiro-wilk test. Homogeneity of variances was checked

using a Levene test. A one-way ANOVA followed by a Fisher-LSD post hoc test (P < 0.05) was performed on

the log2 transformed data (CNRQs) using Statistica 12 software (Statsoft Inc.).

5. CONCLUSIONS

This study provides of a set of reference genes suitable for normalization of RT-qPCR data obtained from G.

fossarum samples. Six candidate genes were tested and five different algorithms allowed the identification

of the most stable sub-set of genes. clathrin and SDH were identified as the most stable genes in our applied

experimental design, while widely used reference genes were unsuitable in G. fossarum in the present work.

Our results highlight how important and crucial is to experimentally define and validate of a set of reference

genes for each RT-qPCR experiment.


112

Acknowledgments: This work is supported by the Fonds National de la Recherche (NanoGAM AFR-PhD-

9229040). The authors are grateful to S. Contal and M. Fossépré for technical supports.

Author Contributions: K.M conceived, designed and performed the experiments, prepared tables and figures

and wrote the manuscript. K.M, S.L, S.C analysed the data; S.C. coordinated its revision; All authors read and

provided helpful discussions and approved the final discussion

Conflicts of Interest: The authors declare no conflict of interest.

Abbreviations

RT-qPCR Reverse transcription quantitative polymerase chain reaction

TUB Tubulin

UB Ubiquitin

SDH Succinate dehydrogenase

GAPDH Glyceraldehyde-3-phosphate dehydrogenase

HSP90 Heat shock protein 90

AgNPs Silver nanoparticles

AuNPs Gold nanoparticles

AgNO3 Silver nitrate

NF Normalization factor

Vn Pairwise variations


113

REFERENCES

1. Guerriero, G.; Legay, S.; Hausman, J.-F. Alfalfa Cellulose Synthase Gene Expression under Abiotic Stress:

A Hitchhiker’s Guide to RT-qPCR Normalization. PLOS ONE 2014, 9, e103808,

doi:10.1371/journal.pone.0103808.

2. Brulle, F.; Bernard, F.; Vandenbulcke, F.; Cuny, D.; Dumez, S. Identification of suitable qPCR reference

genes in leaves of Brassica oleracea under abiotic stresses. Ecotoxicology 2014, 23, 459–471,

doi:10.1007/s10646-014-1209-7.

3. Shaw, J. R.; Colbourne, J. K.; Davey, J. C.; Glaholt, S. P.; Hampton, T. H.; Chen, C. Y.; Folt, C. L.; Hamilton,

J. W. Gene response profiles for Daphnia pulex exposed to the environmental stressor cadmium reveals

novel crustacean metallothioneins. BMC Genomics 2007, 8, 1–20, doi:10.1186/1471-2164-8-477.

4. Lee, S. Y.; Nam, Y. K. Evaluation of reference genes for RT-qPCR study in abalone Haliotis discus hannai

during heavy metal overload stress. Fish. Aquat. Sci. 2016, 19, doi:10.1186/s41240-016-0022-z.

5. Bustin, S. A.; Benes, V.; Garson, J. A.; Hellemans, J.; Huggett, J.; Kubista, M.; Mueller, R.; Nolan, T.; Pfaffl,

M. W.; Shipley, G. L.; Vandesompele, J.; Wittwer, C. T. The MIQE Guidelines: Minimum Information for

Publication of Quantitative Real-Time PCR Experiments. Clin. Chem. 2009, 55, 611–622,

doi:10.1373/clinchem.2008.112797.

6. Heckmann, L.-H.; Connon, R.; Hutchinson, T. H.; Maund, S. J.; Sibly, R. M.; Callaghan, A. Expression of

target and reference genes in Daphnia magna exposed to ibuprofen. BMC Genomics 2006, 7, 1–8,

doi:10.1186/1471-2164-7-175.

7. Pfaffl, M. W.; Tichopad, A.; Prgomet, C.; Neuvians, T. P. Determination of stable housekeeping genes,

differentially regulated target genes and sample integrity: BestKeeper – Excel-based tool using pair-wise

correlations. Biotechnol. Lett. 2004, 26, 509–515, doi:10.1023/B:BILE.0000019559.84305.47.

8. Volland, M.; Blasco, J.; Hampel, M. Validation of reference genes for RT-qPCR in marine bivalve

ecotoxicology: Systematic review and case study using copper treated primary Ruditapes philippinarum

hemocytes. Aquat. Toxicol. 2017, 185, 86–94, doi:10.1016/j.aquatox.2017.01.003.

9. Mangeot-Peter, L.; Legay, S.; Hausman, J.-F.; Esposito, S.; Guerriero, G. Identification of Reference Genes

for RT-qPCR Data Normalization in Cannabis sativa Stem Tissues. Int. J. Mol. Sci. 2016, 17, 1556,

doi:10.3390/ijms17091556.

10. Omondi, B. A.; Latorre-Estivalis, J. M.; Rocha Oliveira, I. H.; Ignell, R.; Lorenzo, M. G. Evaluation of

reference genes for insect olfaction studies. Parasit. Vectors 2015, 8, 243, doi:10.1186/s13071-015-

0862-x.

11. Dedourge-Geffard, O.; Palais, F.; Biagianti-Risbourg, S.; Geffard, O.; Geffard, A. Effects of metals on

feeding rate and digestive enzymes in Gammarus fossarum: An in situ experiment. Chemosphere 2009,

77, 1569–1576, doi:10.1016/j.chemosphere.2009.09.042.


114

12. Ladewig, V.; Jungmann, D.; Köhler, H.-R.; Schirling, M.; Triebskorn, R.; Nagel, R. Population Structure and

Dynamics of Gammarus fossarum (Amphipoda) Upstream and Downstream from Effluents of Sewage

Treatment Plants. Arch. Environ. Contam. Toxicol. 2006, 50, 370–383, doi:10.1007/s00244-005-7039-0.

13. Kelly, D. W.; Dick, J. T. A.; Montgomery, W. I. The functional role of Gammarus(Crustacea, Amphipoda):

shredders, predators, or both? Hydrobiologia 2002, 485, 199–203, doi:10.1023/A:1021370405349.

14. Lacaze, E.; Geffard, O.; Bony, S.; Devaux, A. Genotoxicity assessment in the amphipod Gammarus

fossarum by use of the alkaline Comet assay. Mutat. Res. Toxicol. Environ. Mutagen. 2010, 700, 32–38,

doi:10.1016/j.mrgentox.2010.04.025.

15. Alonso, Á.; Lange, H. J. D.; Peeters, E. T. H. M. Contrasting sensitivities to toxicants of the freshwater

amphipods Gammarus pulex and G. fossarum. Ecotoxicology 2010, 19, 133–140, doi:10.1007/s10646-

009-0398-y.

16. Arce Funck, J.; Danger, M.; Gismondi, E.; Cossu-Leguille, C.; Guérold, F.; Felten, V. Behavioural and

physiological responses of Gammarus fossarum (Crustacea Amphipoda) exposed to silver. Aquat.

Toxicol. 2013, 142–143, 73–84, doi:10.1016/j.aquatox.2013.07.012.

17. Felten, V.; Charmantier, G.; Mons, R.; Geffard, A.; Rousselle, P.; Coquery, M.; Garric, J.; Geffard, O.

Physiological and behavioural responses of Gammarus pulex (Crustacea: Amphipoda) exposed to

cadmium. Aquat. Toxicol. 2008, 86, 413–425, doi:10.1016/j.aquatox.2007.12.002.

18. Mehennaoui, K.; Georgantzopoulou, A.; Felten, V.; Andreï, J.; Garaud, M.; Cambier, S.; Serchi, T.; Pain-

Devin, S.; Guérold, F.; Audinot, J.-N.; Giambérini, L.; Gutleb, A. C. Gammarus fossarum (Crustacea,

Amphipoda) as a model organism to study the effects of silver nanoparticles. Sci. Total Environ. 2016,

doi:10.1016/j.scitotenv.2016.06.068.

19. Baudrimont, M.; Andrei, J.; Mornet, S.; Gonzalez, P.; Mesmer-Dudons, N.; Gourves, P.-Y.; Jaffal, A.;

Dedourge-Geffard, O.; Geffard, A.; Geffard, O.; Garric, J.; Feurtet-Mazel, A. Trophic transfer and effects

of gold nanoparticles (AuNPs) in Gammarus fossarum from contaminated periphytic biofilm. Environ.

Sci. Pollut. Res. 2017, doi:10.1007/s11356-017-8400-3.

20. Trapp, J.; Armengaud, J.; Pible, O.; Gaillard, J.-C.; Abbaci, K.; Habtoul, Y.; Chaumot, A.; Geffard, O.

Proteomic Investigation of Male Gammarus fossarum, a Freshwater Crustacean, in Response to

Endocrine Disruptors. J. Proteome Res. 2014, doi:10.1021/pr500984z.

21. Xuereb, B.; Bezin, L.; Chaumot, A.; Budzinski, H.; Augagneur, S.; Tutundjian, R.; Garric, J.; Geffard, O.

Vitellogenin-like gene expression in freshwater amphipod Gammarus fossarum (Koch, 1835): functional

characterization in females and potential for use as an endocrine disruption biomarker in males.

Ecotoxicology 2011, 20, 1286–1299, doi:10.1007/s10646-011-0685-2.

22. Vandesompele, J.; De Preter, K.; Pattyn, F.; Poppe, B.; Van Roy, N.; De Paepe, A.; Speleman, F. Accurate

normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control

genes. Genome Biol. 2002, 3, research0034, doi:10.1186/gb-2002-3-7-research0034.


115

23. Andersen, C. L.; Jensen, J. L.; Ørntoft, T. F. Normalization of Real-Time Quantitative Reverse

Transcription-PCR Data: A Model-Based Variance Estimation Approach to Identify Genes Suited for

Normalization, Applied to Bladder and Colon Cancer Data Sets. Cancer Res. 2004, 64, 5245–5250,

doi:10.1158/0008-5472.CAN-04-0496.

24. Xie, F.; Xiao, P.; Chen, D.; Xu, L.; Zhang, B. miRDeepFinder: a miRNA analysis tool for deep sequencing of

plant small RNAs. Plant Mol. Biol. 2012, doi:10.1007/s11103-012-9885-2.

25. Silver, N.; Best, S.; Jiang, J.; Thein, S. L. Selection of housekeeping genes for gene expression studies in

human reticulocytes using real-time PCR. BMC Mol. Biol. 2006, 7, 33, doi:10.1186/1471-2199-7-33.

26. Plusquin, M.; DeGheselle, O.; Cuypers, A.; Geerdens, E.; Roten, A. V.; Artois, T.; Smeets, K. Reference

genes for qPCR assays in toxic metal and salinity stress in two flatworm model organisms. Ecotoxicology

2012, 21, 475–484, doi:10.1007/s10646-011-0809-8.

27. Henry, Y.; Piscart, C.; Charles, S.; Colinet, H. Combined effect of temperature and ammonia on molecular

response and survival of the freshwater crustacean Gammarus pulex. Ecotoxicol. Environ. Saf. 2017, 137,

42–48, doi:10.1016/j.ecoenv.2016.11.011.

28. Dong, M.; Zhang, X.; Chi, X.; Mou, S.; Xu, J.; Xu, D.; Wang, W.; Ye, N. The validity of a reference gene is

highly dependent on the experimental conditions in green alga Ulva linza. Curr. Genet. 2012, 58, 13–20,

doi:10.1007/s00294-011-0361-3.

29. Kozera, B.; Rapacz, M. Reference genes in real-time PCR. J. Appl. Genet. 2013, 54, 391–406,

doi:10.1007/s13353-013-0173-x.

30. Zavala, E.; Reyes, D.; Deerenberg, R.; Vidal, R. Selection of reference genes for microRNA analysis

associated to early stress response to handling and confinement in Salmo salar. Sci. Rep. 2017, 7, 1756,

doi:10.1038/s41598-017-01970-3.

31. López-Landavery, E. A.; Portillo-López, A.; Gallardo-Escárate, C.; Del Río-Portilla, M. A. Selection of

reference genes as internal controls for gene expression in tissues of red abalone Haliotis rufescens

(Mollusca, Vetigastropoda; Swainson, 1822). Gene 2014, 549, 258–265, doi:10.1016/j.gene.2014.08.002.

32. Zhang, M. F.; Liu, Q.; Jia, G. X. Reference gene selection for gene expression studies in lily using

quantitative real-time PCR. Genet. Mol. Res. 2016, 15, doi:10.4238/gmr.15027982.

33. Lacroix, C.; Coquillé, V.; Guyomarch, J.; Auffret, M.; Moraga, D. A selection of reference genes and early-

warning mRNA biomarkers for environmental monitoring using Mytilus spp. as sentinel species. Mar.

Pollut. Bull. 2014, 86, 304–313, doi:10.1016/j.marpolbul.2014.06.049.

34. Schopf, F. H.; Biebl, M. M.; Buchner, J. The HSP90 chaperone machinery. Nat. Rev. Mol. Cell Biol. 2017,

18, 345–360, doi:10.1038/nrm.2017.20.

35. Voellmy, R.; Boellmann, F. Chaperone Regulation of the Heat Shock Protein Response. 2007, 89–99,

doi:10.1007/978-0-387-39975-1_9.


116

36. Dohet, A.; Ector, L.; Cauchie, H.-M.; Hoffmann, L. Identification of benthic invertebrate and diatom

indicator taxa that distinguish different stream types as well as degraded from reference conditions in

Luxembourg. Anim. Biol. 2008, 58, 419–472, doi:10.1163/157075608X383719.

37. Sornom, P.; Felten, V.; Médoc, V.; Sroda, S.; Rousselle, P.; Beisel, J.-N. Effect of gender on physiological

and behavioural responses of Gammarus roeseli (Crustacea Amphipoda) to salinity and temperature.

Environ. Pollut. 2010, 158, 1288–1295, doi:10.1016/j.envpol.2010.01.022.

38. Andreï, J.; Pain-Devin, S.; Felten, V.; Devin, S.; Giambérini, L.; Mehennaoui, K.; Cambier, S.; Gutleb, A. C.;

Guérold, F. Silver nanoparticles impact the functional role of Gammarus roeseli (Crustacea Amphipoda).

Environ. Pollut. 2016, 208, Part B, 608–618, doi:10.1016/j.envpol.2015.10.036.

39. Legay, S.; Guerriero, G.; Deleruelle, A.; Lateur, M.; Evers, D.; André, C. M.; Hausman, J.-F. Apple russeting

as seen through the RNA-seq lens: strong alterations in the exocarp cell wall. Plant Mol. Biol. 2015, 88,

21–40, doi:10.1007/s11103-015-0303-4.

40. Legay, S.; Guerriero, G.; André, C.; Guignard, C.; Cocco, E.; Charton, S.; Boutry, M.; Rowland, O.;

Hausman, J.-F. MdMyb93 is a regulator of suberin deposition in russeted apple fruit skins. New Phytol.

2016, 212, 977–991, doi:10.1111/nph.14170.

41. Behr, M.; Legay, S.; Hausman, J.-F.; Guerriero, G. Analysis of Cell Wall-Related Genes in Organs of

Medicago sativa L. under Different Abiotic Stresses. Int. J. Mol. Sci. 2015, 16, 16104–16124,

doi:10.3390/ijms160716104.

42. De Spiegelaere, W.; Dern-Wieloch, J.; Weigel, R.; Schumacher, V.; Schorle, H.; Nettersheim, D.;

Bergmann, M.; Brehm, R.; Kliesch, S.; Vandekerckhove, L.; Fink, C. Reference Gene Validation for RT-

qPCR, a Note on Different Available Software Packages. PLoS ONE 2015, 10,

doi:10.1371/journal.pone.0122515.


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COMPLEMENT 2

Identification of reference genes for RT-qPCR data normalization in Gammarus fossarum

(Crustacea Amphipoda)

Kahina Mehennaoui 1,2, Sylvain Legay 1, Tommaso Serchi 1, François Guérold 2, Laure Giambérini 2,

Arno C. Gutleb 1, Sébastien Cambier 1*

1 Environmental Research and Innovation (ERIN) Department, Luxembourg Institute of Science and

Technology (LIST), 5, avenue des Hauts-Fourneaux, Esch-sur-Alzette, Luxembourg

2 Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360,

Université de Lorraine – Metz, France

Article submitted to Scientific Reports


118

Table S1: Identification of Gammarus fossarum gene sequences


119

Table S2: Cq values (mean ± SD) of the different genes regarding the different exposure conditions.

a CIT: citrate stabilized NPs, PEG: polyethylene-glycol coated NPs b 0: this condition is the control condition for each compound

Compounda Concentrations(µg.L-1)b Actin Clathrin GAPDH SDH TUB UB HSP90

AgNO3

0 21.60± 0.57 21.40± 0.30 18.69± 0.52 19.99± 0.40 18.60± 0.28 24.16± 0.34 18.73± 0.48

0,5 20.93± 0.30 21.23± 0.04 18.93± 0.20 19.82± 0.13 18.01± 0.22 23.86± 0.11 18.48± 0.20

AgNPs40nm

0 21.35± 0.21 21.12± 0.57 18.65± 0.13 20.21± 0.08 18.39± 0.11 23.39± 0.60 19.22± 0.10

CIT 0,5 22.22± 0.24 21.32± 0.03 18.52± 0.38 20.23± 0.10 17.97± 0.21 23.85± 0.12 18.78± 0.09

PEG 0,5 21.67± 0.10 21.34± 0.05 18.60± 0.21 20.31± 0.03 18.02± 0.32 23.94± 0.12 19.03± 0.29

AuNPs40nm

0 21.55± 0.17 21.21± 0.13 18.78± 0.14 20.17± 0.06 18.07± 0.30 24.01± 0.05 18.57± 0.28

CIT 0,5 21.45± 0.15 21.25± 0.08 18.85± 0.18 20.07± 0.03 18.14± 0.29 24.00± 0.11 19.16± 0.09

PEG 0,5 21.56± 0.15 21.26± 0.13 18.71± 0.20 19.98± 0.08 18.69± 0.13 23.58± 0.60 19.04± 0.17

Chapter 4 AgNPs and AuNPs acute toxicity

118

CHAPTER 4

AgNPs and AuNPs acute toxicity

CHAPTER 4

AgNPs and AuNPs acute toxicity


119

KEY FINDINGS

Aim of the study

This study aimed at investigating the influence of the size (20, 40 and 80 nm) and surface coating (CIT and

PEG) on the uptake, tissue distribution and molecular effects of AgNPs and AuNPs on G. fossarum exposed

for 72h to AgNPs and AuNPs, in the absence of food

Experimental design

Key findings

• Coating-dependent uptake of Ag and Au o CIT-AgNPs > PEG-AgNPs

o CIT-AuNPs > PEG-AuNPs

• No size-dependent effects on bioaccumulation

• Concentration-dependent uptake

• Influence of chemical composition on the internal distribution was observed as o AgNPs were found only in G. fossarum gills

o AuNPs were found only in G. fossarum gut

• A metal-dependent effect on gene expression was also observed with (e.g.) o AgNPs leading to an increase in MnSOD and CuZnSOD gene expression

o AuNPs leading to a decrease in MnSOD and CuZnSOD gene expression

Luxembourg

Gammarus fossarum

Organisms Materials

Schwaarzbaach

Sampling site

Ag20nm Ag40nm Ag80nm

Citrate stabilized AgNPs and PEG coated AgNPs

Au20nm Au40nm Au80nm

Citrate stabilized AuNPs and PEG coated AuNPs

Acclimation

Volvic water

D1

1/3

Exposure

T0h T24h T48h T72hD4 D10D2 D3

2/33/3 Temperature : 12 oC



Temperature : 12 oC


Daily renewal of exposure medium

Stalling

Experimental design


120

ARTICLE 3

Submitted to Science of the Total Environment

Influence of size and surface coating on uptake of silver and gold nanoparticles and the molecular effects

on Gammarus fossarum (Crustacea Amphipoda)

Kahina Mehennaoui a,b, Sébastien Cambier a, Tommaso Serchi a, François Guérold b, Johanna Ziebel a, Jean-

Sebastien Thomann c, Nathalie Valle c, Laure Giambérini b, *, Arno C. Gutleb a*

A Environmental Research and Innovation (ERIN) Department, Luxembourg Institute of Science and

Technology, 41 rue du Brill, Belvaux, Luxembourg

B Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360, Université de


C Material Research and Technology (MRT) Department, Luxembourg Institute of Science and Technology, 41

rue du Brill, Belvaux, Luxembourg


121

Influence of size and surface coating on uptake of silver and gold nanoparticles and the

molecular effects on Gammarus fossarum (Crustacea Amphipoda)

Kahina Mehennaoui a,b, Sébastien Cambier a, Tommaso Serchi a, François Guérold b, Johanna Ziebel

a, Jean-Sebastien Thomann c, Nathalie Valle c, Laure Giambérini b, *, Arno C. Gutleb a*



B Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360,

Université de Lorraine – Metz, France

C Material Research and Technology (MRT) Department, Luxembourg Institute of Science and


* Corresponding authors: Arno C. Gutleb

Postal Address: Environmental Research and Innovation Department

Luxembourg Institute of Science and Technology

5, avenue des Hauts-Fourneaux, L-4362 Esch-sur-Alzette, Luxembourg

E-mail : [email protected]

Laure Giamberini

Postal Address : Université de Lorraine, Laboratoire Interdisciplinaire des Environnements

Continentaux (LIEC, CNRS UMR 7360)

Campus Bridoux, rue du General Delestraint, F-57070 Metz

E-mail : [email protected]


122

ABSTRACT

The specific and unique properties of silver nanoparticles (AgNPs) and gold nanoparticles (AuNPs), make

them of high interest for different scientific and industrial applications. Their increasing use will inevitably

lead to their release in the environment and aquatic ecosystems where they may represent a threat to

aquatic organisms. Being a widespread and important component of the aquatic macroinvertebrate

assemblage, amphipods and more specifically Gammarus fossarum will certainly be exposed to AgNPs and

AuNPs. For these reasons, G. fossarum was selected as model organism for this study.

The aim of the present work was the evaluation of the influence of both size (20, 40 and 80 nm) and surface

coating (citrate CIT, polyethylene glycol PEG) on the acute toxicity of AgNPs and AuNPs on G. fossarum. We

investigated the effects of AgNPs and AuNPs on the uptake by G. fossarum, NP tissue distribution and the

expression of stress related genes by the use of ICP-MS NanoSIMS 50 and Cytoviva, and RT-qPCR,

respectively.

Ag and Au bioaccumulation revealed a significant surface-coating dependence, with CIT-AgNPs and CIT-

AuNPs showing the higher bio-accumulation potential in G. fossarum as compared to PEG-NPs. Opposite to

that, no size-dependent effects on the bioaccumulation potential were observed. NanoSIMS 50 and CytoViva

revealed an influence of the type of metal on the tissue distribution after uptake, with AgNPs detected in the

cuticle and the gills of G. fossarum, while AuNPs were detected in the gut area. Furthermore, AgNPs were

found to up-regulate MnSOD and CuZnSOD gene expression while AuNPs led to their down-regulation.

Modulation of SODs may indicate generation of reactive species of oxygen and a possible activation of

antioxidant defence in order to prevent and defend the organism from oxidative stress. However, further

investigations are still needed to better define the mechanisms underlying the observed AgNPs and AuNPs

effects.

Keywords: Gammarus fossarum, Silver nanoparticles, Gold nanoparticles, bioaccumulation, gene expression,

NanoSIMS 50, Cytoviva®


123

1 INTRODUCTION

Engineered nanoparticles (ENPs) are being more and more incorporated in consumers’ products such as

cosmetics, paints, food, computers, medicine, etc. (McGillicuddy et al., 2017; Vale et al., 2016). As a

consequence, their environmental release is raising concerns about their potential toxicity/effects on human

and other living organisms (Klaine et al., 2008; Moreno-Garrido et al., 2015). Among all ENPs, silver

nanoparticles (AgNPs) are the most commercialized and the most used ones in consumer products; such as

clothing, health-care products, food packaging, etc. (Fabrega et al., 2011; Vale et al., 2016) mostly due to

their well-documented antibacterial properties (Bone et al., 2012; Cleveland et al., 2012; McGillicuddy et al.,

2017; Zhang et al., 2016). Gold nanoparticles (AuNPs) represent another example of widely used

nanomaterials. They are mostly used for medical and molecular biology applications mainly for their

supposedly low toxicity and high bio-compatibility (García-Negrete et al., 2013; Lapresta-Fernández et al.,

2012).

The behaviour and fate of AgNPs and AuNPs in aquatic ecosystems and their toxicity towards aquatic

organisms are influenced by their physical-chemical properties such as size and surface chemistry (Angel et

al., 2013; Ellis et al., 2016; Makama et al., 2016). The surface of NPs is commonly modified by addition of

capping agents of surface functionalization in order to prevent aggregation and ion release (Gao et al., 2012;

Levard et al., 2012; Park et al., 2015a; Tolaymat et al., 2010). Common capping agents are citrate (CIT), which

is applied as stabilizer by charge repulsion (Tejamaya et al., 2012), and polyethylene glycol (PEG), which is

used as a sterically stabilizer and described to strongly bind to the core of NPs (Cumberland and Lead, 2009;

Römer et al., 2011; Tejamaya et al., 2012). Although surface-modified AgNPs and AuNPs are widely used, the

influence of these compounds on the behaviour, uptake and toxicity of AgNPs and AuNPs on aquatic

organisms is still not fully understood (Croteau et al., 2011; Kwok et al., 2012; Park et al., 2015).

Amphipods of the genus Gammarus such as Gammarus fossarum represent an important part of the aquatic

macroinvertebrates assemblage (Dedourge-Geffard et al., 2009; Ladewig et al., 2006). Recently, G. fossarum

have been used as a model organism due to their wide distribution in Europe (Janetzky, 1994) and their major

functional role in litter breakdown process and nutrient cycling (Forrow and Maltby, 2000; Kelly et al., 2002;

Lacaze et al., 2010; MacNeil et al., 1997). The ease of use, identification to the species level, differentiation

between gender, sampling and laboratory handling in addition to the well-documented sensitivity to different

kind of pollutants make Gammarus good model organisms for ecotoxicological studies (Arce Funck et al.,

2013; Mehennaoui et al., 2016). Gammarids have been recognized as a relevant tool for water quality and

risk assessment as they allow the assessment of their internal contamination level that is always dependent

and proportional to the amount of contaminants introduced in the environment (Kunz et al., 2010; Vellinger

et al., 2012a).


124

The aim of the present study was to investigate both the influence of coatings (CIT and PEG) and size (20, 40

and 80 nm) on the uptake and molecular effects of AgNPs and AuNPs on the aquatic amphipod G. fossarum.

Effects of AgNO3 were also studied, as a positive control, in order to determine the contribution of the

released ions to the observed biological effects. The biological endpoints that were measured in this study

include mortality, bioaccumulation, localization of AgNPs and AuNPs in G. fossarum tissues and expression

of stress-related genes. Total and ionic Ag and Au were measured in the exposure media in order to

determine the behaviour and aging of AgNPs and AuNPs.

2 MATERIALS AND METHODS

2.1 Particles and Chemicals

Citrate stabilized Ag nanoparticles (CIT-AgNPs 20, 40 and 80 nm) and citrate stabilized Au nanoparticles (CIT-

AuNPs 20, 40 and 80 nm) were purchased from Nanocomposix (San Diego, USA). Coating of AgNPs and AuNPs

with PEG was performed at the LIST’s Material Research and Technology (MRT) Department, (Luxembourg

Institute of Science and Technology (LIST), Luxembourg). AgNO3 (CAS no.7761-88-8) was purchased from

VWR (Leuven, Belgium). Osmium tetroxide (OsO4, CAS no.20816-12-0), glutaraldehyde (CAS no.111-30-8) and

the Spurr low viscosity resin embedding kit were purchased from Sigma Aldrich (Bornem, Belgium).

Phosphate buffer saline (PBS) was obtained from Invitrogen (Merelbeke, Belgium).

2.2 Particle characterization

CIT-AgNPs and CIT-AuNPs were characterized by the manufacturer and the information provided include TEM

images (JEOL1010 Transmission Electron Microscope), mass concentration (Thermo Fisher X series 2 ICP-MS),

Hydrodynamic diameter and zeta potential (Malvern nanoZetasizer), UV-visible extinction spectrum and pH

were provided by Nanocomposix

(https://tools.nanocomposix.com:48/cdn/coa/Silver/Spheres/BioPure/AG20-SS-BP-CIT-

SCM0069.pdf?202%20903).

Functionalization of AgNPs and AuNPs surface was performed at the Material Science and Technology

department (MRT) in the Luxembourg Institute of Science and Technology (LIST). AgNPs and AuNPs were

coated with PEG (MW layer 3000) following a wet-chemistry (Thiol/metal chemistry) method that allowed

functionalization of each isolated NPs. AgNPs and AuNPs coatings were controlled using Scanning Electron

Microscopy (SEM, Figure S1 A) and size distribution in MilliQ water was checked using Nanoparticle Tracking

Analysis (NTA, measured on a NanoSIGHT instrument, Malvern Instrument Ltd, UK(Hole et al., 2013)), (Figure

S1 B).


125

In addition to the information provided by the manufacturer, we performed complementary characterization

including size distribution and zeta potential measurement. Size distribution of CIT-AgNPs, PEG-AgNPs, CIT-

AuNPs and PEG-AuNPs in Volvic water (exposure medium), at T0h and T24h, were characterized using NTA.

Zeta potential in exposure medium was analysed using NanoZetasizer (Malvern Instrument Ltd, UK).

2.3 Organism sampling and acclimation

G. fossarum were collected in June 2015 (CIT-AgNPs and PEG-AgNPs exposure), September 2015 (CIT-AuNPs

exposure) and October 2015 (PEG-AuNPs exposure) at an unpolluted stream (49°48’24.9’’ N and 06°04’53.2’’

E, Schwaarzbaach, Colmar-berg, Luxembourg) (Dohet et al., 2008; Mehennaoui et al., 2016). Animals were

collected using a hand net and were sorted in the field to separate males, which were used for the

experiments, from females, which were placed back in the river. G. fossarum males were selected from pre-

copula pairs or based on sexual dimorphism like gnatopode size (Mehennaoui et al., 2016). Adult males were

kept for the experiments in order to avoid influence of gender on the studied parameters (Arce Funck et al.,

2013; Sornom et al., 2010). Collected male animals were immediately brought in river water to the laboratory

and were kept at 12 °C. Adult males were then acclimated to laboratory conditions (Andreï et al., 2016;

Mehennaoui et al., 2016). The acclimation was conducted in 2 steps. First, Gammarids were acclimated for

72h to mineral water (Volvic, France) by progressively changing field water to Volvic water (30% v/v, 50% v/v,

100% v/v). Then, a stalling period of 10 days was conducted in 100% Volvic water (Andreï et al., 2016;

Mehennaoui et al., 2016). The acclimation was performed under controlled conditions at 12 °C with a 16h

light 8h dark photoperiod. Volvic water was aerated and changed every 24h to avoid organic matter

accumulation and potential increase of ammonium, nitrite and nitrate. Gammarids were fed ad libitum with

alder leaves (Alnus glutinosa) up to 24h before experiment.

2.4 Acute toxicity test

In order to avoid as much as possible the adsorption of AgNO3, AgNPs and AuNPs during the exposure, the

tanks used for exposure were saturated for 96h before exposure (Andreï et al., 2016).

At the end of the acclimation period, groups consisting of 6 males each were transferred into plastic tanks

(250 ml polypropylene tanks) containing 70 ml of exposure medium (Volvic water with or without

contaminants) (Mehennaoui et al., 2016). A piece of mesh was added in each tank to provide a resting surface

for G. fossarum and to reduce as much as possible potential losses linked to cannibalism. Males were exposed

to different treatments: AgNO3 (0.5 – 1 – 2 – 4 – 8 µg.L-1); CIT-AgNPs and PEG-AgNPs 20 nm (1 – 2 -4 – 8 – 10

µg.L-1), CIT-AgNPs 40 nm, PEG-AgNPs 40 nm, CIT-AgNPs 80 nm, PEG-AgNPs 80 nm, CIT-AuNPs and PEG-AuNPs

20, 40 and 80 nm (1 – 5 – 10 – 25 – 50 µg.L-1), and a control for 72h at 12 °C with a photoperiod of 16h light


126

and 8h darkness and in absence of feeding. The different ranges of concentrations tested were selected based

on previous results obtained by exposing D. magna to the same silver and gold nanoparticles (Table S1).

AgNO3, AgNPs and AuNPs stock solutions were diluted to the desired concentrations in mineral water (Volvic,

France). 100% of exposure medium were renewed daily. Dead animals were removed and survivors were

counted.

2.4.1 Total and dissolved silver and gold measurements

At the end of exposure period, 4 mL of exposure medium were analysed for ions release measurements.

Medium were centrifuged at 4000g for 40 min using a 3KDa cut off membrane (Millipore) followed by

mineralization step with 1% HNO3 for Ag ions measurements and 1% HCl for Au ions measurements.

Total Ag and Au concentrations were measured in the exposure medium sampled from the water column at

the end of the exposure period. 5 mL of exposure medium/replicate were mineralized using 1% (v/v) of HNO3

and HCl for Ag and Au measurements, respectively. All measurements of total and dissolved silver and gold

were performed with Inductively Coupled Plasma Mass spectrometer (ICP-MS) (ElanDRC-e, Perkin-

ElmerWaltham, MA, USA) as previously published (Mehennaoui et al., 2016). All measurements were

performed in the absence of animals. The quantification limit of ICP-MS for Ag was around 0.051 µg. L-1 and

0.255 µg. L-1 for Au. A calibration curve was performed following serial dilutions (from 0.01 to 50 µg. L-1 of Ag

and from to for Au) using a multi-element standard solution (Chemlab, certified ISO/IEC 17025). The validity

of the analytical method was checked every 10 measurements within the same series of measurements with

3 quality controls (0.5, 5 and 25 µg.L-1 of Ag and Au) of a second multi-element standard solution (Merck-

VWR, ISO/IEC 17025); Ag and Au values were consistent within the certified ranges (Cambier et al., 2018).

2.4.2 Silver and gold bioaccumulation

At the end of the exposure period, three pools of 4 gammarids each per conditions were rinsed with milliQ

water and dried on filter paper. Animals were frozen at -20 °C, freeze-dried and weighted. Samples were then

mineralized in HNO3 and H2O2 for Ag measurement and in acidic mixture of HNO3 and HCl for Au

measurement at a maximum pressure of 35 bars and maximum temperature of 200 °C using a microwave

(Anton Paar Multiwave Pro) as previously described (Mehennaoui et al., 2016). Results are expressed in µg.g-

1 of dry weight. All measurements were performed in triplicates. A calibration curve was performed following

serial dilutions (from 0.01 to 50 µg.L-1 of Ag and from 0.025 to 10 µg.L-1 for Au). Using a multi-element standard

solution (Chemlab, certified ISO/IEC 17025). The validity of the analytical method was checked every 10

measurements within the same series of measurements with 3 quality controls (0.5, 5 and 25 µg.L-1 of Ag and


127

Au) of a second multi-element standard solution (Merck-VWR, ISO/IEC 17025); Ag and Au values were

consistent within the certified ranges (Cambier et al., 2018).

2.4.3 Nanoparticle uptake

2.4.3.1 Sample preparation

At the end of the exposure period, G. fossarum exposed to 10 µg.L-1 of AgNPs and AuNPs were fixed in 5%

(v/v) glutaraldehyde where they were kept at 4 °C until analyses. After a washing step with PBS, samples

were post-fixed with 1% osmium tetroxide for 2h at room temperature on a horizontal shaker (180 rpm).

After a second washing with PBS, samples were dehydrated with increasing concentrations of ethanol (10%,

20%, 30%, 50%, 70% 90% and 100% v/v) and a final wash in acetone (100% v/v) before being embedded in

Spurr resin for 16h at 60 °C.

2.4.3.2 NanoSIMS50 analyses

Semi-thin layers of 300 nm were cut using ultra-microtome and placed on silicon wafers (Siltronix, Archamps,

France) for Secondary Ion Mass Spectrometry (SIMS) analysis (Georgantzopoulou et al., 2013; Mehennaoui

et al., 2016). Samples were analysed with a NanoSIMS 50 (Cameca, Gennevilliers, France) using Cs+ as primary

ion source (8kev) sputtering the surface of the sample (-8 kev) with a raster of 40 x 40 μm2 to generate

secondary negative ions for silver analyses. Images were recorded in a pixel format of 256 x 256 image points

with a counting time of 30 ms per pixel. The instrument was tuned for a mass resolution (M/ΔM) up to 5000

for the elimination of atomic or molecular isobar interference. The mass calibration of the silver ion was

realized using a silver foil (Goodfelow, Huntingdon, UK). The isotopic ratio between 107Ag- (m= 106.9051,

51.8%) and 109Ag- (m=108.9048, 48.2%) was measured and verified (ratio = 1,074). The mass calibration of

gold ions 197Au- (m= 196.966543 amu) was realized using a gold leaf. The 12C14N- cluster was simultaneously

detected with the silver ions to allow the recognition of the essential anatomical features (Eybe et al., 2009;

Georgantzopoulou et al., 2013).

2.4.3.3 Cytoviva® dark field hyperspectral imaging

Semi-thin layers of 300 nm were cut using ultra-microtome and placed on microscope glass slide and let to

dray for 1 min on a drying workstation. Slides were covered with a glass cover slip, sealed, and kept at 4 °C

until analysis. Visualization was performed using Cytoviva® hyperspectral imaging system (Cytoviva Inc.,

Auburn, Alabama, USA) mounted on Olympus BX-43 optical microscope. Images of the different tissues of G.

fossarum were captured at 60x oil immersion magnification using hyperspectral camera controlled by

environment for visualization ENVI software (version 4.8 from Harris Corporation, Melbourne, FL, USA and


128

modified by CytoViva, Inc.). Spectral libraries of exposed animals were generated manually acquiring about

200 spectra per samples. Acquired libraries were filtered against non-exposed samples to filter out all spectra

non-related to AgNPs or AuNPs using a spectral angle mapper (SAM) algorithm with a 0.05 radians tolerance.

Filtered libraries were mapped onto images of exposed samples using SAM with a 0.05 radian tolerance which

allows highlighting similarities between the spectra in the image and in the spectral library.

2.4.4 Molecular responses

2.4.4.1 RNA extraction and cDNA synthesis

At the end of the exposure period, animals were gently dried and dived in RNA lysis Buffer (RLT buffer)

supplemented with 1% (v/v) β-mercaptoethanol, flash-frozen in liquid nitrogen and stored at -80 °C until RNA

extraction. For RNA extraction, G. fossarum tissues were grinded on ice using a pellet pestle tissue grinder

and the homogenates were centrifuged at 250g for 5 min at 4 °C to remove cuticle fragments as previously

described (Mehennaoui et al., 2016). Total RNA was extracted using RNeasy mini kit for cells and animal

tissues (Qiagen, Leusden, The Netherlands), according to the manufacturer’s instructions (including digestion

with DNase). RNA concentrations and purity were assessed measuring the absorbance at 230, 260 and 280

nm using Nanodrop ND-100 spectrophotometer (ThermoScientific, Villebon-sur-Yvette, France) (A260/280

and A260/230 ratios). RNA integrity was checked using the RNA Nano 6000 assay (Agilent Technologies,

Diegem, Belgium) with a 2100 Bioanalyzer (Agilent, Santa Clara, CA, USA) (Legay et al., 2015). For the

synthesis of cDNA, 1 µg of the extracted RNA were retro-transcribed into cDNA using the Protoscript II reverse

transcriptase (New England Biolabs, Leiden, The Netherlands) and random primers, according to the

manufacturer’s instructions.

2.4.4.2 Primer design and quantitative real-time PCR

All the primers for the RT-PCR were designed using Primer3Plus (http://primer3.ut.ee) with the following

criteria: primer size between 18 and 25 base pairs, GC content between 30% and 70 %, amplicon size from

80 to 150 base pairs, primer annealing temperatures in the 57-61 ºC range. Primers were analysed using

NetPrimer (http://www.premierbiosoft.com/netprimer/) for secondary unexpected structures. PCR

efficiency was checked using decreasing five-fold dilutions from cDNA pool (From 25 ng to 0.04 ng and no

template control). Melting curves were generated at the end of each experiment to assess the specificity of

the amplified products. All primers displayed one clear peak and were all retained for the analysis. All the

used primers are described in Table 4. 1

cDNAs were diluted to 0.8 ng. µL-1 and used for RT-qPCR analyses in 384-well plates. An automated liquid

handling robot (epMotion 5073, Eppendorf, Hamburg, Germany) was used for the preparation of the 384-


129

well plate. cDNA were amplified using Takyon Low ROX SYBR MasterMix dTTP Blue Kit (Eurogentec, Liège,

Belgium) on a ViiA 7 Real-Time PCR System (Thermo Fisher, Waltham, MA, USA) in a final volume of 10 µL

(Behr et al., 2015). All reactions were performed in three technical replicates each one including four

biological replicates. The PCR conditions consisted on an initial denaturation step at 95°C for 3 min, followed

by 40 cycles of denaturation at 95°C for 15 sec and annealing/extension steps at 60°C for 60 sec. Three genes,

namely Ca2+ATPase, Clathrin and SDH, were selected among seven genes to be used as reference and they

were validated using the GeNorm module in the Biogazelle qBase PLUS software (Mehennaoui et al., subm.

Chapter 3). The relative gene expression was calculated including PCR efficiency using Biogazelle qbase Plus

software 2.5 with the ΔΔCT method.


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Table 4. 1. Specific primer pairs used for RT-qPCR analyses on Gammarus fossarum exposed for 72h to AgNPs and AuNPs (coated with CIT or PEG, sizes: 20, 40 and 80

nm, F: forward sequence, R: reverse sequence)

Gene code Description Primer sequence (5’ – 3’) Amplicon size (bp) PCR efficiency

Actin Actin F: CTCATGCTATCCTTCGTCTTGA

103 2.02 R: CGTTCAGCGGTGGTTACAA

Tub Tubulin F: CGGCTGTTGTTGAACCTTAC

93 2.09 R: AGATGGCCTCATTGTCAACC

GAPDH glyceraldehyde 3-phosphate dehydrogenase F: GTCCGTCTCGCTAAGGAGTG

94 1.91 R: TGTATCCGAGGTAGCCCTTG

SDH succinate dehydrogenase F: GGAAGAAGCTGGATGGTCTG

87 1.98 R: ACTTGTCTCCGTTCCACCAG

UB Ubiquitin F: CCCACGATACTCCCTTTGAA

82 2.01 R: ACAATCGGTGGCTTGTTAGG

Ca2+ATPase Calcium ATPase F: CTACGCCAATCAGTTCGTCA

87 2.03 R: TCACAGCGAGAGGAAGACCT

Clathrin Clathrin F: ATCGCCAAGCTTTGTGACTT

107 1.99 R: GCTTTGATAGGCGGACTCTG

MnSOD manganese superoxide dismutase F: AAACGCGACTTTGGAAGTGT

150 1.99 R: TTCTAGCGGATCCTGATTGG

CuZnSOD copper zinc superoxide dismutase F: TTGCCTTGTTGTTGTTGGTG

95 1.92 R: GGCCGTGTTGATGTAGAGGT

P53 Tumour protein P53 F: ACGAGCTCTCCTTCAAGTGC

104 R: CTTGGAAGGTATCCAGTGCAG

HSP90 heat shock protein 90 F: CTGGTTTCTTCTCCCTGCTG

135 1.99 R: GATCTCGAGGTGCTTCTTGG

Gadd45 Growth arrest and DNA damage 45 F: CACACGACGACAAGTTCCTG

91 1.92 R: TGCTGCTTCAATTGCTTCTT

GPx7 glutathione peroxidase 7 F: AGCGTGCTAGTCCAGCTTTC

106 1.95 R: GCATTGATCACGACACCATC


131

HEM hemocyanin F: GGAGAGGATATCGGCATGAA

124 1.94 R: GGTGATGAGCCCAGAAGAAA

CAT catalase F: CTCGTCAACAAGGACAATGC

130 1.97 R: GGTCTCTGATGGCGTAGTCG

CathL Cathepsin L F: GGTTCACTCGAAGGTCAACAC

83 1.95 R: TTGTCTGAACAATCGACCAAG

Cjun C-Jun F: GATCGGTCGTTTGGAAGAAA

94 1.95 R: TGCACACACTTGATCACGTAAC

Cox1 Cyclooxygenase 1 F: GGGAATCAATTTCATTACTACTTGTT

89 1.95 R: TTATTGCCCAAACGAAGAGG

GST Glutathione S transferase F: TGCCTCTAGGTCAACTTCCAG

80 1.98 R: AGCAACATAACGAGCAATCG

NaKATPase Sodium potassium ATPase F: ATTCGCTTGGCTGTGAGG

85 2.02 R: TCCTCATCGGTATCATCGTG

NfkB nuclear factor kappa B F: GCAGAGCATCGACTTGAATG

101 2.09 R: GCGAGACGACAGGAGTAAGG

Chitinase Chitinase F: CTCGTCAACAAGGACAATGC

130 1.96 R: GGTCTCTGATGGCGTAGTCG


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2.5 Statistical analyses

LC50 values were calculated on AgNO3 nominal concentrations based on a logistic curve-fitting procedure

applying the method described earlier (Isnard et al., 2001; Vindimian et al., 1999). The Excel macro REGTOX

allowed us to estimate both LC50 values using Hill model (Vindimian et al., 1983) and their 95% confidence

intervals (Vellinger et al., 2012b).

Bioaccumulation results are reported as mean ± SD (standard deviation) and relative gene expression results

are reported as mean ± SE (standard error). The observed Ag and Au bioaccumulation were compared using

two-way ANOVA followed by Tukey-HSD post-hoc test for multiple comparisons. The weighed means were

calculated to identify in one hand the “coating effects” when results of all sizes were combined and in the

other hand, the “size effects” were compared while the coatings were combined.

Effects on molecular responses were analysed on log2 transformed normalized relative quantities. Two-way

ANOVA followed by a Fisher LSD post hoc test was performed. All statistical calculations were performed

using Statistica software 12 (StatSoft Inc.).

3 RESULTS

3.1 Nanoparticle characterization

Analysis of the hydrodynamic diameter of NPs by NTA, showed that both PEG-AgNPs and PEG-AuNPs

were generally more stable in Volvic water, both at T0h and T24h, than CIT-AgNPs and CIT-AuNPs (Table 4. 2

and Figure S2). After 24h in exposure medium, CIT-AgNPs 20 nm appeared to be the most aggregating particle

with a size of 53 ± 3.7 nm, while CIT-AgNPs 40 and 80 nm appeared to be more stable, showing hydrodynamic

diameter of 45 ±1.5 nm and 72 ± 2.9 nm, respectively. Similarly, CIT-AuNPs 20 and 40 nm tented to aggregate

after 24h in exposure medium with CIT-AuNPs 20 nm being the one that aggregate the most with a size of

154 ± 28 nm. CIT-AuNPs 40 nm and CIT-AuNPs 80 nm had sizes of 67 ± 12.8 nm and 85 ± 0.3 nm, respectively.

Zeta potentials were ranging from -4.8 to -19.1 mV for all the studied AgNPs and AuNPs with CIT-NPs

presenting more negative charges at their surface than PEG-NPs (Table 4. 2).


133

Table 4. 2. Size distribution of particles (mode ± SD, 3 replicates) and ζ potential of AgNPs and AuNPs in Volvic

water (exposure medium) at T0h and T24h.

3.2 Acute toxicity test

3.2.1 Survival

As expected, AgNO3 was the most toxic compound. A significant seasonal variability was observed with LC50

values ranging from 0.78 µg. L-1 (0.001 – 0.85) in autumn (November 2015) to 4.31 µg.L-1 (3.88 – 4.75) in

summer (June 2015) (Figure S3). None of the tested NPs, irrespective of the core material (Ag or Au), coating

or size, induced increase of mortality at any of the tested concentrations (from 1 to 50 µg. L-1).

3.2.2 AgNP exposure

3.2.2.1 Total and dissolved [Ag]

At the end of the experiment, exposure medium was analyzed by ICP-MS to quantify the total concentration

of total silver. Total Ag measured for control groups were under the limit of quantification (< 0.051 µg. L-1,

Table 4.3). Total Ag recovered from CIT-AgNPs 20 nm was higher than Ag recovered from PEG-AgNPs 20 nm

(Table 4.3) while a higher recovery of total Ag introduced as PEG-AgNPs 40 nm and PEG-AgNPs 80 nm was

observed compared to CIT-AgNPs 40 and 80 nm. The recovery rates of total Ag from AgNO3 was in the same

concentration range than PEG-AgNPs 40 and 80 nm (Table 4.3).

Concentrations of dissolved Ag ions from AgNPs were detectable (3.39 ± 0.4 µg. L-1 of Ag only at the highest

concentrations of PEG-AgNPs 40 nm (50 µg.L-1), while in all other treatments the amount of released Ag ions

were below the limit of detection (< 0.051 µg.L-1).

Particles Volvicwater(T0h) Volvicwater(T24h)

Coating Nominalsize(nm) Size(nm) ζpotential(mV) Size(nm) ζpotential(mV)

AgNPs

Citrate

20 61±2.1 -9.8 53±0.7 -19.1

40 41±0.3 -13.6 45±1.5 -7.2

80 78±2.9 -16.8 72±2.9 -18.6

PEG

20 22±0.7 -2.8 20±1.0 -5.3

40 43±1.2 -6.0 47±1.3 -7.2

80 77±1.2 -6.1 75±2.1 -8.6

AuNPs

Citrate

20 42±1.5 -6.0 154±28 -17.1

40 38±0.9 -14.1 67±12.8 -19

80 75±1.0 -14.3 85±0.3 -19

PEG

20 39±1.5 -0.1 40±2.5 -8.8

40 59±1.2 -6.7 38±19.2 -4.8

80 89±0.9 -11.8 86±1.5 -9.2


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Table 4. 3. Total ag concentrations (mean ± SD) and recovery rates (mean ± SD) of AgNO3, CIT-AgNPs and PEG-

AgNPs in Volvic water (exposure medium).

3.2.2.2 [Ag] in Body tissue

Small but detectable amounts of Ag could be detected in the negative control groups of G. fossarum, for

which the observed Ag body concentration was 0.27 ± 0.02 µg. g-1 dry weight (Figure 4. 1). Treatment with

AgNO3 (4 µg. L-1) lead to a significant increase of the Ag body concentrations that were of 7.56 ± 1.89 µg.g-1

d.w. Similarly, treatment with the AgNPs resulted in higher body concentrations of silver in treated animals.

A significant effect of “coating”, “size” and the interaction “coating x size” was observed on the Ag

concentrations in body tissues of G. fossarum (Figure 4. 1 A, Two-way ANOVA, P < 0.05) but the effects were

stronger for the coatings (Figure 4. 1, A, B; Two-way ANOVA, F = 237.3, P < 0.05) than for the size (Figure 4.

1A, C; Two-way ANOVA, F = 1721.4, P < 0.05).

0 <LOQ N.D

0.5 0.20±0.01 39.54±2.78

1 0.45±0.08 44.71±7.91

2 0.91±0.09 45.40±4.38

4 2.76±0.29 68.91±7.29

8 4.92±0.14 61.56±1.77

1 0.80±0.02 80.48±2.08

2 1.68±0.07 84.11±3.67

4 3.22±0.18 80.44±4.38

8 5.91±0.57 73.91±7.16

10 8.70±0.20 87.04±1.98

1 0.47±0.04 46.87±3.83

2 1.10±0.35 54.82±17.56

4 1.94±0.06 48.87±3.83

8 5.17±0.59 64.57±7.33

10 6.97±0.29 69.66±2.86

1 0.49±0.04 48.67±3.91

5 3.21±0.25 64.10±7.91

10 6.79±0.90 67.91±9.04

25 12.58±1.28 50.33±5.12

50 20.26±2.89 40.52±5.77

1 0.97±0.20 97.36±20.15

5 4.33±0.08 86.65±1.51

10 9.01±0.46 90.14±4.57

25 14.67±1.42 56.68±5.66

50 27.60±2.58 55.21±5.16

1 0.77±0.02 77.00±2.34

5 4.37±1.32 87.49±26.42

10 7.21±0.69 72.07±6.93

25 12.17±1.22 48.70±4.86

50 20.38±1.76 40.77±3.51

1 1.10±0.07 >100

5 5.46±0.14 >100

10 10.34±0.55 >100

25 23.68±1.11 94.71±4.45

50 24.43±6.36 48.86±12.72

AgNPs80nm

CIT

PEG

Compounds Nominalconcentrations(µg.L-1)

Control

AgNO3

AgNPs20nm

CIT

PEG

Totalsilverinwater(µg.L-1) Recovery(%)

AgNPs40nm

CIT

PEG


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Figure 4. 1. G. fossarum Ag bioconcentration (mean ± SD) after 72h exposure to CIT and PEG-AgNPs 20, 40 and

80 nm.

(A) Coating x size: mean Ag bioconcentration for each tested coating (CIT vs PEG) and sizes (20, 40 and 80 nm).

Vertical bars represent standard deviation

(B) Coating effect: Weighted mean Ag bioconcentration for each tested coating (results for all sizes tested were

combined). Vertical bars represent 0.95 confidence interval

(C) Size effect: Weighted mean Ag bioconcentration for each size tested (results for all coatings tested were

combined). Vertical bars represent 0.95 confidence interval. Different letters (from a to l) represent the significant

differences in mean Ag bioconcentration values (Two-way ANOVA + Tukey-HSD post hoc test at P < 0.05 level of

significance)

A significant concentration-dependent Ag bioconcentration was observed for CIT-AgNPs. The highest

concentrations (10 µg. L-1) of CIT-AgNPs 20 nm and (25 and 50 µg. L-1) of CIT-AgNPs 40 nm and CIT-AgNPs 80

nm led to a significant higher Ag bioconcentration (Figure 4.1, A, C). On the contrary, no significant

concentration-dependent Ag bioconcentration was observed for PEG-AgNPs 20 nm and PEG-AgNPs 80 nm.

Only the highest concentrations (25 and 50 µg. L-1) of PEG-AgNPs 40 nm led to a significant higher Ag

bioconcentration (Figure 4. 1A). At the same Ag concentrations (≈ 5 µg. L-1), G. fossarum exposed to AgNO3

accumulated more silver than those exposed to AgNPs (Figure 4. 1 A, C).

Control

AgN

O30.5

AgN

O31

AgN

O32

AgN

O34

AgN

O38

AgN

Ps201

AgN

Ps202

AgN

Ps204

AgN

Ps208

AgN

Ps2010

AgN

Ps401

AgN

Ps405

AgN

Ps4010

AgN

Ps4025

AgN

Ps4050

AgN

Ps801

AgN

Ps805

AgN

Ps8010

AgN

Ps8025

AgN

Ps8050

5

10

15

20

25

30

0

35

Size(nm)

1

2

3

4

5

6

0

Citrate PEG

AgBioconcentration(µg.gdw

-1)

Coatings

A

B C


136

3.2.3 AuNP exposure

3.2.3.1 Total and dissolved [Au]

Total gold concentrations in the control tanks were under the limit of detection (< 0.255 µg. L-1). The

concentration of AuNPs in exposure medium were close to the limit of detection leading to missing values

for several conditions and limiting the possibility to exploit the data.

3.2.3.2 [Au] in body tissue

The Au bio-concentration was under the limit of quantification (< 0.075 µg. g-1 d.w) in the non-exposed

animals. Treatment with the AuNPs resulted in higher body concentrations of Au in treated animals. A

significant effect of “coating”, “size” and the interaction “coating x size” was observed (Figure 4.2, Two-way

ANOVA, P < 0.05). For CIT-AuNPs, a significant dose-dependent effect in bio-accumulation is observable, with

higher doses leading to higher body concentrations (Figure 4.2, A B, Two-way ANOVA, F = 669.28, P < 0.05)

while a trend to increasing Au internal concentrations in a dose-dependent manner is also observed for PEG-

AuNPs (Figure 4.2, A,B). On the contrary, no clear influence of NPs size on bio-accumulation is observable.

(Figure 4.2 , A, C).

Legend next page

Citrate PEG

1

2

3

4

5

6

0

7

-1

Coatings

Size(nm)

0

5

10

15

20

25

30

0 0.5 1 2 4 8 1 5 10 25 50 1 5 10 25 50 1 5 10 25 50 1 5 10 25 50 1 5 10 25 50 1 5 10 25 50

Citrate PEG Citrate PEG Citrate PEG

control AgNO3 AuNPs20nm AuNPs40nm AuNPs80nm

Exposureconditions (µg.L-1)

a ababababcabcabc abc abcabc abcd

abcdeabcde

abcde abcdebcdef

cdef def

fggh

gh

i

i

j

ef

h

abcde

AuBioconcentration(µg.gdw

-1)

Control

AgN

O30.5

AgN

O31

AgN

O32

AgN

O34

AgN

O38

AuNPs201

AuNPs205

AuNPs2010

AuNPs2025

AuNPs2050

AuNPs401

AuNPs405

AuNPs4010

AuNPs4025

AuNPs4050

AuNPs801

AuNPs805

AuNPs8010

AuNPs8025

AuNPs8050

5

10

15

20

25

30

0

A

B C


137

Figure 4. 2. G. fossarum au bioconcentration (mean ± SD) after 72h exposure to CIT and PEG AuNPs 20, 40 and

80 nm.

(A) Coating x size: mean Au bioconcentration for each tested coating (CIT vs PEG) and sizes (20, 40 and 80 nm).

Vertical bars represent standard deviation

(B) Coating effect: Weighted mean Au bioconcentration for each tested coating (results for all sizes tested were

combined). Vertical bars represent 0.95 confidence interval

(C) Size effect: Weighted mean Au bioconcentration for each size tested (results for all coatings tested were

combined). Vertical bars represent 0.95 confidence interval. Different letters (from a to j) represent the significant

differences in mean Au bioconcentration values (Two-way ANOVA + Tukey-HSD post hoc test at P < 0.05 level of

significance)

3.2.4 Particle uptake

3.2.4.1 NanoSIMS50 analyses

Optical microscope images (OM) of the cross section of G. fossarum are presented in Figure 4. 3 A and Figure

4. 4 A. CN pictures are used to identify the anatomical features of G. fossarum (Figure 4. 3 B). Ag could be

detectable in gills of G. fossarum exposed to AgNPs 40 nm (Figure 4. 3). A higher signal of Ag was found in

gills of G. fossarum exposed to CIT-AgNPs 40 nm, more specifically in the cuticle (Figure 4. 3). A lower Ag

signal was found in gills of G. fossarum exposed to PEG-AgNPs 40 nm whereas control animals contained

considerably less Ag (Figure 4. 3).

Figure 4. 3. A) 300 nm cross section of G. fossarum observed under optical microscope. Red square represents

the area observed with NanoSIMS 50. Scale bar is 250 µm. B) Elemental distribution of 12C14n- clusters. C) 109Ag-

ions in 300 nm cross sections of G. fossarum gills. Animals were exposed to 10 µg. L-1 of CIT-AgNPs 40 nm and

PEG-AgNPs 40 nm in Volvic water. Scale bar is 2 µm.

Cit-AgNPs

40 nm

PEG-AgNPs

40 nm

107Ag

CN- 107Ag

Control

CN- 107Ag

CN-

A B C


138

Au could only be found close to intestinal caeca of G. fossarum exposed to CIT-AuNPs 40 nm (Figure 4. 4 C)

more specifically between the external wall of two intestinal caeca. No Au signal could be found in G.

fossarum exposed to PEG-AuNPs or in control animals (Figure 4. 4).


the area observed with NanoSIMS 50. Scale bar is 250 µm. B) Elemental distribution of 12C14N- clusters. C) 109Au-

ions in 300 nm cross sections of G. fossarum intestinal caeca. Animals were exposed for 72h to 10 µg. L-1 of CIT-

AuNPs 40 nm and PEG-AuNPs 40 nm in Volvic water. Scale bar is 2 µm

3.2.4.2 CytoViva® dark field hyperspectral imaging

In order to determine whether Ag and Au signals observed with NanoSIMS are internalized as nanoparticles,

Cytoviva dark field hyperspectral imagining microscope was used, since the hyperspectral imaging only allows

the detection of NPs and not of ions. Analysis were performed in different organs and tissues (gills, gut,

muscle and cuticle). Only results for gills and gut are presented in order to compare them with NanoSIMS

results (Figure 4. 5, Figure 4. 6).

The analysis of Ag in G. fossarum gills revealed the presence of a AgNPs in the gills of animals exposed to CIT-

AgNPs and PEG-AgNPs, highlighted with red spots (Figure 4. 5 C). As already observed with the NanoSIMS

analysis, the amount of AgNPs detected in animals exposed to CIT-coated AgNPs is higher than the amount

detected in animals exposed to PEG-coated AgNPs.

Control

Cit-AuNPs

40 nm

PEG-AuNPs

40 nm

CN-

197Au

197Au

CN-

A B C


139


the area observed with Cytoviva darkfield hyperspectral microscope. Scale bar is 250 µm. B) G. fossarum gills

viewed with Cytoviva (60x oil immersion magnification). C) AgNPs accumulation in G. fossarum gills (red spots

and white arrows). Animals were exposed for 72h to 10 µg. L-1 of CIT-AgNPs 40 nm and PEG-AgNPs 40 nm in Volvic

water. Scale bar is 6 µm

The analysis of the intestinal caeca of G. fossarum exposed to CIT-AuNPs 40 nm allowed the detection of

AuNPs within the epithelial tissue (Figure 4. 6 C). Identical to what was observed with NanoSIMS analysis,

PEG-AuNPs were not detectable in exposed animals.

Cit-AgNPs

40 nm

PEG-AgNPs

40 nm

Control

A B C


140


the area observed with Cytoviva darkfield hyperspectral microscope. Scale bar is 250 µm. B) G. fossarum intestinal

caeca viewed with Cytoviva (60x oil immersion magnification). C) AuNPs accumulation in G. fossarum intestinal

caeca (green spots and white arrows). Animals were exposed for 72h to 10 µg. L-1 of CIT-AgNPs 40nm and PEG-

AgNPs 40 nm in Volvic water. Scale bar is 6 µm

3.2.5 Molecular responses

In order to better understand the effects of sub-lethal concentrations AgNPs and AuNPs in G. fossarum and

elucidation of the acute toxic mechanisms, we assessed the expression of genes involved in oxidative stress,

general stress, apoptosis, DNA damage and repair, cytoskeleton remodelling and trafficking, osmoregulation

and respiration. The analysis of gene expression evidenced specific patterns in relation to the metal of the

particle (Table 4. 4 and Table 4. 5). Indeed, the effects observed on gene expression were dependent on the

chemical composition of the studied NPs (Ag or Au). AuNPs 80 nm led to a down-regulation of both MnSOD

and CuZnSOD in G. fossarum whereas AgNPs 80 nm induced both of them. MnSOD was induced in G.

fossarum exposed to 25 and 50 µg. L-1 of PEG-AgNPs 80 nm (1.5-fold increase for both concentrations)

whereas no significant effect of PEG-AuNPs 80 nm was observed. In contrast, MnSOD and CuZnSOD were

down-regulated (2.5- and 4-fold decrease) in G. fossarum exposed to CIT-AuNPs 80 nm (25 and 50 µg. L-1,

respectively), while no significant effect of CIT-AgNPs 80nm was observed. Chitinase was induced 2- and 1.7-

fold by exposure to 10 and 25 µg. L-1 of CIT-AgNPs 40 nm respectively and down regulated by 1.75-fold at 50

µg. L-1 of CIT-AgNPs 40 nm. A 1.4-, 2.3- and 2.8-fold down regulation of GST of was also observed in G.

A B C

Control

Cit-AuNPs

40 nm

PEG-AuNPs

40 nm


141

fossarum exposed to 10, 25, 50 µg. L-1 of PEG-AgNPs 80 nm, respectively. Gpx7 was impacted only in G.

fossarum exposed to the highest concentrations of PEG-AuNPs 20 nm and CIT-AuNPs 40 nm with a 2-fold

increase. However, the observed responses were not coating or size-dependent.


142

Table 4. 4. Relative gene expression of G. fossarum exposed for 72h to CIT-AgNPs and PEG-AgNPs 20, 40 and 80 nm

indicates significant (P < 0.05) up-regulated genes (fold change > 1.5) indicates significant (P < 0.05) down-regulated genes (fold change < 0.5)

indicates non-significant effect on relative gene expression.

exoskeleton/cuticle Lysososmes

Actin TUB UB Chitinase CAT MnSOD CuZnSOD GPx7 GST HSP90 GAPDH Gadd45 NfkB Cjun P53 CathepsinL NaKATPase Hemocyanin

0.5 + + - -

1 +

2

4 + + + + +

8

1 +

2

4 + +

8 +

10 -

1 +

2 +

4

8 +

10

1 +

5

10 + + + -

25 +

50 -

1 + + -

5 -

10 +

25 + + -

50 -

1 +

5 +

10 - + - -

25

50

1 + +

5

10 + + - -

25 + + - - -

50 + - -

AgNPs40nm

citrate

PEG

AgNPs80nm

citrate

PEG

Compound Concentrations(µg.L-1)

AgNO3

AgNPs20nm

Citrate

PEG

Generalstress DNAdamageandrepair OsmoregulationandRespiration

Cytoskeleton Antioxidantdefence

+ - n.s


143

Table 4. 5. Relative gene expression of G. fossarum exposed for 72h to CIT-AuNPs and PEG-AuNPs 20, 40 and 80 nm

indicates significant (P < 0.05) up-regulated genes (fold change > 1.5) indicates significant (P < 0.05) down-regulated genes (fold change < 0.5)

indicates non-significant effect on relative gene expression

exoskeleton/cuticle Lysososmes

Actin TUB UB Chitinase CAT MnSOD CuZnSOD GPx7 GST HSP90 GAPDH Gadd45 NfkB Cjun P53 CathepsinL NaKATPase Hemocyanin

0.5 - - + -

1 +

2 + +

1 - +

5 +

10

25 - - +

50

1

5

10 -

25 - - +

50 - +

1

5 -

10 - -

25

50 - - + +

1

5 - -

10 -

25 - -

50 - -

1

5 -

10 - -

25 + - -

50 + - - +

1 - -

5 -

10 - -

25 -

50 - -

Compound

PEG

AuNPs80nm

citrate

PEG

AuNPs40nm

citrate

PEG

AuNPs20nm

Citrate

AgNO 3

OsmoregulationandRespirationDNAdamageandrepairGeneralstressAntioxidantdefenceCytoskeletonConcentrations(ug.L

-1)

+ - n.s


144

4 DISCUSSION

The fate and behaviour of nanoparticles including AgNPs and AuNPs in the aquatic environment will

undoubtedly influence their toxicity towards aquatic organisms (Lapresta-Fernández et al., 2012;

McGillicuddy et al., 2017; Vale et al., 2016). In the present study, we investigated the influence of size and

surface coating on the uptake and acute effects of AgNPs and AuNPs on G. fossarum following a realistic

exposure scenario in terms of concentrations, which are expected to be in the low µg.L-1, respectively

(Gottschalk et al., 2013; Yang et al., 2016). Three different sizes (20, 40, 80 nm) and two different coatings

(CIT and PEG) were used both for AgNPs and AuNPs. The studied endpoints included uptake, tissue

distribution and short-term effects of the selected AgNPs and AuNPs on expression of stress-related genes of

G. fossarum.

The studied AgNPs and AuNPs appeared to be stable in the exposure medium (commercial mineral water,

Volvic®, France), with limited tendency to agglomeration and sedimentation in the experimental conditions.

Only CIT-AgNPs 20 nm and CIT-AuNPs 20 nm showed the tendency to aggregation with measured values for

hydrodynamic diameters of 53 and 154 nm and zeta potential of -19.1 and -17.1, respectively. It is usually

described that NPs with a zeta potential within -30mV and +30mV range have a tendency to agglomerate in

the tested experimental conditions (Lee and Ranville, 2012). However, in the present study, it seems that

zeta potential is not a good indicator of stability for CIT-AgNPs and CIT-AuNPs as for sterically repulsed NPs

(Sørensen et al., 2016). These results are consistent with previous studies that reported instability and an

increase in size of CIT-AgNPs (Baalousha et al., 2013; Park et al., 2015a; Sakka et al., 2016) and CIT-AuNPs

(Park et al., 2015b) compared to PEG-NPs (Park et al., 2015b; Tejamaya et al., 2012).

AgNPs used in commercial products are generally coated with organic compounds, such as PEG, in order to

increase their colloidal stability by reducing the surface energy (Ju-Nam and Lead, 2008). As a result, capping

agents prevent interaction of the NPs with the surrounding environment and between them, thus avoiding

or reducing their aggregation (Kvítek et al., 2008), which may explain the increased stability of PEG-AgNPs

and PEG-AuNPs in our exposure medium (Volvic water).

Differences in stability can affect behaviour, bioavailability and toxicity of AgNPs and AuNPs (Kwok et al.,

2012; Levard et al., 2012; Park et al., 2015a). Thus, Ag and Au total and ionic concentrations in exposure

medium were evaluated in order to determine the behaviour and fate of the AgNPs and AuNPs. Total Au

concentration could not be determined and further analyses are therefore needed. Ag concentrations in the

exposure medium were close to the nominal ones in the low µg. L-1 range while a plateau around 20 µg. L-1

was observable at the higher exposure concentrations (25 to 50 µg. L-1). This loss in Ag at the highest

concentrations may be linked to a rapid sedimentation (Bacchetta et al., 2017; Farmen et al., 2012; Scown et


145

al., 2010; Zimmermann et al., 2017) or to the uptake by the animals. Furthermore, dissolution assessment

showed that only 6% of Ag ions are released at the highest AgNPs concentrations (50 µg. L-1) indicating that

the tested particles are stable in our experimental conditions as shown by our NTA results for CIT-AgNPs and

CIT-AuNPs 40 and 80 nm and PEG-AgNPs and PEG-AuNPs. This result is in accordance with previous studies

that showed low dissolution rate for CIT-AgNPs, PEG-AgNPs and PVP-AgNPs (Park et al., 2015a, 2015b).

Hence, some studies suggested that presence of organisms or organic matter decreased, through adsorption,

the amount of ionic Ag released from AgNPs (Bone et al., 2012; Cupi et al., 2015; Newton et al., 2013).

Exposure of G. fossarum to AgNPs and AuNPs for 72h led to a significant coating-dependent bioaccumulation

being of CIT-AgNPs and CIT-AuNPs, which were taken up in higher amounts than PEG-AgNPs and PEG-AuNPs,

which is in accordance with previous studies (Oliver et al., 2014; Park et al., 2015b; Sakka et al., 2016). A

coating dependent bioaccumulation of CIT-AgNPs and CIT-AuNPs was reported in Lymnea stagnalis (Oliver et

al., 2014), D. magna (Sakka et al., 2016) and Gammarus pulex (Park et al., 2015b). The observed difference in

the bioaccumulation rate between AgNPs and AuNPs could be due to the sedimentation of CIT-AgNPs and

CIT-AuNPs linked to the reduced colloidal stability of the particles (Croteau et al., 2011; Sakka et al., 2016) as

compared to PEG-coated NPs, which are more stable and thus have a lower tendency for agglomeration and

sedimentation. Since G. fossarum is a benthic invertebrate living at the water/sediment interface, this could

enhance the bioavailability of the CIT-NPs, which will be in the proximity of the animals (Kunz et al., 2010;

Park et al., 2015b). It has been suggested that PEG prevent the interaction of the NPs with the surrounding

environment leading, for instance, to the formation of a reduced and more labile protein corona than for CIT-

coated NPs, which is supposed to be important for the uptake of NPs by aquatic organisms (Lynch and

Dawson, 2008; Nasser and Lynch, 2016; Sasidharan et al., 2015).

In addition to influences of the surface coating, we also investigated the influence of size on the uptake of

AgNPs and AuNPs. While a decrease of uptake with increasing size of AgNPs and AuNPs could be expected

(Georgantzopoulou, 2015; Georgantzopoulou et al., 2013; Mehennaoui et al., 2016), no clear relationship

between size and bioaccumulation in the organisms was observed in the present work. Previous studies

reported that size did not influence the uptake of AgNPs by G. roeseli (Andreï et al., 2016). Same observations

were reported for the uptake of AuNPs by D. magna (Khan et al., 2014; Wray and Klaine, 2015) and G. pulex

(Park et al., 2015b). In these previous studies, rather than the size, a concentration-dependent uptake was

highlighted and it had been described that the particle number seem to be an important factor that influence

the uptake (Park et al., 2015b) and an increase of the particles number led to an increase of bioaccumulation.

When comparing Ag bioaccumulation after treatment with AgNPs or AgNO3, at similar exposure

concentrations (~5 µg. L-1), AgNO3 led to the higher uptake of Ag by G. fossarum. This result is in accordance

with previous studies in D. magna (Croteau et al., 2011; Zhao and Wang, 2010), snails (Chen et al., 2017; Khan


146

et al., 2015) and rainbow trout (Bruneau et al., 2016). This was explained by the high affinity of Ag+ for Na+

channels in membrane transporters (Bianchini and Wood, 2003). Indeed, in our previous study, a decrease in

haemolymph osmolality of G. fossarum exposed to AgNO3 was observed and this was linked to the decrease

of Na+ concentration in haemolymph (Arce Funck et al., 2013; Mehennaoui et al., 2016; Ribeiro et al., 2017).

AgNO3 also showed to be more toxic to G. fossarum leading to a significant mortality rate (Andreï et al., 2016;

Arce Funck et al., 2013; Mehennaoui et al., 2016). The recorded LC50s during the present study showed a

significant seasonal variability in G. fossarum sensitivity as the LC50 recorded in autumn was lower than the

one recoded in summer. It is well established that the season greatly influence the physiological status and

metabolism of gammarids (Lebrun et al., 2015), which are normally field collected, leading to subsequent

changes in their responses to chemicals (Dedourge-Geffard et al., 2013; Gismondi et al., 2012; Lebrun et al.,

2015; Sroda and Cossu-Leguille, 2011). Despite the significant bioaccumulation of AgNPs and AuNPs used in

the present study, none of them were lethal to G. fossarum, nor did they show any sign of acute toxicity.

These results are also consistent with literature that reported an absence of mortality after exposing G.

roeseli to 0.5 and 5 µg.L-1 CIT-AgNPs 10 and 60 nm (Andreï et al., 2016) and G. pulex exposed to up to 1 mg.L-

1 of CIT-AuNPs and PEG-AuNPs (Park et al., 2015b).

To better understand the mechanisms underlying AgNPs and AuNPs bioaccumulation, it is important to know

how NPs behave upon internalization by the organism. NanoSIMS and Cytoviva are good and powerful

techniques to investigate the distribution of NPs within biological matrices. NanoSIMS allows the elemental

detection, with high sensitivity, of virtually the whole periodic table, but it cannot be used to distinguish ions

from intact particles. On the contrary, Cytoviva allows the detection of nanoparticles, through the

hyperspectral analysis, but it does not allow the detection of ions. The use of the two techniques allowed the

detection of AgNPs in gills of G. fossarum and AuNPs close to the intestine of G. fossarum. The obtained

signals, with both techniques (NanoSIMS and Cytoviva) were higher for CIT-AgNPs and CIT-AuNPs which is

consistent with ICP-MS results. The higher uptake of CIT-AgNPs and CIT-AuNPs could be explained by the

attachments on the cuticle of G. fossarum as reported in other studies that used D. Magna as model species

(Asghari et al., 2012; Baumann et al., 2014; Sakka et al., 2016; Zhao and Wang, 2012). The analysis performed

with the NanoSIMS and Cytoviva supports this hypothesis, as presence of CIT-AgNPs and CIT-AuNPs was

observed on the cuticle of G. fossarum with a higher signal obtained for AgNPs than AuNPs. Additionally, CIT-

AgNPs were observed in the gills, more specifically the cuticle and the epithelial tissue of G. fossarum gills.

This result is in accordance with many other studies that reported the presence of AgNPs in the gills of

mussels (Canesi et al., 2012), zebrafish and perch (Bilberg et al., 2010; Griffitt et al., 2009; Scown et al., 2010)

and around the nucleus of gill cells of rainbow trout (Farkas et al., 2011). Our results appear to be consistent

with previous reports since fish and crustaceans’ gills are in direct contact with the external environment and


147

are described to play a protection role as a barrier. Crustaceans’ gills are also described as the major sites of

accumulation of heavy metals and toxic compounds as they can be trapped in the mucus layer or absorbed

by the epithelial cells (Ali, 2014; Georgantzopoulou et al., 2016; Henry et al., 2012). It is suggested that the

main uptake mechanism for AgNPs is the direct uptake from the water through the gills (Ribeiro et al., 2017).

Other studies suggested endocytosis process (pinocytosis and phagocytosis) to explain AgNPs uptake by D.

magna (Ribeiro et al., 2017).

AuNPs were observed in intestinal caeca of G. fossarum. Previous studies reported the presence of AuNPs in

the gut, carapace, antennae and appendages of D. magna (Botha et al., 2016; Jensen et al., 2017; Lee and

Ranville, 2012). AuNPs have been also detected close to the peritrophic membrane and in the microvilli of D.

magna gut, where NPs inhibited the intestinal clearance and thus the excretion of NPs. Ingestion is very likely

the main uptake route of AuNPs in freshwater crustaceans (Wray and Klaine, 2015). Thus, internalization of

nanoparticles by G. fossarum seems to be metal-dependent and can take multiple routes (Lee and Ranville,

2012; Lovern et al., 2008): AgNPs seem to be taken up through the gills while AuNPs may be ingested through

the mouth. It had been suggested that fish and crustaceans’ gills are the primary organ for AgNPs uptake

although Ag was present in the gut, liver, brain depending on the surface coating (Kwok et al., 2012) indicating

that Ag might be distributed to different organs after being taken up through the gills. The intestinal caeca

participated in the production of digestive fluids (Cristescu and Hebert, 2005) and were reported to be the

sites of Cd and Ca2+ uptake (Georgantzopoulou et al., 2016) and seem to be the main site for bioaccumulation

of AuNPs in G. fossarum.

There are many uncertainties about mechanisms of toxicity of AgNPs and AuNPs on aquatic organisms. A

transcriptomic approach, through targeted gene expression method, was used in order to investigate AgNPs

and AuNPs effects at sub-lethal concentrations. A set of genes was selected based on our previous work on

G. fossarum (Mehennaoui et al., 2016) in which a battery of biomarkers was used to asses AgNPs effects. The

set of genes included genes associated with oxidative stress (CAT, MnSOD, CuZnSOD, GPx7), general stress

and detoxification mechanisms (GST and HSP90), cytoskeleton remodelling and trafficking (Actin, TUB, UB),

genotoxicity (c-jun, Gadd45, NfkB, P53), lysosomes (Cathepsin L), moulting (Chitinase), osmoregulation and

respiration (Na+K+ATPase and HEM) were analysed. AgNPs led to an increase of both SODs expression while

AuNPs led to their down regulation. SODs are known to be antioxidant enzymes converting the superoxide

anion to H2O2 (Sroda and Cossu-Leguille, 2011; Vasseur and Cossu-Leguille, 2004) and oxidative stress is

described as one of the most frequent effects reported in aquatic organisms exposed to NPs (Vale et al.,

2016). The down-regulation of MnSOD in G. fossarum exposed to AuNPs was in accordance with a previous

study that reported a decrease of MnSOD expression in gills of C. fulminate (Renault et al., 2008) exposed to

contaminated algae with up to 1.6 x 105 amine-coated AuNPs/cell and in G. fossarum exposed through


148

contaminated diet to up to 46 mg. L-1 of AuNPs 10 nm (Baudrimont et al., 2017). However, our results are in

contrast with other studies that reported the absence of significant effects of AgNPs on SOD expression in

the gills of M. galloprovincialls exposed for 15 days to 10 µg.L-1 of AgNPs (Bebianno et al., 2015) and in liver

of zebrafish exposed to up to 120 mg.L-1 of AgNPs (Choi et al., 2010). It has been described that gene

expression vary dependently of the studied tissue/organ (Guerriero et al., 2014). Therefore, these results

may be explained by the fact that authors looked to the effects of AgNPs and AuNPs in one specific organ

while in the present work gene expression were investigated in the whole animal. In the present study, the

increase in expression of CuZnSOD could indicate an induction of antioxidant defences. Ag is known to

interact with thiol group found in many antioxidant enzymes and it is also known to generate production of

intra and extracellular reactive oxygen species (ROS) (Bebianno et al., 2015; Connolly et al., 2015;

McGillicuddy et al., 2017). Previous studies suggested that generation of ROS was due to the presence of

AgNPs themselves in addition to Ag ions that could be dissolved within the organism (Ahn et al., 2014;

Bebianno et al., 2015). These compounds were described to lead to an overwhelming of antioxidant defence

and metallothionein, which plays an important detoxification role (Gomes et al., 2013; Gomes et al., 2014b).

Thus, it might be suggested that internalization of AgNPs or their adsorption on the cuticles of the gills in G.

fossarum induced to the production of ROS leading to the activation of antioxidant defences. Moreover, the

down-regulation of Cathepsin L and GST by AgNPs 80 nm could be linked to the interaction of AgNPs with

thiol groups of proteins and a potential disruption of glutathione metabolism leading to oxidative stress

(Gismondi et al., 2012; Vellinger et al., 2013). Additionally, the significant effect on Chitinase expression in G.

fossarum exposed to CIT-AgNPs 40 nm may indicate that CIT-AgNPs 40 nm are interacting/interfering with

the structure and the maintenance of exoskeleton. The presence of CIT-AgNPs 40 nm on the cuticle of the

gills observed with NanoSIMS 50 and Cytoviva could therefore explain this result. This effect may disturb ion

transport, gaseous exchange, growth process of G. fossarum that occur through successive moults (Sutcliffe,

1992) and lead to deleterious effects at the population level.

5 CONCLUSION

In the present study, we observed a coating-dependent effect that enhance the uptake of CIT-coated particles

as compared to PEG-coated particles. This uptake led to a metal-dependent distribution of AgNPs and AuNPs

within G. fossarum tissues. AgNPs were mainly found in the gills, while AuNPs were mainly observed in

intestinal caeca. AgNPs were found to up-regulate MnSOD and CuZnSOD gene expression while AuNPs led to

their down-regulation. The observed effects seemed to be directly linked to the NPs themselves as no ions

were detected with ICP-MS and Cytoviva allowed to confirm a significant presence of NPs within the tissues.

Modulation by AgNPs of SODs may indicate a generation of ROS and possible activation of SOD enzymes in


149

order to defend and protect the organism from a potential oxidative stress. However, to affirm this

hypothesis, further investigations are needed in order to better define mechanisms of action at the molecular

level of AgNPs and AuNPs on G. fossarum.

Acknowledgments

This work was supported by National Research Fund, Luxembourg (AFR-PhD-9229040). The authors are

thankful to S. Contal and M. Fossépré for technical support, C. Guignard for chemical analyses, E. Lentzen and

A. Chauvière for NanoSIMS analyses.

REFERENCES

Ahn, J.-M., Eom, H.-J., Yang, X., Meyer, J.N., Choi, J., 2014. Comparative toxicity of silver nanoparticles on oxidative

stress and DNA damage in the nematode, Caenorhabditis elegans. Chemosphere 108, 343–352.


Ali, D., 2014. Oxidative Stress-Mediated Apoptosis and Genotoxicity Induced by Silver Nanoparticles in Freshwater

Snail Lymnea luteola L. Biol. Trace Elem. Res. 162, 333–341. doi:10.1007/s12011-014-0158-6




Angel, B.M., Batley, G.E., Jarolimek, C.V., Rogers, N.J., 2013. The impact of size on the fate and toxicity of

nanoparticulate silver in aquatic systems. Chemosphere. doi:10.1016/j.chemosphere.2013.04.096



143, 73–84. doi:10.1016/j.aquatox.2013.07.012

Asghari, S., Johari, S.A., Lee, J.H., Kim, Y.S., Jeon, Y.B., Choi, H.J., Moon, M.C., Yu, I.J., 2012. Toxicity of various silver

nanoparticles compared to silver ions in Daphnia magna. J. Nanobiotechnology 10, 1–11. doi:10.1186/1477-3155-

10-14

Baalousha, M., Nur, Y., Römer, I., Tejamaya, M., Lead, J.R., 2013. Effect of monovalent and divalent cations, anions

and fulvic acid on aggregation of citrate-coated silver nanoparticles. Sci. Total Environ. 454–455, 119–131.

doi:10.1016/j.scitotenv.2013.02.093







doi:10.1007/s11356-017-8400-3

Baumann, J., Köser, J., Arndt, D., Filser, J., 2014. The coating makes the difference: Acute effects of iron oxide

nanoparticles on Daphnia magna. Sci. Total Environ. 484, 176–184. doi:10.1016/j.scitotenv.2014.03.023

Bebianno, M.J., Gonzalez-Rey, M., Gomes, T., Mattos, J.J., Flores-Nunes, F., Bainy, A.C.D., 2015. Is gene


150

transcription in mussel gills altered after exposure to Ag nanoparticles? Environ. Sci. Pollut. Res. 1–9.

doi:10.1007/s11356-015-5186-z

Behr, M., Legay, S., Hausman, J.-F., Guerriero, G., 2015. Analysis of Cell Wall-Related Genes in Organs of Medicago

sativa L. under Different Abiotic Stresses. Int. J. Mol. Sci. 16, 16104–16124. doi:10.3390/ijms160716104

Bianchini, A., Wood, C.M., 2003. Mechanism of acute silver toxicity in Daphnia magna. Environ. Toxicol. Chem. 22,

1361–1367. doi:10.1002/etc.5620220624

Bilberg, K., Malte, H., Wang, T., Baatrup, E., 2010. Silver nanoparticles and silver nitrate cause respiratory stress in

Eurasian perch (Perca fluviatilis). Aquat. Toxicol. 96, 159–165. doi:10.1016/j.aquatox.2009.10.019

Bone, A.J., Colman, B.P., Gondikas, A.P., Newton, K.M., Harrold, K.H., Cory, R.M., Unrine, J.M., Klaine, S.J., Matson,

C.W., Di Giulio, R.T., 2012. Biotic and Abiotic Interactions in Aquatic Microcosms Determine Fate and Toxicity of

Ag Nanoparticles: Part 2–Toxicity and Ag Speciation. Environ. Sci. Technol. 46, 6925–6933.

doi:10.1021/es204683m





Cambier, S., Røgeberg, M., Georgantzopoulou, A., Serchi, T., Karlsson, C., Verhaegen, S., Iversen, T.-G., Guignard,

C., Kruszewski, M., Hoffmann, L., Audinot, J.-N., Ropstad, E., Gutleb, A.C., 2018. Fate and effects of silver

nanoparticles on early life-stage development of zebrafish (Danio rerio) in comparison to silver nitrate. Sci. Total

Environ. 610, 972–982. doi:10.1016/j.scitotenv.2017.08.115

Canesi, L., Ciacci, C., Fabbri, R., Marcomini, A., Pojana, G., Gallo, G., 2012. Bivalve molluscs as a unique target group

for nanoparticle toxicity. Mar. Environ. Res., Emerging and persistent impacts on Marine Organisms: Detection

methods and action mechanisms 76, 16–21. doi:10.1016/j.marenvres.2011.06.005

Chen, Y., Si, Y., Zhou, D., Dang, F., 2017. Differential bioaccumulation patterns of nanosized and dissolved silver in

a land snail Achatina fulica. Environ. Pollut. 222, 50–57. doi:10.1016/j.envpol.2017.01.007



doi:10.1016/j.aquatox.2009.12.012

Cleveland, D., Long, S.E., Pennington, P.L., Cooper, E., Fulton, M.H., Scott, G.I., Brewer, T., Davis, J., Petersen, E.J.,

Wood, L., 2012. Pilot estuarine mesocosm study on the environmental fate of Silver nanomaterials leached from

consumer products. Sci. Total Environ., Special Section: Reviews of Trace Metal Pollution in China 421–422, 267–

272. doi:10.1016/j.scitotenv.2012.01.025

Connolly, M., Fernandez-Cruz, M.-L., Quesada-Garcia, A., Alte, L., Segner, H., Navas, J.M., 2015. Comparative

Cytotoxicity Study of Silver Nanoparticles (AgNPs) in a Variety of Rainbow Trout Cell Lines (RTL-W1, RTH-149, RTG-

2) and Primary Hepatocytes. Int. J. Environ. Res. Public. Health 12, 5386–5405. doi:10.3390/ijerph120505386

Cristescu, M.E., Hebert, P.D., 2005. The “Crustacean Seas” — an evolutionary perspective on the Ponto–Caspian

peracarids. Can. J. Fish. Aquat. Sci. 62, 505–517. doi:10.1139/f04-210

Croteau, M.-N., Misra, S.K., Luoma, S.N., Valsami-Jones, E., 2011. Silver Bioaccumulation Dynamics in a Freshwater

Invertebrate after Aqueous and Dietary Exposures to Nanosized and Ionic Ag. Environ. Sci. Technol. 45, 6600–

6607. doi:10.1021/es200880c


151

Cumberland, S.A., Lead, J.R., 2009. Particle size distributions of silver nanoparticles at environmentally relevant

conditions. J. Chromatogr. A 1216, 9099–9105. doi:10.1016/j.chroma.2009.07.021

Cupi, D., Hartmann, N.B., Baun, A., 2015. The influence of natural organic matter and aging on suspension stability

in guideline toxicity testing of silver, zinc oxide, and titanium dioxide nanoparticles with Daphnia magna. Environ.


Dedourge-Geffard, O., Charron, L., Hofbauer, C., Gaillet, V., Palais, F., Lacaze, E., Geffard, A., Geffard, O., 2013.

Temporal patterns of digestive enzyme activities and feeding rate in gammarids (Gammarus fossarum) exposed to

inland polluted waters. Ecotoxicol. Environ. Saf. 97, 139–146. doi:10.1016/j.ecoenv.2013.07.016






Biol. 58, 419–472. doi:10.1163/157075608X383719

Ellis, L.-J.A., Valsami-Jones, E., Lead, J.R., Baalousha, M., 2016. Impact of surface coating and environmental

conditions on the fate and transport of silver nanoparticles in the aquatic environment. Sci. Total Environ. 568,

95–106. doi:10.1016/j.scitotenv.2016.05.199

Eybe, T., Bohn, T., Audinot, J.N., Udelhoven, T., Cauchie, H.M., Migeon, H.N., Hoffmann, L., 2009. Uptake

visualization of deltamethrin by NanoSIMS and acute toxicity to the water flea Daphnia magna. Chemosphere 76,

134–140. doi:10.1016/j.chemosphere.2009.02.005



Farkas, J., Christian, P., Gallego-Urrea, J.A., Roos, N., Hassellöv, M., Tollefsen, K.E., Thomas, K.V., 2011. Uptake and

effects of manufactured silver nanoparticles in rainbow trout (Oncorhynchus mykiss) gill cells. Aquat. Toxicol. 101,

117–125. doi:10.1016/j.aquatox.2010.09.010

Farmen, E., Mikkelsen, H.N., Evensen, Ø., Einset, J., Heier, L.S., Rosseland, B.O., Salbu, B., Tollefsen, K.E., Oughton,

D.H., 2012. Acute and sub-lethal effects in juvenile Atlantic salmon exposed to low μg/L concentrations of Ag

nanoparticles. Aquat. Toxicol., Proceedings from the 16th International Symposium on Pollutant Responses in

Marine Organisms (PRIMO16) 108, 78–84. doi:10.1016/j.aquatox.2011.07.007



doi:10.1002/etc.5620190820

Gao, J., Powers, K., Wang, Y., Zhou, H., Roberts, S.M., Moudgil, B.M., Koopman, B., Barber, D.S., 2012. Influence of

Suwannee River humic acid on particle properties and toxicity of silver nanoparticles. Chemosphere 89, 96–101.


García-Negrete, C.A., Blasco, J., Volland, M., Rojas, T.C., Hampel, M., Lapresta-Fernández, A., Jiménez de Haro,

M.C., Soto, M., Fernández, A., 2013. Behaviour of Au-citrate nanoparticles in seawater and accumulation in

bivalves at environmentally relevant concentrations. Environ. Pollut. 174, 134–141.

doi:10.1016/j.envpol.2012.11.014

Georgantzopoulou, A., 2015. Effects of silver nanoparticles and ions and interaction with first line of defense.


152

Wageningen University, Wageningen Netherlands.




doi:10.3109/17435390.2012.715312

Georgantzopoulou, A., Cambier, S., Serchi, T., Kruszewski, M., Balachandran, Y.L., Grysan, P., Audinot, J.-N., Ziebel,

J., Guignard, C., Gutleb, A.C., Murk, A.J., 2016. Inhibition of multixenobiotic resistance transporters (MXR) by silver

nanoparticles and ions in vitro and in Daphnia magna. Sci. Total Environ. 569–570, 681–689.

doi:10.1016/j.scitotenv.2016.06.157

Gismondi, E., Beisel, J.-N., Cossu-Leguille, C., 2012. Influence of gender and season on reduced glutathione

concentration and energy reserves of Gammarus roeseli. Environ. Res. 118, 47–52.

doi:10.1016/j.envres.2012.06.004

Gomes, S.I.L., Soares, A.M.V.M., Scott-Fordsmand, J.J., Amorim, M.J.B., 2013. Mechanisms of response to silver

nanoparticles on Enchytraeus albidus (Oligochaeta): Survival, reproduction and gene expression profile. J. Hazard.

Mater. 254–255, 336–344. doi:10.1016/j.jhazmat.2013.04.005

Gomes, T., Pereira, C.G., Cardoso, C., Sousa, V.S., Teixeira, M.R., Pinheiro, J.P., Bebianno, M.J., 2014. Effects of

silver nanoparticles exposure in the mussel Mytilus galloprovincialis. Mar. Environ. Res.

doi:10.1016/j.marenvres.2014.07.004





Guerriero, G., Legay, S., Hausman, J.-F., 2014. Alfalfa Cellulose Synthase Gene Expression under Abiotic Stress: A

Hitchhiker’s Guide to RT-qPCR Normalization. PLOS ONE 9, e103808. doi:10.1371/journal.pone.0103808



doi:10.3389/fphys.2012.00431

Hole, P., Sillence, K., Hannell, C., Maguire, C.M., Roesslein, M., Suarez, G., Capracotta, S., Magdolenova, Z., Horev-

Azaria, L., Dybowska, A., Cooke, L., Haase, A., Contal, S., Manø, S., Vennemann, A., Sauvain, J.-J., Staunton, K.C.,

Anguissola, S., Luch, A., Dusinska, M., Korenstein, R., Gutleb, A.C., Wiemann, M., Prina-Mello, A., Riediker, M.,

Wick, P., 2013. Interlaboratory comparison of size measurements on nanoparticles using nanoparticle tracking

analysis (NTA). J. Nanoparticle Res. 15, 1–12. doi:10.1007/s11051-013-2101-8

Isnard, P., Flammarion, P., Roman, G., Babut, M., Bastien, P., Bintein, S., Esserméant, L., Férard, J.F., Gallotti-

Schmitt, S., Saouter, E., Saroli, M., Thiébaud, H., Tomassone, R., Vindimian, E., 2001. Statistical analysis of

regulatory ecotoxicity tests. Chemosphere 45, 659–669. doi:10.1016/S0045-6535(00)00600-7



Jensen, L.H.S., Skjolding, L.M., Thit, A., Sørensen, S.N., Købler, C., Mølhave, K., Baun, A., 2017. Not all that glitters

is gold—Electron microscopy study on uptake of gold nanoparticles in Daphnia magna and related artifacts.

Environ. Toxicol. Chem. 36, 1503–1509. doi:10.1002/etc.3697


153

Ju-Nam, Y., Lead, J.R., 2008. Manufactured nanoparticles: An overview of their chemistry, interactions and

potential environmental implications. Sci. Total Environ. 400, 396–414. doi:10.1016/j.scitotenv.2008.06.042



Khan, F.R., Misra, S.K., Bury, N.R., Smith, B.D., Rainbow, P.S., Luoma, S.N., Valsami-Jones, E., 2015. Inhibition of

potential uptake pathways for silver nanoparticles in the estuarine snail Peringia ulvae. Nanotoxicology 9, 493–

501. doi:10.3109/17435390.2014.948519



Chem. 27, 1825–1851. doi:10.1897/08-090.1




Kvítek, L., Panáček, A., Soukupová, J., Kolář, M., Večeřová, R., Prucek, R., Holecová, M., Zbořil, R., 2008. Effect of

Surfactants and Polymers on Stability and Antibacterial Activity of Silver Nanoparticles (NPs). J. Phys. Chem. C 112,

5825–5834. doi:10.1021/jp711616v

Kwok, K.W.H., Auffan, M., Badireddy, A.R., Nelson, C.M., Wiesner, M.R., Chilkoti, A., Liu, J., Marinakos, S.M.,

Hinton, D.E., 2012. Uptake of silver nanoparticles and toxicity to early life stages of Japanese medaka (Oryzias

latipes): Effect of coating materials. Aquat. Toxicol. 120–121, 59–66. doi:10.1016/j.aquatox.2012.04.012



doi:10.1016/j.mrgentox.2010.04.025

Ladewig, V., Jungmann, D., Köhler, H.-R., Schirling, M., Triebskorn, R., Nagel, R., 2006. Population Structure and

Dynamics of Gammarus fossarum (Amphipoda) Upstream and Downstream from Effluents of Sewage Treatment

Plants. Arch. Environ. Contam. Toxicol. 50, 370–383. doi:10.1007/s00244-005-7039-0

Lapresta-Fernández, A., Fernández, A., Blasco, J., 2012. Nanoecotoxicity effects of engineered silver and gold

nanoparticles in aquatic organisms. TrAC Trends Anal. Chem. 32, 40–59. doi:10.1016/j.trac.2011.09.007

Lebrun, J.D., Geffard, O., Urien, N., François, A., Uher, E., Fechner, L.C., 2015. Seasonal variability and inter-species

comparison of metal bioaccumulation in caged gammarids under urban diffuse contamination gradient:

Implications for biomonitoring investigations. Sci. Total Environ. 511, 501–508.

doi:10.1016/j.scitotenv.2014.12.078

Lee, B.-T., Ranville, J.F., 2012. The effect of hardness on the stability of citrate-stabilized gold nanoparticles and

their uptake by Daphnia magna. J. Hazard. Mater. 213–214, 434–439. doi:10.1016/j.jhazmat.2012.02.025

Legay, S., Guerriero, G., Deleruelle, A., Lateur, M., Evers, D., André, C.M., Hausman, J.-F., 2015. Apple russeting as

seen through the RNA-seq lens: strong alterations in the exocarp cell wall. Plant Mol. Biol. 88, 21–40.

doi:10.1007/s11103-015-0303-4






154

Lynch, I., Dawson, K.A., 2008. Protein-nanoparticle interactions. Nano Today 3, 40–47. doi:10.1016/S1748-

0132(08)70014-8



72, 349–364. doi:10.1111/j.1469-185X.1997.tb00017.x

Makama, S., Piella, J., Undas, A., Dimmers, W.J., Peters, R., Puntes, V.F., van den Brink, N.W., 2016. Properties of

silver nanoparticles influencing their uptake in and toxicity to the earthworm Lumbricus rubellus following

exposure in soil. Environ. Pollut. doi:10.1016/j.envpol.2016.08.016




Mehennaoui, K., Georgantzopoulou, A., Felten, V., Andreï, J., Garaud, M., Cambier, S., Serchi, T., Pain-Devin, S.,

Guérold, F., Audinot, J.-N., Giambérini, L., Gutleb, A.C., 2016. Gammarus fossarum (Crustacea, Amphipoda) as a

model organism to study the effects of silver nanoparticles. Sci. Total Environ. doi:10.1016/j.scitotenv.2016.06.068

Moreno-Garrido, I., Pérez, S., Blasco, J., 2015. Toxicity of silver and gold nanoparticles on marine microalgae. Mar.

Environ. Res., Particles in the Oceans: Implication for a safe marine environment 111, 60–73.

doi:10.1016/j.marenvres.2015.05.008



Newton, K.M., Puppala, H.L., Kitchens, C.L., Colvin, V.L., Klaine, S.J., 2013. Silver nanoparticle toxicity to Daphnia

magna is a function of dissolved silver concentration. Environ. Toxicol. Chem. 32, 2356–2364.

doi:10.1002/etc.2300

Oliver, A.L.-S., Croteau, M.-N., Stoiber, T.L., Tejamaya, M., Römer, I., Lead, J.R., Luoma, S.N., 2014. Does water

chemistry affect the dietary uptake and toxicity of silver nanoparticles by the freshwater snail Lymnaea stagnalis?

Environ. Pollut. 189, 87–91. doi:10.1016/j.envpol.2014.02.010

Park, Chung, J., Colman, B.P., Matson, C.W., Kim, Y., Lee, B.-C., Kim, P.-J., Choi, K., Choi, J., 2015a. Ecotoxicity of

bare and coated silver nanoparticles in the aquatic midge, Chironomus riparius. Environ. Toxicol. Chem. 34, 2023–

2032. doi:10.1002/etc.3019

Park, Woodhall, J., Ma, G., Veinot, J.G.., Boxall, A.B.A., 2015b. Do particle size and surface functionality affect

uptake and depuration of gold nanoparticles by aquatic invertebrates? Environ. Toxicol. Chem. 34, 850–859.

doi:10.1002/etc.2868




Ribeiro, F., Van Gestel, C.A.M., Pavlaki, M.D., Azevedo, S., Soares, A.M.V.M., Loureiro, S., 2017. Bioaccumulation

of silver in Daphnia magna: Waterborne and dietary exposure to nanoparticles and dissolved silver. Sci. Total


Römer, I., White, T.A., Baalousha, M., Chipman, K., Viant, M.R., Lead, J.R., 2011. Aggregation and dispersion of

silver nanoparticles in exposure media for aquatic toxicity tests. J. Chromatogr. A 1218, 4226–4233.

doi:10.1016/j.chroma.2011.03.034


155

Sakka, Y., Skjolding, L.M., Mackevica, A., Filser, J., Baun, A., 2016. Behavior and chronic toxicity of two differently

stabilized silver nanoparticles to Daphnia magna. Aquat. Toxicol. 177, 526–535.

doi:10.1016/j.aquatox.2016.06.025

Sasidharan, A., Riviere, J.E., Monteiro-Riviere, N.A., 2015. Gold and silver nanoparticle interactions with human

proteins: impact and implications in biocorona formation. J. Mater. Chem. B 3, 2075–2082.

doi:10.1039/C4TB01926A




Sørensen, S.N., Holten Lützhøft, H.-C., Rasmussen, R., Baun, A., 2016. Acute and chronic effects from pulse

exposure of D. magna to silver and copper oxide nanoparticles. Aquat. Toxicol. 180, 209–217.

doi:10.1016/j.aquatox.2016.10.004



158, 1288–1295. doi:10.1016/j.envpol.2010.01.022




128.

Tejamaya, M., Römer, I., Merrifield, R.C., Lead, J.R., 2012. Stability of Citrate, PVP, and PEG Coated Silver

Nanoparticles in Ecotoxicology Media. Environ. Sci. Technol. 46, 7011–7017. doi:10.1021/es2038596

Tolaymat, T.M., El Badawy, A.M., Genaidy, A., Scheckel, K.G., Luxton, T.P., Suidan, M., 2010. An evidence-based

environmental perspective of manufactured silver nanoparticle in syntheses and applications: A systematic review

and critical appraisal of peer-reviewed scientific papers. Sci. Total Environ. 408, 999–1006.

doi:10.1016/j.scitotenv.2009.11.003

Vale, G., Mehennaoui, K., Cambier, S., Libralato, G., Jomini, S., Domingos, R.F., 2016. Manufactured nanoparticles

in the aquatic environment-biochemical responses on freshwater organisms: A critical overview. Aquat. Toxicol.

170, 162–174. doi:10.1016/j.aquatox.2015.11.019



Vellinger, C., Felten, V., Sornom, P., Rousselle, P., Beisel, J.-N., Usseglio-Polatera, P., 2012a. Behavioural and






doi:10.1016/j.aquatox.2013.05.010

Vellinger, C., Parant, M., Rousselle, P., Immel, F., Wagner, P., Usseglio-Polatera, P., 2012b. Comparison of arsenate

and cadmium toxicity in a freshwater amphipod (Gammarus pulex). Environ. Pollut. 160, 66–73.

doi:10.1016/j.envpol.2011.09.002


156

Vindimian, É., Garric, J., Flammarion, P., Thybaud, É., Babut, M., 1999. An index of effluent aquatic toxicity designed

by partial least squares regression, using acute and chronic tests and expert judgements. Environ. Toxicol. Chem.

18, 2386–2391. doi:10.1002/etc.5620181037

Vindimian, E., Robaut, C., Fillion, G., 1983. A method for cooperative or noncooperative binding studies using

nonlinear regression analysis on a microcomputer. J. Appl. Biochem. 5, 261–268.





doi:10.1016/j.scitotenv.2015.12.150



Zhao, C.-M., Wang, W.-X., 2012. Importance of surface coatings and soluble silver in silver nanoparticles toxicity

to Daphnia magna. Nanotoxicology 6, 361–370. doi:10.3109/17435390.2011.579632

Zhao, C.-M., Wang, W.-X., 2010. Biokinetic Uptake and Efflux of Silver Nanoparticles in Daphnia magna. Environ.

Sci. Technol. 44, 7699–7704. doi:10.1021/es101484s

Zimmermann, S., Ruchter, N., Loza, K., Epple, M., Sures, B., 2017. Nanoparticulate versus ionic silver: Behavior in

the tank water, bioaccumulation, elimination and subcellular distribution in the freshwater mussel Dreissena

polymorpha. Environ. Pollut. 222, 251–260. doi:10.1016/j.envpol.2016.12.048


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COMPLEMENT 3

Submitted to Science of the Total Environment

Influence of size and surface coating on uptake of silver and gold nanoparticles and the molecular effects

on Gammarus fossarum (Crustacea Amphipoda)

Kahina Mehennaoui a,b, Sébastien Cambier a, Tommaso Serchi a, François Guérold b, Johanna Ziebel a, Jean-

Sebastien Thomann c, Nathalie Valle c, Laure Giambérini b, *, Arno C. Gutleb a*



B Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360, Université de


C Material Research and Technology (MRT) Department, Luxembourg Institute of Science and Technology, 41

rue du Brill, Belvaux, Luxembourg


158

Supplementary material

1- Coating of AgNPs and AuNPs by wet chemistry

Commercial citrate stabilized AgNPs and AuNPs (purchased from Nanocomposix) were coated with

polyethylene glycol (PEG layer MW 3000) via wet chemistry method that allowed the control of the correct

functionalization of isolated NPs. This method relied on the established thiol/metal chemistry.

The amount of added PEG was adjusted to ensure the surface saturation of the NPs (Figure S1 A). The

obtained PEG-AgNPs and AuNPs were characterized for size distribution using Nanoparticles Tracking analysis

(NTA, NanoSIGHT 500, Malvern, UK, Figure S1 B). PEG-AgNPs and PEG-AuNPs were stored at 4 °C until use.

Figure S 2 PEG coating on metal Nanoparticles (here silver NPs). SEM shows clearly the conformal silver coating

with the PEG layer. NTA analysis shows a small size increasing for the 40 and 80 nm silver nanoparticles as a

consequence of their coating with the PEG layer. The sensitivity of NTA was not enough to resolve the size increase

for 20 nm nanoparticles.


159

2- Particle character ization

Figure S 3. Size distribution of CIT-AgNPs 20 nm (A), CIT-AgNPs 40 nm (B) and CIT-AgNPs 80 nm (C) in Volvic

Water (T0h) and CIT-AgNPs 20 nm (D), CIT-AgNPs 40 nm (E) and CIT-AgNPs 80 nm (F) in Volvic water after 24h

of incubation. Size distribution is expressed as particle concentration E6.mL-1. The red errors bars indicate ±SD of

the mean of triplicate measurements

A

B

C

D

E

F


160

Figure S 4. Size distribution of PEG-AgNPs 20 nm (A), PEG-AgNPs 40 nm (B) and PEG-AgNPs 80 nm (C) in Volvic

Water (T0h) and PEG-AgNPs 20 nm (D), PEG-AgNPs 40 nm(E) and PEG-AgNPs 80 nm (F) in Volvic water after 24h



A

B

C

D

E

F


161

Figure S 5. Size distribution of CIT-AuNPs 20 nm (A), CIT-AuNPs 40 nm (B) and CIT-AuNPs 80 nm (C) in Volvic

Water (T0h) and CIT-AuNPs 20 nm (D), CIT-AuNPs 40 nm (E) and CIT-AuNPs 80 nm (F) in Volvic water after 24h



A

B

C

D

E

F


162

Figure S 6. Size distribution of PEG-AuNPs 20 nm (A), PEG-AuNPs 40 nm (B) and PEG-AuNPs 80 nm (C) in Volvic

Water (T0h) and PEG-AuNPs 20 nm (D), PEG-AuNPs 40 nm(E) and PEG-AuNPs 80 nm (F) in Volvic water after 24h



A

B

C

D

E

F


163

3- Preliminary test: Daphnia magna Straus mobility inhibition test ( ISO

6341:1996)

In order to determine the concentrations range to be tested with G. fossarum, an acute Daphnia magna test

was performed. Daphnia magna acute test is the most widely used bioassay in ecotoxicology. It aims at

determining the effective concentration of a pollutant that inhibits the mobility of 50% of Daphnia magna

population (EC50).

3.1- Test Organisms

Experiments were conducted with a strain of Daphnia magna Straus 1820, which was maintained in the

laboratory for four years. Parthenogenetic cultures were carried out in 1L aquaria at 20 °C with LCV medium:

a mixture of (20/80) of Lefevre-Czarda (LC) medium and mineral water (Volvic, France). This mixture is

supplemented with:

• Ca and Mg in order to obtain a total hardness of 250 mg.L-1 and a Ca/Mg ration of 4/1

• A mixture of vitamins containing thiamine HCL, vitamin B12 and biotin.

Cultures were maintained under a 16h light 8h dark and at density of 40 animals per liter. The medium was

renewed three times a week and Daphnids were fed with a mixture of two algal species (Pseudokirchneriella

subcapitata and Scenedesmus subspicatus).

3.2- Biological assay

All experiments were performed according to the guideline ISO 6341 – 1996.

3.2.1- Reference test

Daphnia magna were exposed during 24-48h to an increasing range of concentration of potassium

dichromate (K2Cr2O7) (table 1). The aim is to provide data about the sensitivity and the validity of the acute

test.

Four replicates of five neonates (aged < 24h) were placed in glass test tubes containing 10mL of each test

concentration and control. The neonates were not fed during the test. The assessment endpoint examined

was immobilization. More precisely, juveniles that were able to swim were considered mobile and those

which still moved their antennae but did not swim within 15s after a gentle shaking were considered

immobile.

Table S 2 D. magna reference test

Concentration (mg/L) 0 0.58 0.67 0.78 0.9 1.04 1.2 1.39

K2Cr2O7 (µL) 0 290 335 390 450 520 600 695 ISO medium (mL) 10 9.71 9.66 9.61 9.55 9.48 9.5 9.3 Daphnia (n) 5 5 5 5 5 5 5 5


164

3.2.2- Final test

Acute toxicity of AgNPs and AuNPs was determined during 24 and 48h of exposure (serially diluted in ISO

medium, AgNPs 20nm: 1 – 10 µg. L-1, AgNPs 40 and 80nm: 1 – 50 µg.L-1, AuNPs: 10 – 100 µg.L-1). All tests were

conducted at 20 °C in full darkness. Four replicates of five neonates (aged < 24h) were placed in glass test

tubes containing 10mL of each test concentration and control. The neonates were not fed during the test.

The assessment endpoint examined was immobilization.

3.2.3- Results

The inhibition of mobility rate was determined relatively to the total number of Daphnia used. EC50 of each

NPs and their confidence intervals were calculated using Excel Macro REGtox®.

In all tests conditions, immobilisation in controls was lower than 10% and the EC50-48h after exposure to

K2Cr2O7 was 0.9 mg. L-1 which was in accordance with the validity criteria.

The EC50-48h values obtained, after exposure to AgNPs and AuNPs, are expressed in table 6. A size depend

effect was observed in Daphnids exposed to AgNPs (Table 6). AgNPs 20nm was the most toxic one followed

by AgNPs 40 nm. No full dose-responses curves could be obtained for AgNPs 80 nm and AuNPs. These results

were in accordance with previous studies performed in the Lab with the same particles and the same strain

of Daphnia magna.

Table S 3. EC50 values obtained after 48h exposure of D. magna to AgNPs and AuNPs

NPs AgNPs AuNPs

Size EC50 (µg. L-1) EC50 (µg. L-1)

20 nm 6.7 (5.1 – 8.5) > 100

40 nm 30.8 (25.9 – 36.07) > 100 80 nm > 50 > 100


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4- Acute toxicity of AgNO3 on Gammarus fossarum

Figure S 7. Effects AgNO3 on survival of Gammarus fossarum collected in A) June, B) September and C) November

after 72h of exposure.

0

20

40

60

80

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120

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Survivalrates(%)

Concentration (µg.L-1)

Effetcalculé Effetmoyen Réplicats

CE5 CE10 CE15

CE20 CE25

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40

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0 1 2 3 4 5 6

Survivalrates(%)



CE5 CE10 CE15

CE20 CE25

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C

Chapter 5 AgNPs and AuNPs chronic toxicity

167

Chapter 5

Chronic toxicity of AgNPs and AuNPs

CHAPTER 5

AgNPs and AuNPs chronic toxicity


168

KEY FINDINGS

Aim of the study

The aim of the study was to investigate the sub-chronic toxicity of CIT and PEG-AgNPs 40 nm and CIT and

PEG-AuNPs 40 nm on G. fossarum. A multibiomarker approach including molecular effects (stress-related

gene expression), physiological (osmoregulation) and behavioural (locomotion and ventilation) responses

was used. The biological endpoints also included mortality, bioaccumulation and localization of AgNPs and

AuNPs in G. fossarum tissues.

Experimental design

Key findings

• Concentration-dependent uptake

• Coating-dependent uptake o CIT-AgNPs ≥ PEG-AgNPs

o CIT-AuNPs ≥ PEG-AuNPs

• Influence of chemical composition on the internal distribution o CIT-AgNPs40 nm were found only in G. fossarum gills

o CIT-AuNPs 40 nm were found only in the lumen of intestinal caeca

• A metal-dependent effect on gene expression was also observed with o AgNPs leading to an increase in Na+K+ATPase, CAT and Chitinase gene expression

o AuNPs leading to an increase in CuZnSOD gene expression

o Decrease in locomotion

o No effects on osmoregulation, ventilation and survival

Gammarus fossarum

Organisms Materials

Ag40nm

Au40nm

CIT and PEG-AgNPs 40 nm

CIT and PEG-AuNPs 40 nm

Luxembourg

Schwaarzbaach

Sampling site

Acclimation

D-10

Exposure

D-7 D-1

Temperature : 12 oC



Stalling

D15D-30 D-20

Leave

conditioning

Temperature : 12 oC


Exposure medium renewal: every 3 days Addition of food: every 2 days

D0


169

Sub-chronic effects of AgNPs and AuNPs on Gammarus fossarum (Crustacea Amphipoda): from

molecular to behavioural responses

Kahina Mehennaoui a,b, Sébastien Cambier a, Tommaso Serchi a, François Guérold b, Arno C. Gutleb a*, Laure

Giambérini b, *



b Laboratoire Interdisciplinaire des Environnements Continentaux (LIEC), CNRS UMR 7360, Université de


* Corresponding authors:

Arno C. Gutleb

Postal Address: Environmental Research and Innovation Department

Luxembourg Institute of Science and Technology

5, avenue des Hauts-Fourneaux, L-4362 Esch-sur-Alzette, Luxembourg

E-mail: [email protected]

Laure Giamberini

Postal Address: Laboratoire Interdisciplinaire des Environnements Continentaux, Universite de Lorraine,

Campus Bridoux 57000 Metz, France.

Email : [email protected]


170

ABSTRACT

The aim of the present study was the assessment of the sub-chronic effects of silver nanoparticles (AgNPs)

and gold nanoparticles (AuNPs) of 40 nm primary size either stabilized with citrate (CIT-AgNPs and CIT-AuNPs)

or coated with polyethylene glycol (PEG-AgNPs and PEG-AuNPs). AgNO3 was used as a positive control in

order to study the contribution of ions potentially released from the AgNPs on the observed effects. A

multibiomarker approach was used to study the long-term effects of AgNPs and AuNPs 40 nm on molecular,

physiological and behavioural responses of the freshwater invertebrate Gammarus fossarum.

G. fossarum were exposed for 15 days to 0.5 and 5 µg. L-1 of CIT and PEG AgNPs and AuNPs 40 nm in the

presence of food. A significant uptake of Ag and Au was observed in exposed animals, with CIT-AgNPs

detected in gills and CIT-AuNPs 40 nm within the lumen of intestinal caeca of G. fossarum using hyperspectral

darkfield microscopy (Cytoviva). PEG-AgNPs 40 nm led to an up-regulation of CAT and Chitinase and CIT-

AgNPs and PEG-AgNPs 40 nm led to an up-regulation of Na+K+ATPase. CIT and PEG-AuNPs 40 nm led to an

increase of CuZnSOD gene expression. Both AgNPs and AuNPs 40 nm significantly impacted locomotor activity

of G. fossarum while no effects were observed on haemolymphatic ions and ventilation.

Keywords: Gammarus fossarum, silver nanoparticles; gold nanoparticles; gene expression; osmoregulation;

behaviour; Cytoviva; long term exposure.


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1. INTRODUCTION

The recent advances in nanotechnologies led to an increasing use of engineered nanoparticles (ENPs) in

various consumers’ products. Silver nanoparticles (AgNPs) are among the most promising and most widely

used ENPs due to their broad spectrum antimicrobial activities (Vance et al., 2015). They are currently found

in many products of daily life such as textiles, plastics, health care products, water filters or food packaging.

Gold nanoparticles (AuNPs) are used in a wide range of applications in the field of biomedicine, biology and

chemistry (Nel et al., 2006; Volland et al., 2015) mostly because of their low toxicity and high bio-compatibility

(García-Cambero et al., 2013; Lapresta-Fernández et al., 2012). The increased production and use of AgNPs

and AuNPs has raised concerns about their release in the environment (Farkas et al., 2011; Kaegi et al., 2010).

The predicted environmental concentrations in surface water for both AgNPs and AuNPs are expected to be

in the range from 0.01 µg.L-1 to 0.32 µg.L-1 and from 0.14 µg.L-1 to 1.43 µg.L-1, respectively for AgNPs and

AuNPs (Gottschalk et al., 2013; Tiede et al., 2009; Yang et al., 2016). The NPs’ unique physical-chemical

properties in terms of size, shape, surface coatings and charges will substantially influence their behaviour,

fate and their effects/toxicity towards living organisms (Ju-Nam and Lead, 2008; McGillicuddy et al., 2017).

These properties may influence AgNPs and AuNPs entry into the cells (Nel et al., 2006; Oberdörster et al.,

2006; Volland et al., 2015) potentially leading to the generation of reactive species of oxygen (ROS) and

resulting in oxidative stress that has been reported as one of the principal mechanisms of toxicity of NPs in

aquatic organisms (Klaine et al., 2008; Vale et al., 2016; Volland et al., 2015).

Many studies have investigated the impact of AgNPs and AuNPs on key freshwater. Most of the studies

focused on acute toxicity through direct exposure (Buffet et al., 2013). However, in order to better predict

the potential effects of ENPs at higher biological level, such as population and community levels, it is

important to include dietary exposure and assess, through a multi-biomarker approach, the chronic toxicity

of these ENPs (Blinova et al., 2012; Gaiser et al., 2011; Mackevica et al., 2015). It is recognized that one

important uptake route for metals and particles is through the diet (Croteau et al., 2011; Luoma et al., 2014;

Luoma and Rainbow, 2005). Ag was found in the form of nanoparticle in the gut of polychaete exposed

through the diet to AgNPs (García-Alonso et al., 2014). Some studies reported effects of AgNPs and AuNPs

after trophic exposure of aquatic organisms. For instance, trophic exposure of the clam Corbicula fluminae

to AuNPs led to their presence in the epithelia tract of digestive gland and gills of the animals (Renault et al.,

2008). Additionally, an activation of phase II antioxidant enzymes encoding genes and an increase in

metallothionein gene expression was observed (Renault et al., 2008). Trophic exposure (15 days) of zebrafish

to sub-lethal concentrations of AgNPs 20 nm resulted in a significant Ag accumulation in Liver blood vessels

and in the interstitial tissue between the intestine and the liver. Gene expression profiles revealed that AgNPs


172

20 nm impacted photo-transduction system, circadian clock regulation and photoreception (Cambier et al.,

2018).

Amphipods of the genus Gammarus such as Gammarus fossarum represent an important part of the aquatic

macroinvertebrate assemblage (Dedourge-Geffard et al., 2009; Ladewig et al., 2006). Recently, G. fossarum

have been used as a model species due to their wide distribution in Europe (Janetzky, 1994) and their major

functional role in litter breakdown process and nutrient cycling (Forrow and Maltby, 2000; Kelly et al., 2002;

Lacaze et al., 2010; MacNeil et al., 1997). The ease of use, identification to the species level, differentiation

between gender, sampling and laboratory handling in addition to the well-documented sensitivity to different

kind of pollutants make Gammarus a good model organisms for ecotoxicological studies (Alonso et al., 2010;

Arce Funck et al., 2013; Felten et al., 2008; Mehennaoui et al., 2016). In our previous study, influence of size

and surface coatings on the acute toxicity of AgNPs and AuNPs were assessed (Mehennaoui et al., subm.,

Chapter 4). Acute exposure of G. fossarum to AgNPs and AuNPs showed:

• A coating dependent bio-accumulation, which was higher for citrate stabilized NPs as compared to

polyethylene glycol-coated (PEG) NPs;

• A metal dependent tissues distribution, with AgNPs found in G. fossarum gills while AuNPs were

found in intestinal caeca;

• An induction of antioxidant genes by AgNPs while AuNPs led to their down regulation.

However, the previous study only investigated the acute effects of AgNPs and AuNPs, but there is still a lack

of information about the chronic toxicity of AgNPs and AuNPs in G. fossarum. Hence, the aim of the present

study was to investigate the chronic toxicity of differently coated AgNPs and AuNPs 40 nm (citrate (CIT) and

polyethylene glycol (PEG) as surface coatings) on G. fossarum. Effects of AgNO3 were also studied in order to

determine the contribution of the released ions to the observed biological effects. A multibiomarker

approach including molecular effects (stress-related gene expression), physiological (osmoregulation) and

behavioural (locomotion and ventilation) responses was used (Mehennaoui et al., 2016). Biological endpoints

also included mortality, bioaccumulation and localization of AgNPs and AuNPs in G. fossarum tissues.

2. MATERIALS AND METHODS

2.1. Particles and Chemicals

Citrate stabilized AgNPs (CIT-AgNPs 40 nm) and citrate stabilized AuNPs (CIT-AuNPs 40 nm) were purchased

from Nanocomposix (San Diego, USA). Coating of AgNPs and AuNPs with PEG was performed at the LIST’s

Material Research and Technology (MRT) Department, (Luxembourg Institute of Science and Technology

(LIST), Luxembourg, Mehennaoui et al., subm., see Chapter 4). AgNO3 (CAS no.7761-88-8) was purchased


173

from VWR (Leuven, Belgium). Osmium tetroxide (OsO4, CAS no.20816-12-0), glutaraldehyde (CAS no.111-30-

8) and the Spurr low viscosity resin embedding kit were purchased from Sigma Aldrich (Bornem, Belgium).

Phosphate buffer saline (PBS) was obtained from Invitrogen (Merelbeke, Belgium).

2.2. Particle characterization

Size distribution of CIT-AgNPs, PEG-AgNPs, CIT-AuNPs and PEG-AuNPs 40 nm in Volvic water (exposure

medium), at T0h and T72h, were characterized using Nanoparticle Tracking Analysis (NTA, measured on a

NanoSIGHT instrument, Malvern Instrument Ltd, UK using validated protocols (Hole et al., 2013). Zeta

potential analyses are still on progress and will be performed on a Zeta-sizer Nano (Malvern, Hoeilaart,

Belgium).

2.3. Alder leaves conditioning

Leaves of Alnus glutinosa (Alder) were collected, air-dried and kept at room temperature until use. Leaves

were rinsed in deionized water for few minutes and cut into 16 mm diameter disks. Sets of 50 leaf disks were

placed into fine-net bags (0.5 mm pore size) and immersed in an unpolluted stream (49°48’24.9’’ N and

06°04’53.2’’ E, Schwaarzbaach, Colmar-berg, Luxembourg) (Dohet et al., 2008; Mehennaoui et al., 2016) for

10 days to allow microbial colonization. At the end of the immersion period, leaves were returned to the

laboratory where they were kept at -20 °C until use (Andreï et al., 2016; Batista et al., 2017).

2.4. Organisms sampling and acclimation

G. fossarum were collected in November 2016 at the same unpolluted site in Schwaarzbaach studied in our

previous study (Mehennaoui et al., 2016). Animals were collected using a hand net and they were sorted in

the field to separate males, which were used for the experiments, from females, which were immediately

returned to the water stream. Collected male animals were transported in river water to the laboratory once

collection was finished and they were kept at 12 °C. The decision to keep only adult males for the experiment

is justified by the fact that we wished to avoid influence of gender on the studied parameters (Arce Funck et

al., 2013; Sornom et al., 2010). G. fossarum males were selected from pre-copula pairs or based on sexual

dimorphism like gnatopode size (Mehennaoui et al., 2016). Adult males were then acclimated to laboratory

conditions (Andreï et al., 2016; Mehennaoui et al., 2016). The acclimation was conducted in 2 steps. First,

Gammarids were acclimated for 72h to mineral water (Volvic, France) by progressively changing field water

to Volvic water (30% v/v, 50% v/v, 100% v/v). Then, a stalling period of 10 days was conducted in 100% Volvic

water (Andreï et al., 2016; Mehennaoui et al., 2016). Acclimation was performed under controlled conditions

at 12 °C with a 16h light 8h dark photoperiod. 100% of Volvic water was aerated and changed every 24h to


174

avoid organic matter accumulation and potential increase of ammonium, nitrite and nitrate. Gammarids were

fed ad libitum with alder leaves (Alnus glutinosa) up to 24h before experiment.

2.5. Trophic exposure

2.5.1. Experimental design

In order to avoid as much as possible the adsorption of AgNO3, AgNPs 40nm and AuNPs 40 nm, the tanks

used for exposure were saturated with AgNO3, AgNPs 40 nm and AuNPs 40 nm for 96h before exposure

(Andreï et al., 2016).

At the end of the acclimation period, groups consisting of 15 males (compartment: animals) were transferred

into the pre-saturated plastic tanks (500 ml polypropylene tanks) containing 400 ml of exposure medium

(Volvic water with or without contaminants) (Mehennaoui et al., 2016) and 6 leave disks (compart: food). A

piece of mesh was added in each tank to provide a resting surface for G. fossarum and to reduce as much as

possible potential losses linked to cannibalism. Males were exposed to different treatments: AgNO3 (0 – 0.5

µg.L-1); CIT-AgNPs 40 nm, PEG-AgNPs 40 nm, CIT-AuNPs 40nm and PEG-AuNPs 40 nm (0 – 0.5 –5 µg.L-1) for

15 days at 12 °C with a photoperiod of 16h light and 8h darkness. The different ranges of concentrations

tested were selected based on previous results obtained by exposing G. fossarum to the same Ag and Au

nanoparticles for 72h (Mehennaoui et al., subm., see Chapter 4). AgNO3, AgNPs and AuNPs stock solutions

were diluted to the desired concentrations in mineral water (Volvic, France). 100% of exposure medium

(compartment: water colomn) were renewed every 72h to maintain the exposure conditions stable, to ensure

sufficient amount of nutrients and to remove waste products. Dead animals were removed and survivors

were counted every 24h. 6 leave disks per replicates were added every 48h.

2.5.2. Total and dissolved silver and gold measurements

In order to better describe the fate of Ag and Au in the studied compartment (water column, food or animals),

water samples were filtered on a nitrate cellulose filter (47 mm diameter, 0.45 µm pore size) previously rinsed

with Milliq water, dried for 24h at 80 °C and weighed. After filtration, the filters were dried for 24h at 80 °C

and weighed in order to determine the quantity of fine particle suspended matter produced after 72h of

exposure (Andreï et al., 2016). 5 mL of filtered water were kept for total Ag and Au measurement and 4 mL

of filtered water were used for ion release measurements and were centrifuged at 4000 g for 40 min using a

3KDa cut off membrane (Millipore) followed by a mineralization step with 1% HNO3 for Ag ion measurements

and 1% of HCl for Au ion measurements (Georgantzopoulou et al., 2013). The analyses are currently ongoing.

All measurements will be performed on an Inductively Coupled Plasma Mass spectrometer (ICP-MS, ElanDRC-

e, Perkin-Elmer, Waltham, MA, USA) as previously described (Mehennaoui et al., 2016).


175

2.5.3. Silver and gold bioaccumulation

At the end of the exposure period, three pools of 4 gammarids per each condition were rinsed with milliQ

water and dried on filter paper. Animals were frozen at -20 °C, freeze-dried and weighed. Samples were then

mineralized in HNO3 and H2O2 for Ag measurement and in acidic mixture of HNO3 and HCl for Au

measurement at a maximum pressure of 35 bars and maximum temperature of 200 °C using a microwave

system (Anton Paar Multiwave Pro) as previously described (Mehennaoui et al., 2016). All measurements

were performed in triplicates and are expressed in µg.g-1 of dry weight. A calibration curve was performed

following serial dilutions (from 0.01 to 50 µg.L-1 of Ag and from to for Au). Using a multi-element standard

solution (Chemlab, certified ISO/IEC 17025). The validity of the analytical method was controlled every 10

measurements within the same series of measurements with 3 quality controls (0.5, 5 and 25 µg.L-1 of Ag and

Au) of a second multi-element standard solution (Merck-VWR, ISO/IEC 17025). Ag and Au values were

consistent within the certified ranges (Cambier et al., 2018).

2.5.4. Particles uptake: Cytoviva ® analyses

2.5.4.1. Sample preparation

At the end of the exposure period, G. fossarum exposed to 5 µg. L-1 of AgNPs and AuNPs were fixed in 5%

(v/v) glutaraldehyde where they were kept at 4 °C until analyses. After a washing step with PBS, samples were

post-fixed with 1% osmium tetroxide for 2h at room temperature on a horizontal shaker (180 rpm). After a

second washing with PBS, samples were dehydrated with increasing concentrations of ethanol (10%, 20%,

30%, 50%, 70%, 90% and 100% v/v) and a final wash in acetone (100% v/v) before being embedded in Spurr

resin for 16h at 60 °C.

2.5.4.2. Cytoviva® dark field hyperspectral imaging

Semi-thin layers of 300 nm were cut using ultra-microtome and placed on microscope glass slide and let dry

for 1 min on a drying workstation. Slides were covered with a glass cover slip, sealed, and kept at 4 °C until

analysis. Visualization was performed using a Cytoviva® darkfield hyperspectral imaging system (Cytoviva Inc.,

Auburn, Alabama, USA) mounted on an Olympus BX-43 optical microscope. Images of the different tissues of

G. fossarum were captured at 60x magnification with oil immersion using hyperspectral camera controlled

by ENVI software (version 4.8 from Harris Corporation, Melbourne, FL, USA and modified by CytoViva, Inc.).

Spectral libraries of exposed animals were generated manually acquiring about 200 spectra per sample.

Acquired libraries were filtered against non-exposed samples to filter out all spectra non-related to AgNPs or

AuNPs using a spectral angle mapper (SAM) algorithm with a 0.1 radians tolerance. Filtered libraries were


176

mapped onto images of exposed samples using SAM with a 0.1 radians tolerance which allows highlighting

similarities between the spectra in the image and in the spectral library.

2.5.5. Molecular effects

2.5.5.1. RNA extraction and cDNA synthesis

At the end of the exposure period, animals were gently dried and dived in RNA lysis Buffer (RLT buffer)

supplemented with 1% (v/v) β-mercaptoethanol, flash-frozen in liquid nitrogen and stored at -80 °C until RNA

extraction. For RNA extraction, G. fossarum tissues were grinded on ice using a pellet pestle tissue grinder

and the homogenates were centrifuged at 250g for 5 min at 4 °C to remove cuticle fragments as previously

described (Mehennaoui et al., 2016). Total RNA was extracted using RNeasy mini kit for cells and animal

tissues (Qiagen, Leusden, The Netherlands), according to the manufacturer’s instructions (including digestion

with DNase). RNA concentrations and purity were assessed measuring the absorbance at 230, 260 and 280

nm using Nanodrop ND-1000 spectrophotometer (ThermoScientific, Villebon-sur-Yvette, France) (A260/280

and A260/230 ratios). RNA integrity was controlled using the RNA Nano 6000 assay (Agilent Technologies,

Diegem, Belgium) with 2100 Bioanalyzer (Agilent, Santa Clara, CA, USA) (Legay et al., 2015). For the synthesis

of cDNA, 1 µg of the extracted RNA were retro-transcribed into cDNA using the Protoscript II reverse

transcriptase (New England Biolabs, Leiden, The Netherlands) and random primers, according to the

manufacturer’s instructions.

2.5.5.2. Primer design and quantitative real-time PCR

All the primers for the RT-qPCR were designed using Primer3Plus (http://primer3.ut.ee) with the previously

described criteria (Mehennaoui et al., subm., chapter 3; Mehennaoui et al., subm., chapter 4). Primers were

analysed using NetPrimer (http://www.premierbiosoft.com/netprimer/) for secondary unexpected

structures. PCR efficiency was checked using decreasing five-fold dilutions from cDNA pool (From 25 ng to

0.04 ng and no template control). Melting curves were generated at the end of each experiment to assess

the specificity of the amplified products. All primers displayed one clear peak and were all retained for the

analysis. All the used primers are described in our previous study (Mehennaoui et al., subm., chapter 3 and

4)

cDNAs were diluted to 0.8 ng. µL-1 and used for RT-qPCR analyses in 384-well plates. An automated liquid

handling robot (epMotion 5073, Eppendorf, Hamburg, Germany) was used for the preparation of the 384-

well plate. cDNA were amplified using Takyon Low ROX SYBR MasterMix dTTP Blue Kit (Eurogentec, Liège,

Belgium) on a ViiA 7 Real-Time PCR System (Thermo Fisher, Waltham, MA, USA) in a final volume of 10 µL

(Behr et al., 2015). All reactions were performed in three technical replicates each one including four


177

biological replicates. The PCR conditions consisted on an initial denaturation step at 95 °C for 3 min, followed

by 40 cycles of denaturation at 95 °C for 15 sec and annealing/extension steps at 60 °C for 60 sec. Two genes,

namely Clathrin and SDH, were selected among six genes to be used as reference and they were validated

using the GeNorm module in the Biogazelle qBase PLUS software (Mehennaoui et al., subm., chapter 3). The

relative gene expression was calculated including PCR efficiency using Biogazelle qbase Plus software 2.5 with

the ΔΔCT method.

2.5.6. Behavioural responses and osmoregulation

For each treatment, measurements of locomotor activity, ventilation and osmoregulation were performed

on the same pool of 10 organisms (Mehennaoui et al., 2016).

2.5.6.1. Locomotor activity

At the end of the exposure period, locomotor activity was firstly assessed by counting the number of animals

in movement in a 80 mL glass tank containing 10 organisms with a piece of net added to provide a resting

surface (Felten et al., 2008). Measurements were performed after 5 min of acclimation at the same time of

the day with similar light conditions and in a quiet environment. Moving G. fossarum were counted for a

period of 2 sec and this process was repeated 40 times (Mehennaoui et al., 2016).

2.5.6.2. Ventilation activity

Ventilation activity was recorded immediately after locomotion on the same animals, by measuring the

frequency of pleopod beats. Ten gammarids from each treatment group were placed individually in a glass

tube containing Volvic water and left for a 30 sec acclimation period. Then, pleopod beats were visually

counted three times for 10 seconds, using a manual cell counter, only when animals were at rest.

Measurements were performed at the same period of the day to avoid possible effects of a circadian rhythm

on respiration (Felten et al., 2008; Mehennaoui et al., 2016; Rosas et al., 1992).

2.5.6.3. Osmoregulation

2.5.6.3.1. Haemolymph sampling

Immediately after behavioural response measurements, animals were dried on filter paper. Samples of

haemolymph were taken from the telson of each individual (n = 10) using a modified microcapillary (Felten

et al., 2008). To prevent haemolymph coagulation, samples were Kept at 4 °C until Na+, Cl- and Ca2+

measurements (Arce Funck et al., 2013).


178

2.5.6.3.2. Haemolymph Na+, Cl- and Ca2+ concentrations measurements

Samples of haemolymph (0.8 – 1.2 µL) were transferred to a gauged 5 µL microcapillary tube and were

centrifuged for 10 min at 6600 g at 4 °C. Samples were then diluted in 2.5 mL of MilliQ water to determine

Na+ and Ca2+ concentrations using atomic absorption spectrophotometry (Perkin Elmer Analyst 100) and Cl-

concentrations using ionic chromatography (Dionec 4500i equipped with an Ion Pac AS4A column) (Arce

Funck et al., 2013; Felten et al., 2008).

2.6. Statistical analyses

Survival, bioaccumulation, osmoregulation and behavioural response results are reported as mean ± SD

(standard deviation) and relative gene expression results are reported as mean ± SE (standard error). Results

of locomotor activity were obtained from the same set of individuals. Therefore, a one-way ANOVA with

repeated measurements following a general linear model was used as statistical test. The observed Ag and

Au bioaccumulation, survival, ventilation and gene expression on log2 transformed normalized relative

quantities were compared using a one-way ANOVA followed by Tukey-HSD post-hoc test for multiple

comparisons when normality and homogeneity of variances were respected. When this was not the case, a

non-parametric Kruskal-Wallis ANOVA followed by a Mann-Whitney U test was used. All statistical

calculations were performed using Statistica software 12 (StatSoft Inc.).

3. RESULTS

3.1. Particle character ization

In our previous study CIT-AgNPs, PEG-AgNPs 40 nm were the most stable in Volvic water with sizes of 45 ±

1.5 and 47 ± 1.3 nm, respectively (Mehennaoui et al., subm., chapter 4). CIT-AuNPs 40 nm had a size of 67 ±

13 nm after 24h in Volvic water, while PEG-AuNPs was stable after 24 hours in Volvic water with a size of 38

± 19 nm. In the present study, size distribution, which is measured using the NanoSIGHT, could not be

characterized due to the presence of food preventing the distinction between AgNPs, AuNPs and leave

particles. Thus, further analyses are needed in order to define the behaviour of AgNPs and AuNPs in Volvic

water in the presence of food.

3.2. Survival

A general tendency for a decrease in survival rates following exposure time (15 days) was observed in all the

exposure conditions (Figure 5. 1). However, only 0.5 µg. L-1 of AgNO3 led to a significant decrease in survival

rates of G. fossarum at day 15 (65 ± 3.3 %, One-way ANOVA, F= 11.63, 8df, P < 0.001) (Figure 5. 1). Survival


179

rates of G. fossarum tended to decrease with increasing exposure time to AgNPs and AuNPs even though

effects of AgNPs and AuNPs on survival were not statistically significant.

Figure 5. 1. Survival rates (mean ± SD) of G. fossarum exposed for 15 days to 0.5 and 5 µg. L-1 of CIT and PEG-

AgNPs and PEG-AuNPs 40 nm

3.3. Silver and gold b ioaccumulation

At the end of the exposure period, small but detectable amounts of Ag could be detected in the negative

control groups of G. fossarum (0.24 ± 0.1 µg. g-1 dry weight), for which the observed Ag body concentration

was 0.24 ± 0.01 µg. g-1 dry weight (Table 5. 1). Treatment with AgNO3 (0.5 µg. L-1) lead to the highest increase

of silver body concentrations that were of 5.16 ± 1.20 µg. g-1 d.w. Similarly, treatment with the AgNPs resulted

in a significant concentration-dependent Ag bioconcentration for both CIT-AgNPs and PEG-AgNPs 40 nm

(One-way ANOVA, F=45.12, 17df, P < 0.01). The highest concentrations (5 µg. L-1) of CIT-AgNPs and PEG-

AgNPs 40 nm led to a significant higher Ag bioconcentration in exposed animals compared to non-exposed

ones (P < 0.0001, Table 5.1).

The Au bio-concentration was under the limit of quantification (< 0.075 µg. g-1 d.w) in the non-exposed

animals. G. fossarum exposed to the highest concentration (5 µg. L-1) of CIT and PEG-AuNPs 40 nm tended to

accumulate more Au than those exposed to 0.5 µg. L-1 of the same AuNPs (P > 0.05, Table 5. 1).

0

20

40

60

80

100

120

0 0.5 5 0 0.5 5 0 0.5 5 0 0.5 5 0 0.5

CIT PEG CIT PEG


Survival(%)

Exposureconditions


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Table 5. 1. G. fossarum Ag and Au uptake (mean ± SD) after 15 days of exposure to CIT and PEG-AgNPs and

AuNPs 40 nm.

Different letters (a-d) indicate significant differences (one-way ANOVA + Tukey HSD post hoc test at P < 0.05). Bold

values indicate significant differences compared to control G. fossarum.

Nominalconcentrations

(µg.L-1)

Gammarus internal[Ag]and

[Au](µg.g-1gammaridsdry

weight)

0 0.24±0.01a

0.5 0.42±0.05ab

5 2.09±0.19c

0 0.24±0.01a

0.5 0.37±0.28ab

5 1.55±0.28bc

0 <LOQa

0.5 0.11±0.01a

5 1.03±0.18abc

0 <LOQa

0.5 0.10±0.03a

5 0.63±0.16ab

0 0.21±0.04a

0.5 5.16±1.21d

NPs

AgNO3

AgNPs40nm

CIT

PEG

AuNPs40nm

CIT

PEG


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3.4. Cytoviva® dark f ie ld hyperspectral imaging

A Cytoviva dark field hyperspectral imaging microscope was used in order to localize the internalized AgNPs

and AuNPs 40 nm. Currently only results for G. fossarum exposed to CIT-AgNPs and CIT-AuNPs 40 nm are

available (Figure 5. 2, Figure 5. 3).

Figure 5. 2 A) 300 nm cross section of G. fossarum observed under optical microscope. Red square represents

the area observed with cytoviva darkfield hyperspectral microscope. Scale bar is 250 µm. B) G. fossarum gills or

intestinal caeca viewed with Cytoviva (60x oil immersion magnification). C) AgNPs accumulation in G. fossarum

gills (red spots and white arrows). Animals were exposed for 72h to 10 µg. L-1 of CIT-AgNPs 40nm and PEG-AgNPs

40 nm in Volvic water. Scale bar is 6µm.

Analyses of samples exposed to CIT-AgNPs and CIT-AuNPs 40 nm were performed in different organs and

tissues (gills, gut and muscle). Tissues were selected based on the results obtained in the previous study

where AgNPs were found in the gills while AuNPs were found in the gut area of G. fossarum (Mehennaoui et

al., subm., chapter 4). In the present study, in addition to the long-term effects, the influence of food on the

internal distribution of AgNPs and AuNPs was investigated.

The analysis of Ag revealed the presence of AgNPs in the gills of G. fossarum exposed to CIT-AgNPs 40 nm

(Figure 5. 2C) while the analyses of Au showed the presence of AuNPs within the lumen and epithelial tissues

of intestinal caeca on G. fossarum exposed to CIT-AuNPs (Figure 5. 3C).

Cit-AgNPs

40 nm

Gills

Control

A B C


182


the area observed with CytoViva darkfield hyperspectral microscope. Scale bar is 250 µm. B) G. fossarum intestinal

caeca viewed with CytoViva (60x oil immersion magnification). C) AuNPs accumulation in G. fossarum caeca (red

spots and white arrows). Animals were exposed for 72h to 10 µg. L-1 of CIT-AgNPs 40nm and PEG-AgNPs 40 nm in

Volvic water. Scale bar is 6µm.

3.5. Molecular effects

In the present study, the same set of stress-related genes (Mehennaoui et al., subm., chapter 4) was tested

in order to define the influence of the presence of food and the longer exposure time (15 days) on the

molecular responses of G. fossarum to AgNPs and AuNPs 40 nm. Both concentrations (0.5 and 5 µg. L-1) of

PEG-AgNPs 40 nm led to an up-regulation of catalase and chitinase and CIT-AgNPs and PEG-AgNPs 40 nm led

to an up-regulation of Na+K+ATPase. The highest concentrations of CIT and PEG-AuNPs 40 nm (5 µg. L-1) led

to an increase of CuZnSOD gene expression (Table 5. 2).

3.6. Behavioural responses and osmoregulation

3.6.1. Locomotion and venti lation activity

G. fossarum exposed to CIT-AgNPs, PEG-AgNPs 40 nm, CIT-AuNPs and PEG-AuNPs 40 nm and AgNO3 showed

a significant reduced locomotor activity when compared to non-exposed animals (Figure 5. 4 A, GLM - one-

way ANOVA, P < 0.05). Animals exposed to 0.5 µg. L-1 of CIT-AgNPs 40 nm showed a significant reduced

locomotion activity when compared to G. fossarum exposed to 5 µg. L-1 CIT-AgNPs 40 nm and 0.5 µg. L-1 of

AgNO3 (GLM- Tukey-HSD post-hoc test, P < 0.05). At the highest concentration (5 µg. L-1), PEG-AgNPs 40 nm

led to a significant decrease in locomotor activity compared to CIT-AgNPs 40 nm (GLM- Tukey-HSD post-hoc

test, P < 0.05).

CIT-AuNPs

40 nm

Intestinal caeca

Control

A B C


183

Table 5. 2. Relative gene expression of G. fossarum exposed for 15 days to CIT- and PEG-AgNPs 40 nm and CIT and PEG-AuNPs 40 nm

Exoskeleton Lysosomes

Actin TUB UB Chitinase CAT CuZnSOD GPx7 GST HSP90 GAPDH Gadd45 Cjun CathL Ca2+ATPase NaKATPase HEM Cox1

0 1±0.18 1±0.13 1±0.06 1±0.14 1±0.12 1±0.32 1±0.08 1±0.17 1±0.12 1±0.1 1±0.05 1±0.12 1±0.19 1±0.15 1±0.14 1±0.12 1±0.26

0.5 1.44±0.29 1.33±0.28 1.16±0.13 0.71±0.17 0.73±0.17 0.66±0.16 0.83±0.08 1.5±0.21 1.07±0.16 0.77±0.08 1.04±0.05 1.37±0.11 1.54±0.35 0.89±0.44 1.13±0.08 0.95±0.04 1.07±0.14

Control 0 1±0.15 1±0.12 1±0.48 1±0.11 1±0.1 1±0.15 1±0.04 1±0.22 1±0.13 1±0.1 1±0.12 1±0.26 1±0.23 1±0.1 1±0.13 1±0.14 1±0.25

0.5 0.56±0.11 0.79±0.09 1.33±0.09 1.15±0.12 1.02±0.14 1.13±0.08 0.8±0.08 0.71±0.15 1.33±0.08 0.89±0.04 1.15±0.08 1.18±0.2 0.61±0.17 1.07±0.15 1.39±0.15 0.79±0.04 1.11±0.1

5 0.74±0.19 0.89±0.04 0.93±0.35 1.48±0.3 1.43±0.23 0.88±0.19 0.92±0.11 0.86±0.16 0.93±0.11 0.91±0.07 1.06±0.19 1.09±0.2 0.76±0.16 1.1±0.18 1.55±0.16 0.89±0.07 1.4±0.4

0.5 0.77±0.07 0.97±0.22 1.09±0.25 2.17±0.95 1.98±0.83 1.2±0.03 0.84±0.07 0.84±0.08 1.09±0.07 1.03±0.15 1.27±0.03 1.47±0.22 0.74±0.13 1.11±0.04 1.49±0.08 0.97±0.15 1.51±0.3

5 0.77±0.22 0.92±0.12 0.58±0.33 1.68±0.57 1.7±0.63 0.9±0.32 0.88±0.08 0.81±0.12 1.42±0.08 0.82±0.11 1.26±0.32 1.28±0.07 0.69±0.11 1.12±0.03 1.53±0.12 0.68±0.11 0.97±0.43

Control 0 1±0.19 1±0.15 1±0.22 1±0.11 1±0.06 1±0.24 1±0.11 1±0.13 1±0.11 1±0.09 1±0.24 1±0.07 1±0.15 1±0.05 1±0.13 1±0.09 1±0.39

0.5 1.02±0.16 0.93±0.22 0.96±0.06 1.2±0.2 1.31±0.24 1.71±0.35 1.04±0.16 1.02±0.09 0.66±0.16 0.91±0.11 0.89±0.35 1.05±0.13 1.09±0.23 0.97±0.06 0.95±0.09 0.83±0.11 0.68±0.15

5 1±0.21 0.77±0.13 0.85±0.06 1.46±0.13 1.43±0.09 1.21±0.28 1.05±0.12 0.83±0.14 0.86±0.12 1.11±0.04 1.15±0.28 1.43±0.27 0.88±0.16 0.89±0.06 0.81±0.14 1.18±0.04 0.97±0.36

0.5 0.97±0.04 0.61±0.24 1.23±0.05 1.29±0.23 1.24±0.27 1.25±0.18 0.99±0.08 0.87±0.04 0.69±0.08 1±0.14 0.81±0.18 0.83±0.15 0.82±0.09 0.79±0.07 0.66±0.04 1.23±0.14 1.12±0.15

5 1.26±0.2 0.94±0.1 1±0.14 1.42±0.27 1.44±0.23 2.14±0.54 1.01±0.08 1.13±0.14 0.96±0.08 0.68±0.17 1.06±0.54 1.27±0.17 1.31±0.27 0.88±0.07 0.8±0.14 0.92±0.17 1.15±0.56

Generalstress DNAdamageandrepair Osmoregulation Respiration

AuNPs40nmCitrate

PEG

Cytoskeleteon Antioxidantdefensecompound concentrations(µg.L

-1)

AgNO3

AgNPs40nmCitrate

PEG


184

A significant decrease in the number of moving Gammarids was observed when animals were exposed to 0.5

µg. L-1 of CIT-AuNPs 40 nm compared to those exposed to 5 µg. L-1 of CIT-AuNPs 40 nm (GLM- Tukey-HSD post-

hoc test, P < 0.05) while no significant effects were observed in Gammarids exposed to PEG-AuNPs 40 nm

(GLM- Tukey-HSD post-hoc test, P > 0.05).

Pleopod beats frequency of G. fossarum exposed to 5 µg. L-1 of CIT-AgNPs 40 nm and CIT-AuNPs 40 nm as not

significantly affected by AgNPs, AuNPs and AgNO3 (Figure 5. 4 B, Kruskal-Wallis ANOVA, P < 0.05).

Figure 5. 4. Behavioural responses of G. fossarum exposed for 15 days to CIT and PEG-AgNPs and AuNPs 40 nm.

A) Locomotor activity (mean percentage of moving G. fossarum ± SD) and b) Ventilation (mean pleopods beat

frequency ± SD). Different letters (a-f) indicates significant differences (one-way ANOVA + Tukey HSD post hoc test

at P < 0.05 level of significance, n = 10).

0

10

20

30

40

50

60

70

80

90

100

0 0.5 5 0 0.5 5 0 0.5 5 0 0.5 5 0 0.5

CIT PEG CIT PEG


%Gammarusinmouvement

Exposureconditions(µg.L-1)

a

a ab

f

c

c

c

cd

ee

ff

de

f

A

0

50

100

150

200

250

300

350

400

450

0 0.5 5 0 0.5 5 0 0.5 5 0 0.5 5 0 0.5

CIT PEG CIT PEG


Ventilation(Pleopodsbeatperminute)

Exposureconditions(µg.L-1)

abab

abab ab

b

aa

ab

ab

ab

abab b

B


185

3.6.2. Osmoregulation

The basal level of haemolymphatic Cl- in control gammarids was constant with values of 70.4, 101.9 and 95.5

mmol. L-1 in the three different control groups. The basal level of haemolymphatic Na+ was 108.8, 169.4 and

114.2 mmol. L-1 in the non-exposed animals. Basal level of haemolymphatic Ca2+ were 39.7, 72.7, and 28.7

mmol. L-1 in the non-exposed animals (Table 5. 3). No significant effects of AgNPs and AuNPs 40 nm were

observed on the concentration of the different haemolymphatic ions indicating that osmoregulation of G.

fossarum was not impacted by the applied treatment (Kruskal-Wallis ANOVA, P > 0.05).

Table 5. 3. Haemolymph [Cl-], [Na+] and [Ca2+] of G. fossarum exposed for 15 days to CIT and PEG-AgNPs 40 nm

and CIT and PEG-AuNPs 40 nm.

4. Discussion

The chronic toxicity of AgNPs and AuNPs on G. fossarum is still not fully understood. Hence, in the present

study we investigated the chronic effects of differently coated (CIT and PEG) AgNPs and AuNPs 40 nm on G.

fossarum. Individuals were exposed to realistic environmental concentrations (0.5 and 5 µg. L-1) of AgNPs and

AuNPs 40 nm for 15 days and in presence of food (leaves disk of A. glutinosa). AgNPs and AuNPs 40 nm were

selected based on the results of our previous study (Mehennaoui et al., subm., chapter 4). AgNPs and AuNPs

40 nm were selected as they were the most stable in exposure medium and the NPs with the highest uptake

rate by G. fossarum as described in our previous study (Mehennaoui et al., subm. Chapter 4). The studied

endpoints include mortality, bioaccumulation, tissue distribution, gene expression, osmoregulation and

behavioural responses (locomotion and ventilation).

Nominal

concentrations

(µg.L-1)

haemolymph[Cl-]

(mmol.L-1)

haemolymph[Na+]

(mmol.L-1)

haemolymph[Ca2+]

(mmol.L-1)

0 70.4±31 108.9±42 39.7±5

0.5 66.4±22 103.4±37 41.6±39

5 69.2±12 110.0±27 31.9±12

0 70.4±31 108.9±42 39.7±5

0.5 82.1±7 123.2±7 36.3±7

5 78.5±23 129.4±17 49.2±27

0 101.9±18 169.4±54 72.7±33

0.5 102.8±8 125.3±17 36.8±20

5 86.5±5 137.5±22 40.2±4

0 101.9±18 169.4±54 72.7±33

0.5 84.5±10 137.4±9 34.6±5

5 116.3±20 130.8±22 22.5±3

0 95.6±23 114.3±17 28.7±7

0.5 75.3±24 105.5±40 107.8±62

AuNPs40nm

CIT

PEG

AgNO3

NPs

AgNPs40nm

CIT

PEG


186

4.1. Particle character ization

AgNPs and AuNPs 40 nm were characterized in the exposure media (Volvic water) that contained food debris.

For the characterization of the size distribution in water we, first, used the nanoparticle tracking analysis

(NTA), which, however, did not allow a complete and relevant characterization of the studied NPs at the used

concentrations (0.5 and 5 µg. L-1) since the presence of the food debris interferes with the detection of

particles. Further analyses using NanoZetasizer are on progress. Thus, the necessity of feeding in the present

experiment can be considered as an influencing/confounding factor on the behaviour of AgNPs and AuNPs

as observed in previous studies (Mackevica et al., 2015; Sørensen et al., 2016). Potentially AgNPs and AuNPs

interact with the different compound present in the exposure medium and may be complexed or aggregated

(Andreï et al., 2016; Levard et al., 2012; Petersen et al., 2014) but further analyses are needed in order to

verify this hypothesis. This behaviour makes the commonly used techniques for characterization (DLS, NTA

and electron microscopy) unsuitable for environmental samples analysis due to their detection limits (Farré

et al., 2008; Gallego-Urrea et al., 2011; Yang et al., 2016). One possibility is to test new techniques such as

the combination of the ORION NanoFab helium ion microscope (HIM) with a secondary ion mass

spectrometer (SIMS) that may produce an elemental SIMS maps with a high resolution images (Dowsett et

al., 2016). This technique might allow distinction between metallic element and carbon element although it

might not allow the determination of the size distribution of AgNPs and AuNPs.

4.2. Biological endpoints

Trophic exposure of G. fossarum for 15 days to CIT and PEG-AgNPs and CIT-and PEG-AuNPs 40 nm led to a

concentration-dependent uptake of Ag and Au. These findings are in agreement with what was already

described in our previous study in which G. fossarum were exposed for 72h to an increasing gradient of

different sizes of differently coated AgNPs and AuNPs (Mehennaoui et al., subm., chapter4; Mehennaoui et

al., 2016). Dose-dependent accumulation was also reported in G. roeseli exposed for 72h to 0.5 and 5 µg.L-1

of CIT-AgNPs 10 and 60 nm (Andreï et al., 2016). G. fossarum exposed to the highest concentration of CIT-

AuNPs 40 nm (5 µg. L-1) tended to accumulate more Au than non-exposed animals and those exposed to PEG-

AgNPs and PEG-AuNPs. Similar observations were done in previous studies reporting a rapid uptake of Au by

the clams Ruditapes philippinarum exposed for 7 days to 0.75 µg.L-1 of CIT-AuNPs 20 nm and Scorbicularia

plana exposed for 16 days to 100 µg.L-1 of AuNPs 40 nm (Pan et al., 2012). The internal concentrations of Ag

tended to be higher than those of Au which could be linked to different metal speciation in the exposure

medium and/or their affinity to the binding sites (Sørensen et al., 2016). The uptake of Ag and Au after a

dietary exposure in G. fossarum may be relevant of a potential transfer of these NPs in the food chain.


187

AgNO3 led to the highest uptake compared to AgNPs and was associated with a significant mortality rate in

G. fossarum, which is consistent to the short-term exposure (Mehennaoui et al., 2016). Similarly, other

studies reported a significant uptake associated with a significant mortality of G. roeseli exposed to AgNO3

for 72h (Andreï et al., 2016) and G. fossarum exposed to up to 8 µg. L-1 of AgNO3 for 96h (Arce Funck et al.,

2013). However, despite an uptake of Ag and Au, no significant effects on survival were observed when

animals were exposed to CIT and PEG-AgNPs 40 nm and CIT and PEG-AuNPs 40 nm. The internal

concentration of Ag recorded in G. fossarum reached only 2 µg.g-1 gammarid d.w. which is less than the toxic

threshold of 8 µg.g-1 gammarid d.w. previously reported in G. fossarum (Arce Funck et al., 2013). Observed

differences in toxicities are most likely due to the ratio of NPs to free ions which has been shown to be an

important factor for toxicity. NPs used in the current study have low dissolution of below 6% (Mehennaoui

et a., subm., chapter 4) which may indicate the low toxicity observed for AgNPs.

Knowing the behaviour of AgNPs and AuNPs, once they are taken up by an organism, is crucial for better

definition of their fate and their potential toxic mechanisms. Thus, the internal distribution of AgNPs and

AuNPs was evaluated using enhanced darkfield hyperspectral microscopy. AgNPs were observed in the gills

of G. fossarum exposed to CIT-AgNPs 40 nm similar to short-term exposure (Mehennaoui et al., subm.,

chapter 4). However, in the presence of food during exposure of the organisms less AgNPs were observed

compared to our previous study with exposure via water. CIT-AuNPs 40 nm were observed within the lumen

of intestinal caeca of G. fossarum. This may suggest that AuNPs were ingested with the food and might not

cross the intestinal barrier and might be eliminated with faeces. This observation may also explain the

absence of significant effects of AuNPs on G. fossarum survival rates.

Acute waterborne exposure of G. fossarum to CIT-AgNPs and CIT-AuNPs 40 nm showed an uptake of AgNPs

in the gills and AuNPs in membrane of intestinal caeca of AuNPs (Mehennaoui et al., subm., chapter4). The

necessity of feeding in the present work had a significant effect on the pattern of internal distribution of

AgNPs and AuNPs. Fewer AgNPs were observed in G. fossarum gills when compared to gammarids exposed

to AgNPs in the absence of food (Mehennaoui et al., subm. Chapter 4), while AuNPs were observed only

within the lumen of intestinal caeca which may suggest a rapid elimination of AuNPs with faeces. AuNPs were

observed in the gut of D. magna after direct exposure and were not eliminated while the presence of food

during the experiment resulted in their elimination (Botha et al., 2016). Other study reported an adsorption

of AuNPs on algae surface which led to their ingestion by D. magna followed by a rapid elimination from the

gut (Khan et al., 2014; Volland et al., 2015; Wray and Klaine, 2015).

No significant effects of AgNPs or AuNPs on concentrations of haemolymphatic ions, namely Na+, Cl- and Ca2+,

were observed. As described above, only small amounts of AgNPs were observed in G. fossarum gills

indicating a potential lower Ag uptake through the gills. The dietary exposure may have decreased the


188

bioavailability of Ag, leading to the formation of different complex with biomolecules, thus in turn making Ag

less toxic or favouring a different uptake route (Mann et al., 2004; Webb and Wood, 1998). Thus, the low

presence of AgNPs in the gills may explain the absence of significant effects on concentrations of

haemolymphatic ions (Mann et al., 2004). Similarly, the presence of AuNPs within the lumen of intestinal

caeca of G. fossarum may indicate that these compounds are likely to be eliminated with faeces without

crossing intestinal barrier and reaching the circulatory system. Gammarids exposed to metallic contaminants

are usually described to exhibit an iono/osmotic disruption including changes in the Na+K+ATPase activity,

ions influx and gills surface permeability (Lignot et al., 2000). Osmoregulation impairment can therefore be

noticed by an impact on haemolymphatic Na+ and Cl- concentrations which represent 90% of the osmotic

pressure in gammarids (Mantel and Farmer, 1983). It is also known that an acute exposure to Ag generally

lead to the inhibition of Na+ uptake through the inhibition of Na+K+ATPase activity (Bianchini and Wood, 2002,

2003; Bury et al., 2002; Grosell et al., 2002; Webb and Wood, 1998). This mechanism was previously observed

in G. fossarum exposed to AgNO3 and AgNPs 23 nm for up to 96h (Arce Funck et al., 2013; Mehennaoui et al.,

2016). Even though, the major haemolymphatic ions were not impacted by AgNPs and AuNPs, an

osmoregulation impairment could occur as other elements present in haemolymph, such as amino-acids,

may be affected (Felten et al., 2008) and thus, further analyses are needed to better define the chronic effects

of AgNPs and AuNPs at the physiological level of G. fossarum.

A set of stress-related genes was used in order to investigate the effects of AgNPs and AuNPs 40 nm at the

molecular level (Mehennaoui et al., subm., chapter 3-4.). Both concentrations (0.5 and 5 µg. L-1) of PEG-AgNPs

40 nm led to an up-regulation of catalase and chitinase and CIT-AgNPs and PEG-AgNPs 40 nm led to an up-

regulation of Na+K+ATPase. The highest concentrations of CIT and PEG-AuNPs 40 nm (5 µg. L-1) led to an

increase of CuZnSOD gene expression. One previous study reported an over-expression of SOD and catalase

in G. fossarum exposed for 48h to 4.56 mg.L-1 of positively charged AuNPs 10 nm (Baudrimont et al., 2017).

Authors linked the up-regulation of antioxidant defense to the increase of the number of mitochondria,

observed via induction of 16S gene, as these mechanisms require energy (Baudrimont et al., 2017). Similarly,

catalase was induced in Chironomus riparus exposed to up to 1 mg.L-1 of AgNPs (Nair et al., 2013). At a higher

biological level, an activation of phase II antioxidant enzymes after exposing R. philippinarum to 0.75 µg.L-1 of

CIT-AuNPs 20 nm for 24h was reported (Volland et al., 2015). However, by increasing the exposure time,

authors observed significant alterations in the gene expression of GPx and genes related to inflammatory

responses (Gadd45) as well as MT (Volland et al., 2015). Additionally, digestive gland of bivalve appeared as

the target organ for AuNPs under short-term exposure. 1 day of exposure led to the activation of phase II

antioxidant enzymes (GST, CAT, GPx and GR) (Volland et al., 2015) while 7 days of exposure led to a persistent

oxidative stress linked to an increase in GPx and SOD expression and a decrease of Gadd 45 (Volland et al.,


189

2015). However, the absence of effects on CAT, GPx and GR in addition to an activation of MT may suggest

an adaptative handling and/or detoxification strategies due to the low doses of AuNPs administered for a

prolonged exposure time (Pan et al., 2012; Volland et al., 2015). This hypothesis may also explain the increase

in Na+K+ATPase gene expression in G. fossarum exposed to AgNPs. As previously described, the acute toxicity

of Ag is exerted through the inhibition of Na+K+ATPase activity. A prolonged exposure may have led to an

adaptative strategy leading to the up-regulation of Na+K+ATPase in order to offset the inhibition of the

enzymatic activity. These results highlight the differences or evolution of gene expression responses observed

after an acute waterborne exposure of G. fossarum to AgNPs and AuNPs (Mehennaoui et al., subm., chapter

4) and chronic exposure in the presence of food to the same particles.

Exposing G. fossarum for 15 days to CIT and PEG-AgNPs 40 nm led to a significant decrease in the number of

G. fossarum in movement. This result is in accordance with our previous finding as a decrease in locomotor

activity was observed in G. fossarum exposed for 72h to AgNPs 23 nm (Mehennaoui et al., 2016). Similarly, a

decrease in locomotor activity of G. roeseli exposed for 72h to CIT-AgNPs 10 and 60 nm was observed (Andreï

et al., 2016). Authors reported a size-dependent effect on locomotion as CIT-AgNPs 10 nm was more effective

than CIT-AgNPs 60 nm (Andreï et al., 2016). The effects of CIT-AgNPs 10 nm was as strong as the effects

observed for AgNO3 (Andreï et al., 2016; Arce Funck et al., 2013). Therefore, it may be suggested that the

readily taken up AgNPs dissolve internally and release Ag+ ions that are more potent in inhibiting G. fossarum

locomotor activity (Arce Funck et al., 2013; Mehennaoui et al., 2016). The effects on locomotion could also

be linked to sensorial disruption linked to the adsorption of AgNPs and AuNPs on the carapace of G. fossarum

as observed in our previous study (Mehennaoui et al., subm., chapter 4) and which is consistent with the

significant effect of AgNPs on the expression of chitinase. The decrease in locomotor activity could also be a

result of an energy reallocation in favour of defence mechanisms. Indeed, an increase in the internal

concentrations of metals or NPs usually lead individuals to invest their energy in highly energy consuming

mechanisms such as detoxification, homeostasis maintenance (iono/osmoregulation) or cellular repairing

process (Arce Funck et al., 2013; Vellinger et al., 2013).

This study highlights once more the great sensitivity of behavioural responses that are fast, simple to perform,

cheap and non-invasive biological endpoints that can highlight sub-lethal effects of contaminants.

Behavioural responses allow linking responses observed at the physiological level to potential effects at the

population and community levels regarding the central position of G. fossarum in aquatic ecosystems.

Indeed, an impairment of locomotion may have consequent effects on the fitness of an organism and lead to

“ecological death” (Scott and Sloman, 2004).


190

5. CONCLUSIONS

The present study allowed obtaining first insights on key mechanisms of sub-chronic effects of AgNPs and

AuNPs. Gene expression and locomotion appeared good biological endpoins as they allow the detection of

sub-lethal effects of AgNPs and AuNPs at very low concentration (0.5 µg. L-1). Similarly to the acute exposure

(Chapter 4), chemical composition of AgNPs and AuNPs appear as the main factor influencing the fate and

effects of these compounds on G. fossarum. Indeed, different patterns of internal distribution were observed

as AgNPs were observed in the gills whereas AuNPs were localized within the lumen of intestinal caeca

indicating potential elimination with faeces. Gene expression results may allow an early prediction of effects

of AgNPs and AuNPs on G. fossarum. Locomotion may predict effects at the population and community level

regarding the central position of G. fossarum in aquatic ecosystems.

Acknowledgments

This work was supported by National Research Fund, Luxembourg (AFR-PhD-9229040). The authors are

thankful to S. Contal for technical support, P. Rouselle, C. Guignard and J. Ziebel for chemical analyses and A.

Chauvière for ultramicrotome sections.

REFERENCES

Allen, H.J., Impellitteri, C.A., Macke, D.A., Heckman, J.L., Poynton, H.C., Lazorchak, J.M., Govindaswamy, S.,

Roose, D.L., Nadagouda, M.N., 2010. Effects from filtration, capping agents, and presence/absence of food

on the toxicity of silver nanoparticles to Daphnia magna. Environ. Toxicol. Chem. 29, 2742–2750.

doi:10.1002/etc.329

Alonso, Á., Lange, H.J.D., Peeters, E.T.H.M., 2010. Contrasting sensitivities to toxicants of the freshwater

amphipods Gammarus pulex and G. fossarum. Ecotoxicology 19, 133–140. doi:10.1007/s10646-009-0398-y

Andreï, J., Pain-Devin, S., Felten, V., Devin, S., Giambérini, L., Mehennaoui, K., Cambier, S., Gutleb, A.C.,

Guérold, F., 2016. Silver nanoparticles impact the functional role of Gammarus roeseli (Crustacea

Amphipoda). Environ. Pollut. 208, Part B, 608–618. doi:10.1016/j.envpol.2015.10.036


physiological responses of Gammarus fossarum (Crustacea Amphipoda) exposed to silver. Aquat. Toxicol.

142–143, 73–84. doi:10.1016/j.aquatox.2013.07.012

Batista, D., Pascoal, C., Cássio, F., 2017. How do physicochemical properties influence the toxicity of silver

nanoparticles on freshwater decomposers of plant litter in streams? Ecotoxicol. Environ. Saf. 140, 148–155.

doi:10.1016/j.ecoenv.2017.02.039




doi:10.1007/s11356-017-8400-3


191

Behr, M., Legay, S., Hausman, J.-F., Guerriero, G., 2015. Analysis of Cell Wall-Related Genes in Organs of

Medicago sativa L. under Different Abiotic Stresses. Int. J. Mol. Sci. 16, 16104–16124.

doi:10.3390/ijms160716104

Bianchini, A., Wood, C.M., 2003. Mechanism of acute silver toxicity in Daphnia magna. Environ. Toxicol. Chem.

22, 1361–1367. doi:10.1002/etc.5620220624

Bianchini, A., Wood, C.M., 2002. Physiological effects of chronic silver exposure in Daphnia magna. Comp.

Biochem. Physiol. Toxicol. Pharmacol. CBP 133, 137–145.

Blinova, I., Niskanen, J., Kajankari, P., Kanarbik, L., Käkinen, A., Tenhu, H., Penttinen, O.-P., Kahru, A., 2012.

Toxicity of two types of silver nanoparticles to aquatic crustaceans Daphnia magna and Thamnocephalus

platyurus. Environ. Sci. Pollut. Res. 1–8. doi:10.1007/s11356-012-1290-5

Botha, T.L., Boodhia, K., Wepener, V., 2016. Adsorption, uptake and distribution of gold nanoparticles in

Daphnia magna following long term exposure. Aquat. Toxicol. 170, 104–111.

doi:10.1016/j.aquatox.2015.11.022

Buffet, P.-E., Pan, J.-F., Poirier, L., Amiard-Triquet, C., Amiard, J.-C., Gaudin, P., Faverney, C.R., Guibbolini, M.,

Gilliland, D., Valsami-Jones, E., Mouneyrac, C., 2013. Biochemical and behavioural responses of the

endobenthic bivalve Scrobicularia plana to silver nanoparticles in seawater and microalgal food. Ecotoxicol.


Bury, N.R., Shaw, J., Glover, C., Hogstrand, C., 2002. Derivation of a toxicity-based model to predict how water

chemistry influences silver toxicity to invertebrates. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol. 133,

259–270. doi:10.1016/S1532-0456(02)00096-0

Cambier, S., Røgeberg, M., Georgantzopoulou, A., Serchi, T., Karlsson, C., Verhaegen, S., Iversen, T.-G.,

Guignard, C., Kruszewski, M., Hoffmann, L., Audinot, J.-N., Ropstad, E., Gutleb, A.C., 2018. Fate and effects of

silver nanoparticles on early life-stage development of zebrafish (Danio rerio) in comparison to silver nitrate.

Sci. Total Environ. 610, 972–982. doi:10.1016/j.scitotenv.2017.08.115

Croteau, M.-N., Misra, S.K., Luoma, S.N., Valsami-Jones, E., 2011. Silver Bioaccumulation Dynamics in a

Freshwater Invertebrate after Aqueous and Dietary Exposures to Nanosized and Ionic Ag. Environ. Sci.

Technol. 45, 6600–6607. doi:10.1021/es200880c

Dedourge-Geffard, O., Palais, F., Biagianti-Risbourg, S., Geffard, O., Geffard, A., 2009. Effects of metals on

feeding rate and digestive enzymes in Gammarus fossarum: An in situ experiment. Chemosphere 77, 1569–

1576. doi:10.1016/j.chemosphere.2009.09.042

Dohet, A., Ector, L., Cauchie, H.-M., Hoffmann, L., 2008. Identification of benthic invertebrate and diatom

indicator taxa that distinguish different stream types as well as degraded from reference conditions in

Luxembourg. Anim. Biol. 58, 419–472. doi:10.1163/157075608X383719

Dowsett, D., Audinot, J.-N., Vollnhals, F., Eswara, S., Wirtz, T., 2016. High Resolution Chemical Imaging on the

Helium Ion Microscope, in: European Microscopy Congress 2016: Proceedings. Wiley-VCH Verlag GmbH &

Co. KGaA. doi:10.1002/9783527808465.EMC2016.6559

Farkas, J., Christian, P., Gallego-Urrea, J.A., Roos, N., Hassellöv, M., Tollefsen, K.E., Thomas, K.V., 2011. Uptake

and effects of manufactured silver nanoparticles in rainbow trout (Oncorhynchus mykiss) gill cells. Aquat.



192

Farré, M., Gajda-Schrantz, K., Kantiani, L., Barceló, D., 2008. Ecotoxicity and analysis of nanomaterials in the

aquatic environment. Anal. Bioanal. Chem. 393, 81–95. doi:10.1007/s00216-008-2458-1


Physiological and behavioural responses of Gammarus pulex (Crustacea: Amphipoda) exposed to cadmium.

Aquat. Toxicol. 86, 413–425. doi:10.1016/j.aquatox.2007.12.002

Forrow, D.M., Maltby, L., 2000. Toward a mechanistic understanding of contaminant-induced changes in

detritus processing in streams: Direct and indirect effects on detritivore feeding. Environ. Toxicol. Chem. 19,

2100–2106. doi:10.1002/etc.5620190820

Gaiser, B.K., Biswas, A., Rosenkranz, P., Jepson, M.A., Lead, J.R., Stone, V., Tyler, C.R., Fernandes, T.F., 2011.

Effects of silver and cerium dioxide micro- and nano-sized particles on Daphnia magna. J. Environ. Monit. 13,

1227–1235. doi:10.1039/C1EM10060B

Gallego-Urrea, J.A., Tuoriniemi, J., Hassellöv, M., 2011. Applications of particle-tracking analysis to the

determination of size distributions and concentrations of nanoparticles in environmental, biological and food

samples. TrAC Trends Anal. Chem., Characterization, Analysis and Risks of Nanomaterials in Environmental

and Food Samples II 30, 473–483. doi:10.1016/j.trac.2011.01.005

García-Alonso, J., Rodriguez-Sanchez, N., Misra, S.K., Valsami-Jones, E., Croteau, M.-N., Luoma, S.N., Rainbow,

P.S., 2014. Toxicity and accumulation of silver nanoparticles during development of the marine polychaete

Platynereis dumerilii. Sci. Total Environ. 476–477, 688–695. doi:10.1016/j.scitotenv.2014.01.039

García-Cambero, J.P., Núñez García, M., López, G.D., Herranz, A.L., Cuevas, L., Pérez-Pastrana, E., Cuadal, J.S.,

Castelltort, M.R., Calvo, A.C., 2013. Converging hazard assessment of gold nanoparticles to aquatic organisms.

Chemosphere 93, 1194–1200. doi:10.1016/j.chemosphere.2013.06.074

Georgantzopoulou, A., Balachandran, Y.L., Rosenkranz, P., Dusinska, M., Lankoff, A., Wojewodzka, M.,

Kruszewski, M., Guignard, C., Audinot, J.-N., Girija, S., Hoffmann, L., Gutleb, A.C., 2013. Ag nanoparticles: size-

and surface-dependent effects on model aquatic organisms and uptake evaluation with NanoSIMS.

Nanotoxicology 1–11. doi:10.3109/17435390.2012.715312

Gottschalk, F., Sun, T., Nowack, B., 2013. Environmental concentrations of engineered nanomaterials: Review

of modeling and analytical studies. Environ. Pollut. 181, 287–300. doi:10.1016/j.envpol.2013.06.003

Grosell, M., Nielsen, C., Bianchini, A., 2002. Sodium turnover rate determines sensitivity to acute copper and

silver exposure in freshwater animals. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol. 133, 287–303.

doi:10.1016/S1532-0456(02)00085-6

Hole, P., Sillence, K., Hannell, C., Maguire, C.M., Roesslein, M., Suarez, G., Capracotta, S., Magdolenova, Z.,

Horev-Azaria, L., Dybowska, A., Cooke, L., Haase, A., Contal, S., Manø, S., Vennemann, A., Sauvain, J.-J.,

Staunton, K.C., Anguissola, S., Luch, A., Dusinska, M., Korenstein, R., Gutleb, A.C., Wiemann, M., Prina-Mello,

A., Riediker, M., Wick, P., 2013. Interlaboratory comparison of size measurements on nanoparticles using

nanoparticle tracking analysis (NTA). J. Nanoparticle Res. 15, 1–12. doi:10.1007/s11051-013-2101-8

Janetzky, W., 1994. Distribution of the genus Gammarus (Amphipoda: Gammaridae) in the River Hunte and

its tributaries (Lower Saxony, northern Germany). Hydrobiologia 294, 23–34. doi:10.1007/BF00017622




193

Kaegi, R., Sinnet, B., Zuleeg, S., Hagendorfer, H., Mueller, E., Vonbank, R., Boller, M., Burkhardt, M., 2010.

Release of silver nanoparticles from outdoor facades. Environ. Pollut. 158, 2900–2905.

doi:10.1016/j.envpol.2010.06.009



Khan, F.R., Kennaway, G.M., Croteau, M.-N., Dybowska, A., Smith, B.D., Nogueira, A.J.A., Rainbow, P.S.,

Luoma, S.N., Valsami-Jones, E., 2014. In vivo retention of ingested Au NPs by Daphnia magna: No evidence

for trans-epithelial alimentary uptake. Chemosphere. doi:10.1016/j.chemosphere.2013.12.051

Klaine, S.J., Alvarez, P.J.J., Batley, G.E., Fernandes, T.F., Handy, R.D., Lyon, D.Y., Mahendra, S., McLaughlin,

M.J., Lead, J.R., 2008. Nanomaterials in the environment: Behavior, fate, bioavailability, and effects. Environ.

Toxicol. Chem. 27, 1825–1851. doi:10.1897/08-090.1

Lacaze, E., Geffard, O., Bony, S., Devaux, A., 2010. Genotoxicity assessment in the amphipod Gammarus

fossarum by use of the alkaline Comet assay. Mutat. Res. Toxicol. Environ. Mutagen. 700, 32–38.

doi:10.1016/j.mrgentox.2010.04.025

Ladewig, V., Jungmann, D., Köhler, H.-R., Schirling, M., Triebskorn, R., Nagel, R., 2006. Population Structure

and Dynamics of Gammarus fossarum (Amphipoda) Upstream and Downstream from Effluents of Sewage

Treatment Plants. Arch. Environ. Contam. Toxicol. 50, 370–383. doi:10.1007/s00244-005-7039-0

Lam, I.K.S., Wang, W.-X., 2006. Accumulation and elimination of aqueous and dietary silver in Daphnia magna.




Legay, S., Guerriero, G., Deleruelle, A., Lateur, M., Evers, D., André, C.M., Hausman, J.-F., 2015. Apple

russeting as seen through the RNA-seq lens: strong alterations in the exocarp cell wall. Plant Mol. Biol. 88,

21–40. doi:10.1007/s11103-015-0303-4

Levard, C., Hotze, E.M., Lowry, G.V., Brown, G.E., 2012. Environmental Transformations of Silver

Nanoparticles: Impact on Stability and Toxicity. Environ. Sci. Technol. 46, 6900–6914. doi:10.1021/es2037405

Lignot, J.-H., Spanings-Pierrot, C., Charmantier, G., 2000. Osmoregulatory capacity as a tool in monitoring the

physiological condition and the effect of stress in crustaceans. Aquaculture 191, 209–245.

doi:10.1016/S0044-8486(00)00429-4




Luoma, S.N., Rainbow, P.S., 2005. Why Is Metal Bioaccumulation So Variable? Biodynamics as a Unifying

Concept. Environ. Sci. Technol. 39, 1921–1931. doi:10.1021/es048947e

Mackevica, A., Skjolding, L.M., Gergs, A., Palmqvist, A., Baun, A., 2015. Chronic toxicity of silver nanoparticles

to Daphnia magna under different feeding conditions. Aquat. Toxicol. 161, 10–16.

doi:10.1016/j.aquatox.2015.01.023



194

(crustacea:amphipoda): Problems and Perspectives Concerning the Functional Feeding Group Concept. Biol.

Rev. 72, 349–364. doi:10.1111/j.1469-185X.1997.tb00017.x

Mann, R.M., Grosell, M., Bianchini, A., Wood, C.M., 2004. Biologically incorporated dietary silver has no

ionoregulatory effects in American red crayfish (Procambarus clarkii). Environ. Toxicol. Chem. 23, 388–395.

doi:10.1897/02-572

Mantel, L.H., Farmer, L.L., 1983. 2 - Osmotic and Ionic Regulation, in: Linda Mantel (Ed.), Internal Anatomy

and Physiological Regulation. Academic Press, pp. 53–161.

McGillicuddy, E., Murray, I., Kavanagh, S., Morrison, L., Fogarty, A., Cormican, M., Dockery, P., Prendergast,

M., Rowan, N., Morris, D., 2017. Silver nanoparticles in the environment: Sources, detection and

ecotoxicology. Sci. Total Environ. 575, 231–246. doi:10.1016/j.scitotenv.2016.10.041

Mehennaoui, K., Georgantzopoulou, A., Felten, V., Andreï, J., Garaud, M., Cambier, S., Serchi, T., Pain-Devin,

S., Guérold, F., Audinot, J.-N., Giambérini, L., Gutleb, A.C., 2016. Gammarus fossarum (Crustacea, Amphipoda)

as a model organism to study the effects of silver nanoparticles. Sci. Total Environ.

doi:10.1016/j.scitotenv.2016.06.068

Naddy, R.B., McNerney, G.R., Gorsuch, J.W., Bell, R.A., Kramer, J.R., Wu, K.B., Paquin, P.R., 2011. The effect

of food on the acute toxicity of silver nitrate to four freshwater test species and acute-to-chronic ratios.

Ecotoxicology 20, 2019. doi:10.1007/s10646-011-0745-7

Nair, P.M.G., Park, S.Y., Choi, J., 2013. Evaluation of the effect of silver nanoparticles and silver ions using

stress responsive gene expression in Chironomus riparius. Chemosphere 92, 592–599.


Nel, A., Xia, T., Mädler, L., Li, N., 2006. Toxic Potential of Materials at the Nanolevel. Science 311, 622–627.

doi:10.1126/science.1114397

Oberdörster, E., Zhu, S., Blickley, T.M., McClellan-Green, P., Haasch, M.L., 2006. Ecotoxicology of carbon-

based engineered nanoparticles: Effects of fullerene (C60) on aquatic organisms. Carbon 44, 1112–1120.

doi:10.1016/j.carbon.2005.11.008

Pan, J.-F., Buffet, P.-E., Poirier, L., Amiard-Triquet, C., Gilliland, D., Joubert, Y., Pilet, P., Guibbolini, M., Risso

de Faverney, C., Roméo, M., Valsami-Jones, E., Mouneyrac, C., 2012. Size dependent bioaccumulation and

ecotoxicity of gold nanoparticles in an endobenthic invertebrate: The Tellinid clam Scrobicularia plana.


Petersen, E.J., Henry, T.B., Zhao, J., MacCuspie, R.I., Kirschling, T.L., Dobrovolskaia, M.A., Hackley, V., Xing, B.,

White, J.C., 2014. Identification and Avoidance of Potential Artifacts and Misinterpretations in Nanomaterial

Ecotoxicity Measurements. Environ. Sci. Technol. 48, 4226–4246. doi:10.1021/es4052999




Rosas, C., Sanchez, A., Escobar, E., Soto, L., Bolongaro-Crevenna, A., 1992. Daily variations of oxygen

consumption and glucose hemolymph level related to morphophysiological and ecological adaptations of

crustacea. Comp. Biochem. Physiol. A Physiol. 101, 323–328. doi:10.1016/0300-9629(92)90540-7


195

Scott, G.R., Sloman, K.A., 2004. The effects of environmental pollutants on complex fish behaviour:

integrating behavioural and physiological indicators of toxicity. Aquat. Toxicol. 68, 369–392.

doi:10.1016/j.aquatox.2004.03.016



doi:10.1016/j.aquatox.2016.10.004

Sornom, P., Felten, V., Médoc, V., Sroda, S., Rousselle, P., Beisel, J.-N., 2010. Effect of gender on physiological

and behavioural responses of Gammarus roeseli (Crustacea Amphipoda) to salinity and temperature. Environ.


Stevenson, L.M., Dickson, H., Klanjscek, T., Keller, A.A., McCauley, E., Nisbet, R.M., 2013. Environmental

Feedbacks and Engineered Nanoparticles: Mitigation of Silver Nanoparticle Toxicity to Chlamydomonas

reinhardtii by Algal-Produced Organic Compounds. PLoS ONE 8. doi:10.1371/journal.pone.0074456

Stevenson, L.M., Krattenmaker, K.E., Johnson, E., Bowers, A.J., Adeleye, A.S., McCauley, E., Nisbet, R.M., 2017.

Standardized toxicity testing may underestimate ecotoxicity: Environmentally relevant food rations increase

the toxicity of silver nanoparticles to Daphnia. Environ. Toxicol. Chem. n/a-n/a. doi:10.1002/etc.3869

Tiede, K., Hassellöv, M., Breitbarth, E., Chaudhry, Q., Boxall, A.B.A., 2009. Considerations for environmental

fate and ecotoxicity testing to support environmental risk assessments for engineered nanoparticles. J.

Chromatogr. A, Tools for the REACH Programme - analytical methods for the evaluation of industrial

contaminants 1216, 503–509. doi:10.1016/j.chroma.2008.09.008

Vale, G., Mehennaoui, K., Cambier, S., Libralato, G., Jomini, S., Domingos, R.F., 2016. Manufactured

nanoparticles in the aquatic environment-biochemical responses on freshwater organisms: A critical

overview. Aquat. Toxicol. 170, 162–174. doi:10.1016/j.aquatox.2015.11.019

Vance, M.E., Kuiken, T., Vejerano, E.P., McGinnis, S.P., Hochella, M.F., Rejeski, D., Hull, M.S., 2015.

Nanotechnology in the real world: Redeveloping the nanomaterial consumer products inventory. Beilstein J.

Nanotechnol. 6, 1769–1780. doi:10.3762/bjnano.6.181

Vellinger, C., Gismondi, E., Felten, V., Rousselle, P., Mehennaoui, K., Parant, M., Usseglio-Polatera, P., 2013.

Single and combined effects of cadmium and arsenate in Gammarus pulex (Crustacea, Amphipoda):

Understanding the links between physiological and behavioural responses. Aquat. Toxicol. 140–141, 106–

116. doi:10.1016/j.aquatox.2013.05.010

Volland, M., Hampel, M., Martos-Sitcha, J.A., Trombini, C., Martínez-Rodríguez, G., Blasco, J., 2015. Citrate

gold nanoparticle exposure in the marine bivalve Ruditapes philippinarum: uptake, elimination and oxidative

stress response. Environ. Sci. Pollut. Res. 22, 17414–17424. doi:10.1007/s11356-015-4718-x

Webb, N.A., Wood, C.M., 1998. Physiological analysis of the stress response associated with acute silver

nitrate exposure in freshwater rainbow trout (Oncorhynchus mykiss). Environ. Toxicol. Chem. 17, 579–588.

doi:10.1002/etc.5620170408

Wray, A.T., Klaine, S.J., 2015. Modeling the influence of physicochemical properties on gold nanoparticle

uptake and elimination by Daphnia magna. Environ. Toxicol. Chem. 34, 860–872. doi:10.1002/etc.2881

Yang, Y., Long, C.-L., Li, H.-P., Wang, Q., Yang, Z.-G., 2016. Analysis of silver and gold nanoparticles in

environmental water using single particle-inductively coupled plasma-mass spectrometry. Sci. Total Environ.

Chapter 6 General discussion

198

CHAPTER 6

General discussion

CHAPTER 6

General discussion


199

Amphipods are one of the most important components of freshwater ecosystems. Among them, gammarids

are the most widespread group in Europe and are often used as bioindicators of the quality of aquatic

ecosystems and model organisms in many ecotoxicological studies due to their well-known ecology and

sensitivity to different kind of pollutants (Farkas et al., 2003; Kunz et al., 2010). However, their use, especially

of Gammarus fossarum for the study of the environmental impact of nanoparticles has been rather limited

so far (Mehennaoui et al., 2016).

Engineered nanoparticles (ENPs) are being increasingly incorporated in consumer products and as a

consequence, their environmental release is raising concerns about their potential toxicity/effects on living

organisms (Klaine et al., 2008; Moreno-Garrido et al., 2015). Silver nanoparticles (AgNPs) are among the most

promising and used ENPs for application in numerous consumer products due to their well-documented

antibacterial properties (Bone et al., 2012; Cleveland et al., 2012; Vance et al., 2015). Gold nanoparticles

(AuNPs) are used in a wide range of applications in the field of biomedicine, biology and chemistry (Nel et al.,

2006; Volland et al., 2015) mostly for their low toxicity and high bio-compatibility (García-Cambero et al.,

2013; Lapresta-Fernández et al., 2012).

The research conducted within this thesis aimed at the evaluation of potential threats of AgNPs and AuNPs

and identification of their key mechanisms of toxicity on G. fossarum. The influence of size, surface coating

and chemical composition on the toxicity of AgNPs and AuNPs was assessed. An integrated approach

including molecular, enzymatic, physiological and behavioural responses, was applied in order to define

and potentially predict effects of AgNPs and AuNPs at higher biological organisation (population,

community, ecosystems) regarding the central and important position of G. fossarum in the aquatic

ecosystems.

Regarding the lack of information on the toxicity mechanisms of ENPs on G. fossarum and its scare use as a

model organism in nanotoxicology studies, the first study of this thesis (Chapter 2) aimed at evaluating the

effects of a well-characterised and well-studied set of AgNPs (Georgantzopoulou, 2015). Synthetically

produced AgNPs of different sizes (20 and 200 nm) as well as AgNPs synthetized by a biological method (using

plant leaf extracts of Ocinum sanctum and Azadirachta indica, AgNPs 23 and 27 nm, respectively;

(Balachandran et al., 2012, 2013) were used in order to elucidate the relation between size, synthesis

method, NPs surface properties, ions dissolution and their toxicity (Georgantzopoulou et al., 2013;

Mehennaoui et al., 2016). A multi-biomarker approach was applied in order to determine the effects at the

physiological and behavioural level. Biological endpoints including survival, bioaccumulation, tissue

distribution, osmoregulation, antioxidant responses, defence mechanisms, cellular damage, energy reserves

and behaviour were investigated and allowed obtaining first information about the effects of AgNPs on G.

fossarum (Figure 6. 1).


200

Figure 6. 1. Physiological and behavioural effects of synthetic and biological AgNPs on G. fossarum exposed for

72h: summary of results presented in Chapter 2. G. fossarum were exposed for 72h to synthetic AgNPs 20 and

200 nm and biologically synthetized from plant leaf extract AgNPs 23 and 27 nm. Effects on antioxidant responses

(GPx, TAC, CAT), defense mechanism (GST, ACP), cellular damage (LDH, CASP3, LOOH), energy reserves (Prot, Chol,

Trig, ETS), osmoregulation and behaviour (Locomotion and ventilation) were assessed.

Despite the rapid advances in the “omics” field marked by the development of next generation sequencing

(NGS) methods, RT-qPCR remains one of the most accurate and reliable techniques for target gene expression

and NGS data validation. This method is recognized for its capacity to highlight sensitive changes in gene

transcription levels (Guerriero et al., 2014; Plusquin et al., 2012) which represent an early warning of

pollution-induced stress and may be linked to important physiological changes (Lacroix et al., 2014).

Therefore, a set of stress related genes, selected based on the battery of biomarkers developed earlier by

our research group (Garaud et al., 2015a; 2015b, 2016), was used to investigate the toxicity/effects of

different sized AgNPs and AuNPs (20, 40 and 80 nm) with two different coatings (CIT and PEG) on G. fossarum

(Chapter 4). This study aimed, firstly, at investigating the influence of the size, chemical composition and

surface coating on the uptake, tissue distribution and molecular effects of AgNPs and AuNPs on G. fossarum

exposed for 72h to AgNPs and AuNPs, in the absence of food (Figure 6. 2). Additionally, in order to obtain

accurate results at the molecular level, a set of reference genes for RT-qPCR data normalization was

identified. To our best knowledge, this study was the first attempt of identifying a set of reference genes for

Sub-individual

Physiologicaleffects

• ↓Osmoregulation

• Antioxidantresponses

• Defensemechanisms

• Cellulardamage

• Energyreserves

Individual

Behavioraleffects

• ↓ Locomotion

• Ventilation

• ↓Survival

SyntheticAgNPs Biological AgNPs

20 200 23 27


201

the freshwater crustacean G. fossarum. Six candidate genes (actin, TUB, UB, SDH, clathrin and GAPDH) were

tested in order to determine the most stable ones in different stress conditions and to increase the

robustness of obtained RT-qPCR data (Chapter 3).

Figure 6. 2. Influence of size (20, 40 and 80 nm) and surface coating (CIT and PEG) of AgNPs and AuNPs on their

uptake, tissue distribution and molecular effects of G. fossarum exposed for 72h: summary of the results

presented in Chapter 4. G. fossarum were exposed for 72h to up to 50 µg. L-1 of AgNPs and AuNPs in absence of

food. Influence of size and surface coating on bioaccumulation was assessed using ICP-MS, internal distribution of

AgNPs and AuNPs were evaluated using NanoSIMS50 and Cytoviva, and molecular effects were assessed using RT-

qPCR. A set of stress-related genes expression including genes implied in cytoskeleton trafficking (Actin, TUB, UB),

exoskeleton cuticle (Chitinase), antioxidant defence (CAT, MnSOD, CuZnSOD, GPx7), general stress (GST, HSP90,

GAPDH), DNA damage and repair (Gadd45, NfkB, c-jun, P53), lysosomes (Cathepsin L), osmoregulation and

respiration (Na+K+ATPase, HEM) was used.

The second aim of this approach was the identification of sub-chronic effects of AgNPs and AuNPs on G.

fossarum. Therefore, a more realistic exposure scenario was applied by exposing G. fossarum for 15 days

to CIT and PEG-AgNPs 40 nm and CIT and PEG-AuNPs 40 nm in presence of food. Effects on the molecular

(gene expression), physiological (osmoregulation) and behavioural responses (locomotion and ventilation)

were investigated (Chapter 5, Figure 6. 3).

AgNPs

CIT PEG

AuNPs

CIT PEG

20

40

80

Bioaccumulation

• CIT-AuNPs>PEG-AuNPs

• Concentration

dependentuptake

• Nosizeeffect

TissueDistribution

• ↑↑Intestine

• ↑Cuticle

• Gills

Moleculareffects

• ↑GPx7

• ↓MnSOD

• ↓CuZnSOD

• ↓HSP90

• ↓Hemocyanin

Bioaccumulation

• CIT-AgNPs>PEG-AgNPs

• Concentration

dependentuptake

• Nosizeeffect

TissueDistribution

• ↑↑Gills

• ↑Cuticle

• Intestine

Moleculareffects

• ↑MnSOD

• ↑CuZnSOD

• ↑Chitinase

• ↓GST

• ↓Hemocyanin

• ↓CathepsinL

20

40

80

20

40

80

20

40

80


202

Figure 6. 3. Sub-chronic toxicity of CIT and PEG-AgNPs and CIT and PEG-AuNPs 40 nm on molecular, physiological

and behavioural responses of G. fossarum: summary of the results presented in Chapter 5. G. fossarum were

exposed for 15 days to 0.5 and 5 µg. L-1 of CIT and PEG-AgNPs and AuNPs 40 nm in presence of food.

Bioaccumulation was assessed using ICP-MS, internal distribution of AgNPs and AuNPs were evaluated using

Cytoviva and molecular effects were assessed using a set of stress-related genes expression including genes

implied in cytoskeleton trafficking (Actin, TUB, UB), exoskeleton cuticle (Chitinase), antioxidant defence (CAT,

MnSOD, CuZnSOD, GPx7), general stress (GST, HSP90, GAPDH), DNA damage and repair (Gadd45, NfkB, c-jun, P53),

lysosomes (Cathepsin L), osmoregulation and respiration (Na+K+ATPase, HEM).

AgNPs

CIT PEG

AuNPs

CIT PEG

Molecularresponses

• ↑CuZnSOD

Physiologicalresponses

• [Na+]

• [Cl-]

• [Ca2+]

Behavioralresponses

• ↓ Locomotion

• Ventilation

Molecularresponses

• ↑CAT

• ↑Chitinase

• ↑Na+K+ATPase

Physiologicalresponses

• [Na+]

• [Cl-]

• [Ca2+]

Behavioralresponses

• ↓ Locomotion

• Ventilation40 40

40 40


203

1. NPS SIZE, SURFACE COATING AND CHEMICAL COMPOSITION INFLUENCE THE

UPTAKE

As reported in many different studies, NPs size, surface coating and chemical composition appeared as critical

factors controlling the uptake and effects of AgNPs and AuNPs (Baumann et al., 2014; Blinova et al., 2013;

Ivask et al., 2014; Silva et al., 2014; Chapter 2 and 4). A size-dependent toxicity was observed on G. fossarum

exposed for 72h to AgNPs 20, 23, 27 and 200 nm with AgNPs 23 nm, as the one that aggregated the less

(smallest hydrodynamic diameter of 53 nm), being the most toxic one with an LC50 of at least 2.3 µg. L-1

(Figure 6. 1). The order of toxicity was the same as reported in a previous study using V. fisheri, D. subspicatus

and D. magna as model organisms (Georgantzopoulou et al., 2013). Size-dependent effects for the bare

AgNPs 20 and 200 nm with AgNPs 20 nm being more potent to D. magna than AgNPs 200 nm were reported.

Moreover, biologically synthetized AgNPs 23 and 27 nm were always more toxic to D. magna with AgNPs 23

nm being the most toxic one (Georgantzopoulou et al., 2013). Similarly, in addition to dose-dependent

effects of AgNPs on a multi-trophic test battery including algae and D. magna, size-dependent effects were

observed with the smallest AgNPs (10 nm) being more toxic to D. magna than the bigger ones (20, 40, 60 and

80 nm) (Ivask et al., 2014).

Interestingly, using commercial AgNPs, stabilized with citrate or coated with PEG, did not induce size-

dependent effects. Indeed, short-term exposure of G. fossarum to three different sizes of CIT and PEG-AgNPs,

namely 20, 40 and 80 nm, led to a concentration-dependent rather than size-dependent Ag uptake (Chapter

4, Figure 6. 2). The same results were obtained for AuNPs with a concentration-dependent rather than a size-

dependent Au uptake. A surface coating-dependent bioaccumulation of Ag and Au was observed with CIT-

AgNPs and CIT-AuNPs being more taken up by G. fossarum than PEG-AgNPs and PEG-AuNPs. Our results were

consistent with previous studies that reported a coating dependent bioaccumulation of CIT-AgNPs and CIT-

AuNPs in aquatic organisms (Oliver et al., 2014; Park et al., 2015; Sakka et al., 2016). These differences may

be linked to the behaviour of CIT-NPs in the water column. As they appeared to aggregate more than PEG-

NPs, it could be expected that CIT-NPs settle down and reach the water/sediment interface where G.

fossarum tend to be found. Therefore, it could be expected that CIT-NPs may be more bioavailable for benthic

macro-invertebrates than PEG-NPs. It has been reported that G. roeseli accumulated more cerium (Ce) than

Dreissena polymorpha exposed to CeO-NPs (Garaud et al., 2015). The higher bioaccumulation could be linked

to the ingestion of aggregate that may settle in the aquarium or via the ingestion of faeces and pseudo-faeces

of contaminated bivalves by gammarids (Garaud et al., 2015). However, in chapter 2, we showed that the

bare AgNPs 20 and 200 nm were less absorbed by G. fossarum than biological AgNPs 23 and 27 nm despite a

higher aggregation (Mehennaoui et al., 2016). This result enhances the key role played by surface coating of

AgNPs and AuNPs bioaccumulation by G. fossarum. Exposing G. fossarum to CIT and PEG-AgNPs 40 nm and


204

CIT and PEG-AuNPs 40 nm for 15 days and in the presence of food confirmed the role of surface coating on

the bioaccumulation of Ag and Au (Chapter 5, Figure 6. 3). Indeed, G. fossarum exposed to the highest

concentration of CIT-AgNPs and CIT-AuNPs 40 nm (5 µg. L-1) tended to accumulate more Ag and Au than

animals exposed to PEG-AgNPs and PEG-AuNPs.

One important step to understand NPs’ fate, behaviour and adverse effects is by investigating their internal

distribution in tissues. Usually, transmission electron microscopy (TEM) is used for internal localization of NPs

(Heinlaan et al., 2008; Lovern et al., 2008). However, this method does not allow a direct elemental

identification of NPs leading to ambiguous results (Georgantzopoulou et al., 2013). In the beginning of the

present thesis, we have used one of the latest generation of Secondary Ion Mass Spectrometry instruments,

the NanoSIMS 50. This probe provides information about the elemental distribution of NPs within biological

materials. However, it does not allow the distinction between ions and NPs. Later during this work, a dark-

field hyperspectral microscope, Cytoviva, was acquired, which allows detection of only the NPs in biological

tissues but not of the ions. In Chapter 2, NanoSIMS allowed localization of Ag from AgNPs 23 and 27 nm in

the gills of G. fossarum. The other organs could not be analysed due to sample preparation limits. Thus, the

protocol of sample preparation for NanoSIMS 50 analyses was optimized and different tissues of G. fossarum

were analysed, namely gills, intestines, muscle and external cuticle (Chapter 4, Figure 6. 2), when Gammarus

were exposed to CIT and PEG AgNPs and AuNPs. NanoSIMS50 and Cytoviva allowed the confirmation of

bioaccumulation results from ICP-MS as the signal of Ag and Au obtained in G. fossarum exposed to CIT-

AgNPs and CIT-AuNPs was higher than PEG-AgNPs and PEG-AuNPs.

Interestingly, the chemical composition of the studied NPs appeared also as a critical factor determining their

internal distribution. Indeed, different uptake routes were observed (Chapter 2, 4 and 5). AgNPs were found

in the gills (cuticle and epithelial tissue) while AuNPs were observed in the epithelial tissue of intestinal caeca

(Chapter 4, Figure 6. 2). The sub-chronic exposure of G. fossarum to CIT and PEG-AgNPs and CIT and PEG-

AgNPs 40 nm in the presence of food confirmed the different uptake routes as AgNPs were observed in the

gills while AuNPs were observed in the gut area (Figure 6. 3). However, the presence of the food led to the

localization of AuNPs within the lumen on intestinal caeca indicating a potential elimination with faeces while

in the absence of the food, it seems that AuNPs were able to cross the gut barrier.


205

2. SYNTHESIS METHODS AND CHEMICAL COMPOSITION INFLUENCE AGNPS AND

AUNPS EFFECTS

The NPs synthesis method proved to be an important parameter in determining their toxicity. The biological

AgNPs 23 and 27 nm synthesised using two different plant leaf extracts, A. indica (AgNPs 27 nm) and O.

sanctum (AgNPs 23 nm) were the most potent ones (Chapter 2, Figure 6. 1). Using two different plant leaf

extracts led to the presence of different biomolecules on the surface of AgNPs 23 and 27 nm as one additional

molecule was observed on the surface of AgNPs 27 which may confer it more stability compared to AgNPs 23

nm (Georgantzopoulou et al., 2013; Mehennaoui et al., 2016). These observations confirm once more the

importance of surface coating on bioaccumulation potential of AgNPs. Even though both AgNPs 23 and 27

nm led to a significant mortality of G. fossarum, the sub-lethal effects were different. Only AgNPs 23 nm led

to a higher mortality rate, osmoregulation impairment and a decrease in locomotion (Mehennaoui et al.,

2016). In the same way, the chemical composition played an important role on the sub-individual effects of

AgNPs and AuNPs on G. fossarum (Chapter 4 and 5). The acute exposure of G. fossarum to AgNPs led to an

up-regulation of MnSOD, CuZnSOD and Chitinase, and a down-regulation of GST, HEM and Cathepsin L gene

expression whereas AuNPs led to an up-regulation of GPx 7, and a decrease in MnSOD, CuZnSOD, HSP90 and

HEM gene expression (Chapter 4, Figure 6. 2). It is important to note that these effects were independent

from AgNPs and AuNPs size and surface coatings.

Gene expression appear as a very sensitive biomarker level. Although the effects at the protein and enzyme

activity level were not performed in the experiments described in Chapter 4 and 5, the observed effects at

the gene level allow to highlight that AgNPs and AuNPs induced different responses. The analysis of the gene

expression in G. fossarum also revealed the importance of the exposure route. For instance, acute exposure

of G. fossarum to AgNPs led to the induction of SODs while AuNPs led to their down-regulation. The chronic

exposure led to the up-regulation of Na+K+ATPase by AgNPs whereas no effects were observed on this gene

when G. fossarum were exposed to AuNPs. However, it can be suggested that both AgNPs and AuNPs might

activate antioxidant defense, which is coherent with literature as oxidative stress via ROS generation was

reported as the main mechanism of action of NPs (Vale et al., 2016).


206

3. CONTRIBUTION OF SOLUBLE IONS TO THE EFFECTS

In the present work, AgNO3 was used as a positive control in order to compare AgNPs effects to Ag ions effects

while ion release from AuNPs was not expected. Ions release from AgNPs is size, surface coating and exposure

media dependent (Georgantzopoulou, 2015). In the present thesis, significant ions release was observed only

with biologically synthetized AgNPs 23 nm (Chapter 2) with a dissolution rate of up to 29% in Volvic water

(Mehennaoui et al., 2016). Bare AgNPs 20 and 200 nm, AgNPs 27 nm, CIT and PEG-AgNPs and CIT and PEG-

AuNPs showed a dissolution rate of maximum 6% (24h) with very low concentration of Ag and Au ions

(chapter 2 and 4). According to the results obtained in Chapter 2 and 4, it seems that surface coating is the

main factor influencing ions release as bare, CIT and PEG-AgNPs in addition to AgNPs 27 nm appeared to be

more stable in the exposure medium (Volvic water) than AgNPs 23 nm.

Interestingly, AgNPs 23 nm were more toxic to G. fossarum than AgNPs 20 and 27 nm. Indeed, the EC50s

causing effects were similar for AgNPs 23 nm and AgNO3. Both of them led to a decrease in survival rates and

locomotion in addition to an osmoregulation impairment at similar concentrations (Mehennaoui et al., 2016).

These results highlight the implication of Ag ions in the potency of AgNPs to G. fossarum. However, ion

release seems to be not the only mechanism of effects of AgNPs on G. fossarum. Indeed, gene expression

results showed that AgNPs affect G. fossarum in a different way compared to AgNO3 (chapter 4 and 5)

although common mechanisms were reported. For instance, both AgNPs and AgNO3 led to a down-regulation

of hemocyanin and an up-regulation of CuZnSOD and Chitinase whereas only AgNPs increased MnSOD gene

expression and decreased GST, Cathepsin L and AgNO3 led to an up-regulation of TUB, GST and Cathepsin L

and to a down-regulation of NfkB. These results are in accordance with results observed on the MXR

responses of in vitro gastrointestinal co-culture model exposed to AgNPs (Georgantzopoulou et al., 2016).

AgNPs 23 nm were more potent than AgNO3 and AgNPs 27 nm while the level of dissolved Ag in the media

were comparable for AgNO3 and AgNPs 23 nm (Georgantzopoulou et al., 2016). This suggest that the

differences in the observed effects are not only linked to Ag ions release as AgNPs appeared to be sometime

more toxic than AgNO3 at the same nominal exposure concentrations (García-Alonso et al., 2014; Ivask et al.,

2014). Moreover, different gene expression profiles were observed in D. magna exposed to AgNO3 and AgNPs

(Poynton et al., 2012) which is in support of our findings for the gene expression study (Chapter 4).


207

4. LIMITS OF THE MULTI-BIOMARKERS APPROACH

The application of a multi-biomarkers approach allowed the assessment of effects of AgNPs and AuNPs on

different biological levels (from molecular to behavioural responses). Biomarkers need to be sensitive

indicators and are defined as early warning of pollution-induced stress on ecosystems (Lagadic, 2002; Vasseur

and Leguille, 2004). The results obtained in the present work allowed to confirm the sensitivity of sub-

individual and individual responses (behaviour) as effects were observed at very low and environmentally

realistic concentrations of AgNPs and AuNPs (from 0.5 µg. L-1). Behavioural responses allow obtaining rapid

and non-invasive responses even though no effects on ventilation could be detected in the present work

(Chapter 2 and 5). However, locomotion was one of the most sensitive responses as effects were detected at

very low concentrations of AgNPs and AuNPs (0.5 µg. L-1). The effects on locomotion may suggest an energy

reallocation strategy in favour of defense mechanisms in G. fossarum. Our results were in accordance with

previous studies performed within the ANR MESONNET project by exposing G. roeseli to different sizes of

CIT-AgNPs (Andreï et al., 2016). Effects on locomotion may also allow predicting the effects of AgNPs and

AuNPs at the population and community level. Indeed, locomotion is a highly ecologically relevant biomarker

as it is necessary for foraging, mate finding and escaping from predators. By interfering with locomotion

AgNPs and AuNPs may reduce the fitness of the organism and have a potential effect at the population level.

The use of numerous stress-related genes allowed to highlight the influence of chemical composition on the

effects of AgNPs and AuNPs on G. fossarum. However, the obtained results should be considered with

precaution as one of the difficulties of the target genes approach is the important number of potential

isoforms present encoding for each protein family. As no functional studies (proteomic analyses, enzymatic

analyses; Chapter 4 and 5) were performed, no clear conclusions can be formulated as an induction of a given

gene does not systematically result in the expression/activation at the protein level. Hence, further analyses

are still needed in order to better define and predict the effects of AgNPs and AuNPs on G. fossarum and to

better link responses observed at the tissue level to those observed at the whole individual level.

Furthermore, the responses studied in chapter 2 seem not to be the most appropriate endpoints for studying

the effects of NPs on G. fossarum. The absence of significant effects may be due to exposure time and the

chemical composition of the studied materials. Indeed, it has been reported that antioxidant responses were

significantly impacted in goldfish after 6h of exposure to a mixture of pesticides while no significant effects

on antioxidant responses were observed after 96h or 16 days of exposure (Gandar et al., 2017). This may

suggest the setting up of different sampling times in order to unsure detection of early effects on G. fossarum.


208

5. CONCLUDING REMARKS AND FUTURE PERSPECTIVES

The present work provides a substantial amount of data which open new windows for further exploratory

work. The results of this thesis highlight the importance of considering on one hand the NPs characteristics

such as size, methods of synthesis, surface coating and chemical composition and on the other hand, the

exposure mode of the studied AgNPs and AuNPs regarding their significant influence on the behaviour of NPs

in the exposure medium and uptake routes.

It can be concluded that the chemical composition is the main factor influencing the effects of NPs on aquatic

organisms. It is important to keep in mind that for a specific NP, physico-chemical characteristics should be

considered as these parameters play a crucial role in modulating the behaviour of NPs in the environment

and their interactions with organisms. The results of the present thesis show that both the NPs and the

released ions play a role for the toxicity to G. fossarum. It seems that surface coating and chemical

composition, and to a lower extent size, are the main factors favouring uptake, strong interaction with

biological sites and thus relative toxicity of AgNPs and AuNPs.

The knowledge acquired in the present work through the exposure of G. fossarum to realistic concentrations

of NPs contributed to a better understanding of mechanisms of action of AgNPs and AuNPs. However,

establishment of more realistic exposure scenarios including river water and organic matter should provide

complementary information with a biological continuum linking molecular to functional responses as these

latter appeared as sensitive responses (Andreï et al., 2016). Linking molecular to individual and functional

responses will allow a better prediction of the effects of NPs at the population and community level. With

the acquirement of new sequences of G. fossarum, it might be valuable to apply NGS methods like RNAseq

(Illumina) in order to identify new biomarkers in nano-ecotoxicology. This highly sensitive method might

allow a better coverage and determination of mechanisms of action of NPs. The use of SIMS in combination

with Cytoviva allows localisation of AgNPs and AuNPs within G. fossarum. However, these techniques do not

allow a determination of effects of the studied NPs on the studied tissues. Hence, the use of histopathological

methods might also bring complementary data to better link responses at the molecular level to those

observed at the individual level.


209

References Andreï, J., Pain-Devin, S., Felten, V., Devin, S., Giambérini, L., Mehennaoui, K., Cambier, S., Gutleb, A.C., Guérold,

F., 2016. Silver nanoparticles impact the functional role of Gammarus roeseli (Crustacea Amphipoda). Environ. Pollut. 208, Part B, 608–618. doi:10.1016/j.envpol.2015.10.036

Balachandran, Y.L., Girija, S., Selvakumar, R., Tongpim, S., Gutleb, A.C., Suriyanarayanan, S., 2013. Differently Environment Stable Bio-Silver Nanoparticles: Study on Their Optical Enhancing and Antibacterial Properties. PLoS ONE 8, e77043. doi:10.1371/journal.pone.0077043

Balachandran, Y.L., Peranantham, P., Selvakumar, R., Gutleb, A.C., Girija, S., 2012. Size-Controlled Green Synthesis of Silver Nanoparticles using Dual Functional Plant Leaf Extract at Room Temperature. Int. J. Green Nanotechnol. 4, 310–325. doi:10.1080/19430892.2012.706183

Baumann, J., Köser, J., Arndt, D., Filser, J., 2014. The coating makes the difference: Acute effects of iron oxide nanoparticles on Daphnia magna. Sci. Total Environ. 484, 176–184. doi:10.1016/j.scitotenv.2014.03.023

Blinova, I., Niskanen, J., Kajankari, P., Kanarbik, L., Käkinen, A., Tenhu, H., Penttinen, O.-P., Kahru, A., 2013. Toxicity of two types of silver nanoparticles to aquatic crustaceans Daphnia magna and Thamnocephalus platyurus. Environ. Sci. Pollut. Res. 20, 3456–3463. doi:10.1007/s11356-012-1290-5

Bone, A.J., Colman, B.P., Gondikas, A.P., Newton, K.M., Harrold, K.H., Cory, R.M., Unrine, J.M., Klaine, S.J., Matson, C.W., Di Giulio, R.T., 2012. Biotic and Abiotic Interactions in Aquatic Microcosms Determine Fate and Toxicity of Ag Nanoparticles: Part 2–Toxicity and Ag Speciation. Environ. Sci. Technol. 46, 6925–6933. doi:10.1021/es204683m

Cleveland, D., Long, S.E., Pennington, P.L., Cooper, E., Fulton, M.H., Scott, G.I., Brewer, T., Davis, J., Petersen, E.J., Wood, L., 2012. Pilot estuarine mesocosm study on the environmental fate of Silver nanomaterials leached from consumer products. Sci. Total Environ., Special Section: Reviews of Trace Metal Pollution in China 421–422, 267–272. doi:10.1016/j.scitotenv.2012.01.025

Farkas, A., Salánki, J., Varanka, I., 2003. Crustaceans as biological indicators of heavy metal pollution in Lake Balaton (Hungary). Hydrobiologia 506–509, 359–364. doi:10.1023/B:HYDR.0000008615.86330.1d

Gandar, A., Laffaille, P., Canlet, C., Tremblay, M., Gautier, R., Perrault, A., Gress, L., Mormède, P., Tapie, N., Budzinski, H., Jean, S., 2017. Adaptive response under multiple stress exposure in fish: From the molecular to individual level. Chemosphere 188. doi:10.1016/j.chemosphere.2017.08.089

Garaud, M., Auffan, M., Devin, S., Felten, V., Pagnout, C., Pain-Devin, S., Proux, O., Rodius, F., Sohm, B., Giamberini, L., 2016. Integrated assessment of ceria nanoparticle impacts on the freshwater bivalve Dreissena polymorpha. Nanotoxicology 10, 935–944. doi:10.3109/17435390.2016.1146363

Garaud, M., Auffan, M., Devin, S., Felten, V., Pagnout, C., Pain-Devin, S., Proux, O., Rodius, F., Sohm, B., Giambérini, L., 2015. Fate and integrated assessment of ceria nanoparticle impacts on the freshwater bivalve Dreissena polymorpha: a mesocosm approach. Nanotoxicology.

Garaud, Trapp, J., Devin, S., Cossu-Leguille, C., Pain-Devin, S., Felten, V., Giamberini, L., 2015. Multibiomarker assessment of cerium dioxide nanoparticle (nCeO2) sublethal effects on two freshwater invertebrates, Dreissena polymorpha and Gammarus roeseli. Aquat. Toxicol. 158, 63–74. doi:10.1016/j.aquatox.2014.11.004

García-Alonso, J., Rodriguez-Sanchez, N., Misra, S.K., Valsami-Jones, E., Croteau, M.-N., Luoma, S.N., Rainbow, P.S., 2014. Toxicity and accumulation of silver nanoparticles during development of the marine polychaete Platynereis dumerilii. Sci. Total Environ. 476–477, 688–695. doi:10.1016/j.scitotenv.2014.01.039

García-Cambero, J.P., Núñez García, M., López, G.D., Herranz, A.L., Cuevas, L., Pérez-Pastrana, E., Cuadal, J.S., Castelltort, M.R., Calvo, A.C., 2013. Converging hazard assessment of gold nanoparticles to aquatic organisms. Chemosphere 93, 1194–1200. doi:10.1016/j.chemosphere.2013.06.074

Georgantzopoulou, A., 2015. Effects of silver nanoparticles and ions and interaction with first line of defense. Wageningen University, Wageningen Netherlands.


210

Georgantzopoulou, A., Balachandran, Y.L., Rosenkranz, P., Dusinska, M., Lankoff, A., Wojewodzka, M., Kruszewski, M., Guignard, C., Audinot, J.-N., Girija, S., Hoffmann, L., Gutleb, A.C., 2013. Ag nanoparticles: size- and surface-dependent effects on model aquatic organisms and uptake evaluation with NanoSIMS. Nanotoxicology 1–11. doi:10.3109/17435390.2012.715312

Georgantzopoulou, A., Cambier, S., Serchi, T., Kruszewski, M., Balachandran, Y.L., Grysan, P., Audinot, J.-N., Ziebel, J., Guignard, C., Gutleb, A.C., Murk, A.J., 2016. Inhibition of multixenobiotic resistance transporters (MXR) by silver nanoparticles and ions in vitro and in Daphnia magna. Sci. Total Environ. 569–570, 681–689. doi:10.1016/j.scitotenv.2016.06.157

Guerriero, G., Legay, S., Hausman, J.-F., 2014. Alfalfa Cellulose Synthase Gene Expression under Abiotic Stress: A Hitchhiker’s Guide to RT-qPCR Normalization. PLOS ONE 9, e103808. doi:10.1371/journal.pone.0103808

Heinlaan, M., Ivask, A., Blinova, I., Dubourguier, H.-C., Kahru, A., 2008. Toxicity of nanosized and bulk ZnO, CuO and TiO2 to bacteria Vibrio fischeri and crustaceans Daphnia magna and Thamnocephalus platyurus. Chemosphere 71, 1308–1316. doi:10.1016/j.chemosphere.2007.11.047

Ivask, A., ElBadawy, A., Kaweeteerawat, C., Boren, D., Fischer, H., Ji, Z., Chang, C.H., Liu, R., Tolaymat, T., Telesca, D., Zink, J.I., Cohen, Y., Holden, P.A., Godwin, H.A., 2014. Toxicity Mechanisms in Escherichia coli Vary for Silver Nanoparticles and Differ from Ionic Silver. ACS Nano 8, 374–386. doi:10.1021/nn4044047

Klaine, S.J., Alvarez, P.J.J., Batley, G.E., Fernandes, T.F., Handy, R.D., Lyon, D.Y., Mahendra, S., McLaughlin, M.J., Lead, J.R., 2008. Nanomaterials in the environment: Behavior, fate, bioavailability, and effects. Environ. Toxicol. Chem. 27, 1825–1851. doi:10.1897/08-090.1

Kunz, P.Y., Kienle, C., Gerhardt, A., 2010. Gammarus spp. in Aquatic Ecotoxicology and Water Quality Assessment: Toward Integrated Multilevel Tests, in: Whitacre, D.M. (Ed.), Reviews of Environmental Contamination and Toxicology Volume 205, Reviews of Environmental Contamination and Toxicology. Springer New York, pp. 1–76.

Lacroix, C., Coquillé, V., Guyomarch, J., Auffret, M., Moraga, D., 2014. A selection of reference genes and early-warning mRNA biomarkers for environmental monitoring using Mytilus spp. as sentinel species. Mar. Pollut. Bull. 86, 304–313. doi:10.1016/j.marpolbul.2014.06.049

Lagadic, L., 2002. Biomarkers: Useful tools for the monitoring of aquatic environments. Rev. Med. Veterinaire 153, 581–588.

Lapresta-Fernández, A., Fernández, A., Blasco, J., 2012. Nanoecotoxicity effects of engineered silver and gold nanoparticles in aquatic organisms. TrAC Trends Anal. Chem. 32, 40–59. doi:10.1016/j.trac.2011.09.007

Lovern, S.B., Owen, H.A., Klaper, R., 2008. Electron microscopy of gold nanoparticle intake in the gut of Daphnia magna. Nanotoxicology 2, 43–48. doi:10.1080/17435390801935960

Mehennaoui, K., Georgantzopoulou, A., Felten, V., Andreï, J., Garaud, M., Cambier, S., Serchi, T., Pain-Devin, S., Guérold, F., Audinot, J.-N., Giambérini, L., Gutleb, A.C., 2016. Gammarus fossarum (Crustacea, Amphipoda) as a model organism to study the effects of silver nanoparticles. Sci. Total Environ. 566–567, 1649–1659. doi:10.1016/j.scitotenv.2016.06.068

Moreno-Garrido, I., Pérez, S., Blasco, J., 2015. Toxicity of silver and gold nanoparticles on marine microalgae. Mar. Environ. Res., Particles in the Oceans: Implication for a safe marine environment 111, 60–73. doi:10.1016/j.marenvres.2015.05.008

Nel, A., Xia, T., Mädler, L., Li, N., 2006. Toxic Potential of Materials at the Nanolevel. Science 311, 622–627. doi:10.1126/science.1114397

Oliver, A.L.-S., Croteau, M.-N., Stoiber, T.L., Tejamaya, M., Römer, I., Lead, J.R., Luoma, S.N., 2014. Does water chemistry affect the dietary uptake and toxicity of silver nanoparticles by the freshwater snail Lymnaea stagnalis? Environ. Pollut. 189, 87–91. doi:10.1016/j.envpol.2014.02.010


211

Park, Woodhall, J., Ma, G., Veinot, J.G.., Boxall, A.B.A., 2015. Do particle size and surface functionality affect uptake and depuration of gold nanoparticles by aquatic invertebrates? Environ. Toxicol. Chem. 34, 850–859. doi:10.1002/etc.2868

Plusquin, M., DeGheselle, O., Cuypers, A., Geerdens, E., Roten, A.V., Artois, T., Smeets, K., 2012. Reference genes for qPCR assays in toxic metal and salinity stress in two flatworm model organisms. Ecotoxicology 21, 475–484. doi:10.1007/s10646-011-0809-8

Poynton, H.C., Lazorchak, J.M., Impellitteri, C.A., Blalock, B.J., Rogers, K., Allen, H.J., Loguinov, A., Heckman, J.L., Govindasmawy, S., 2012. Toxicogenomic Responses of Nanotoxicity in Daphnia magna Exposed to Silver Nitrate and Coated Silver Nanoparticles. Environ. Sci. Technol. 46, 6288–6296. doi:10.1021/es3001618

Sakka, Y., Skjolding, L.M., Mackevica, A., Filser, J., Baun, A., 2016. Behavior and chronic toxicity of two differently stabilized silver nanoparticles to Daphnia magna. Aquat. Toxicol. 177, 526–535. doi:10.1016/j.aquatox.2016.06.025

Silva, T., Pokhrel, L.R., Dubey, B., Tolaymat, T.M., Maier, K.J., Liu, X., 2014. Particle size, surface charge and concentration dependent ecotoxicity of three organo-coated silver nanoparticles: Comparison between general linear model-predicted and observed toxicity. Sci. Total Environ. 468–469, 968–976. doi:10.1016/j.scitotenv.2013.09.006

Vale, G., Mehennaoui, K., Cambier, S., Libralato, G., Jomini, S., Domingos, R.F., 2016. Manufactured nanoparticles in the aquatic environment-biochemical responses on freshwater organisms: A critical overview. Aquat. Toxicol. 170, 162–174. doi:10.1016/j.aquatox.2015.11.019

Vance, M.E., Kuiken, T., Vejerano, E.P., McGinnis, S.P., Hochella, M.F., Rejeski, D., Hull, M.S., 2015. Nanotechnology in the real world: Redeveloping the nanomaterial consumer products inventory. Beilstein J. Nanotechnol. 6, 1769–1780. doi:10.3762/bjnano.6.181

Vasseur, P., Leguille, C., 2004. Defense systems of benthic invertebrates in response to environmental stressors. Environ. Toxicol. 19, 433–436. doi:10.1002/tox.20024

Volland, M., Hampel, M., Martos-Sitcha, J.A., Trombini, C., Martínez-Rodríguez, G., Blasco, J., 2015. Citrate gold nanoparticle exposure in the marine bivalve Ruditapes philippinarum: uptake, elimination and oxidative stress response. Environ. Sci. Pollut. Res. 22, 17414–17424. doi:10.1007/s11356-015-4718-x

213

Reference

Adam, N., Vergauwen, L., Blust, R., Knapen, D., 2015. Gene transcription patterns and energy reserves in Daphnia

magna show no nanoparticle specific toxicity when exposed to ZnO and CuO nanoparticles. Environ. Res. 138, 82–

92. doi:10.1016/j.envres.2015.02.014

Adam, O., Badot, P.-M., Degiorgi, F., Crini, G., 2009. Mixture toxicity assessment of wood preservative pesticides

in the freshwater amphipod Gammarus pulex (L.). Ecotoxicol. Environ. Saf. 72, 441–449.

doi:10.1016/j.ecoenv.2008.07.017

Ahn, J.-M., Eom, H.-J., Yang, X., Meyer, J.N., Choi, J., 2014. Comparative toxicity of silver nanoparticles on oxidative

stress and DNA damage in the nematode, Caenorhabditis elegans. Chemosphere 108, 343–352.


Ali, D., 2014. Oxidative Stress-Mediated Apoptosis and Genotoxicity Induced by Silver Nanoparticles in Freshwater

Snail Lymnea luteola L. Biol. Trace Elem. Res. 162, 333–341. doi:10.1007/s12011-014-0158-6

Ali, D., Yadav, P.G., Kumar, S., Ali, H., Alarifi, S., Harrath, A.H., 2014. Sensitivity of freshwater pulmonate snail

Lymnaea luteola L., to silver nanoparticles. Chemosphere. doi:10.1016/j.chemosphere.2013.10.081

Alonso, Á., Lange, H.J.D., Peeters, E.T.H.M., 2010. Contrasting sensitivities to toxicants of the freshwater

amphipods Gammarus pulex and G. fossarum. Ecotoxicology 19, 133–140. doi:10.1007/s10646-009-0398-y

Andersen, C.L., Jensen, J.L., Ørntoft, T.F., 2004. Normalization of Real-Time Quantitative Reverse Transcription-

PCR Data: A Model-Based Variance Estimation Approach to Identify Genes Suited for Normalization, Applied to

Bladder and Colon Cancer Data Sets. Cancer Res. 64, 5245–5250. doi:10.1158/0008-5472.CAN-04-0496




Angel, B.M., Batley, G.E., Jarolimek, C.V., Rogers, N.J., 2013. The impact of size on the fate and toxicity of

nanoparticulate silver in aquatic systems. Chemosphere. doi:10.1016/j.chemosphere.2013.04.096

Arab, K., Steghens, J.-P., 2004. Plasma lipid hydroperoxides measurement by an automated xylenol orange

method. Anal. Biochem. 325, 158–163. doi:10.1016/j.ab.2003.10.022



143, 73–84. doi:10.1016/j.aquatox.2013.07.012

Asghari, S., Johari, S.A., Lee, J.H., Kim, Y.S., Jeon, Y.B., Choi, H.J., Moon, M.C., Yu, I.J., 2012. Toxicity of various silver

nanoparticles compared to silver ions in Daphnia magna. J. Nanobiotechnology 10, 1–11. doi:10.1186/1477-3155-

10-14

AshaRani, P., Hande, M.P., Valiyaveettil, S., 2009. Anti-proliferative activity of silver nanoparticles. BMC Cell Biol.

10, 65. doi:10.1186/1471-2121-10-65

AshaRani, P.V., Low Kah Mun, G., Hande, M.P., Valiyaveettil, S., 2009. Cytotoxicity and Genotoxicity of Silver

Nanoparticles in Human Cells. ACS Nano 3, 279–290. doi:10.1021/nn800596w

Auffan, D.M., Santaella, D.C., Thiéry, P.A., Paillès, C., Rose, J., Achouak, D.W., Thill, D.A., Masion, A., Wiesner, M.,

Bottero, J.-Y., 2012. Ecotoxicity of Inorganic Nanoparticles: From Unicellular Organisms to Invertebrates, in:

214

Bhushan, P.B. (Ed.), Encyclopedia of Nanotechnology. Springer Netherlands, pp. 623–636.

Auffan, M., Rose, J., Wiesner, M., Bottero, J.-Y., 2009. Chemical stability of metallic nanoparticles : a parameter

controlling their potential cellular toxicity in vitro.

Baalousha, M., Nur, Y., Römer, I., Tejamaya, M., Lead, J.R., 2013. Effect of monovalent and divalent cations, anions

and fulvic acid on aggregation of citrate-coated silver nanoparticles. Sci. Total Environ. 454–455, 119–131.

doi:10.1016/j.scitotenv.2013.02.093




Balachandran, Y.L., Girija, S., Selvakumar, R., Tongpim, S., Gutleb, A.C., Suriyanarayanan, S., 2013. Differently

Environment Stable Bio-Silver Nanoparticles: Study on Their Optical Enhancing and Antibacterial Properties. PLoS

ONE 8, e77043. doi:10.1371/journal.pone.0077043

Balachandran, Y.L., Peranantham, P., Selvakumar, R., Gutleb, A.C., Girija, S., 2012. Size-Controlled Green Synthesis

of Silver Nanoparticles using Dual Functional Plant Leaf Extract at Room Temperature. Int. J. Green Nanotechnol.

4, 310–325. doi:10.1080/19430892.2012.706183

Barata, C., Varo, I., Navarro, J.C., Arun, S., Porte, C., 2005. Antioxidant enzyme activities and lipid peroxidation in

the freshwater cladoceran Daphnia magna exposed to redox cycling compounds. Comp. Biochem. Physiol. Part C

Toxicol. Pharmacol. 140, 175–186. doi:10.1016/j.cca.2005.01.013

Barnard, J.L., Barnard, C.M., 1983. Freshwater amphipoda of the world / J.L. & C.M. Barnard. - Version details

[WWW Document]. Mt Vernon Va Hayfield Assoc. URL http://trove.nla.gov.au/version/26012490 (accessed

4.22.16).

Batista, D., Pascoal, C., Cássio, F., 2017. How do physicochemical properties influence the toxicity of silver

nanoparticles on freshwater decomposers of plant litter in streams? Ecotoxicol. Environ. Saf. 140, 148–155.

doi:10.1016/j.ecoenv.2017.02.039




doi:10.1007/s11356-017-8400-3

Baumann, J., Köser, J., Arndt, D., Filser, J., 2014. The coating makes the difference: Acute effects of iron oxide

nanoparticles on Daphnia magna. Sci. Total Environ. 484, 176–184. doi:10.1016/j.scitotenv.2014.03.023

BCC RESEARCH, 2017. NAN021G_Report Overview.pdf [WWW Document]. URL

https://cdn2.hubspot.net/hubfs/308401/NAN%20Report%20Overviews/NAN021G_Report%20Overview.pdf?t=1

507039495677&utm_campaign=NAN021G&utm_source=hs_automation&utm_medium=email&utm_content=3

9598398&_hsenc=p2ANqtz-8aMzzG8s1ejWU5vfx62L2v9e7nraO-

2Zo5IxMvgn1QGf168euhj5LTcihQ1hQNtJhW680aqJqccZGQGLNnRuMjC9fE2g&_hsmi=39598398 (accessed

10.3.17).

Bebianno, M.J., Gonzalez-Rey, M., Gomes, T., Mattos, J.J., Flores-Nunes, F., Bainy, A.C.D., 2015. Is gene

transcription in mussel gills altered after exposure to Ag nanoparticles? Environ. Sci. Pollut. Res. 1–9.

doi:10.1007/s11356-015-5186-z

Beers, R.F., Sizer, I.W., 1951. A Spectrophotometric Method for Measuring the Breakdown of Hydrogen Peroxide

by Catalase. J. Biol. Chem. 195, 133–140.

215

Behr, M., Legay, S., Hausman, J.-F., Guerriero, G., 2015. Analysis of Cell Wall-Related Genes in Organs of Medicago

sativa L. under Different Abiotic Stresses. Int. J. Mol. Sci. 16, 16104–16124. doi:10.3390/ijms160716104

Besse, J.-P., Coquery, M., Lopes, C., Chaumot, A., Budzinski, H., Labadie, P., Geffard, O., 2013. Caged Gammarus

fossarum (Crustacea) as a robust tool for the characterization of bioavailable contamination levels in continental

waters: Towards the determination of threshold values. Water Res. 47, 650–660.

doi:10.1016/j.watres.2012.10.024

Bianchini, A., Wood, C.M., 2003. Mechanism of acute silver toxicity in Daphnia magna. Environ. Toxicol. Chem. 22,

1361–1367. doi:10.1002/etc.5620220624

Bianchini, A., Wood, C.M., 2002. Physiological effects of chronic silver exposure in Daphnia magna. Comp.

Biochem. Physiol. Toxicol. Pharmacol. CBP 133, 137–145.

Bickford, D., Lohman, D.J., Sodhi, N.S., Ng, P.K.L., Meier, R., Winker, K., Ingram, K.K., Das, I., 2007. Cryptic species

as a window on diversity and conservation. Trends Ecol. Evol. 22, 148–155. doi:10.1016/j.tree.2006.11.004

Bilberg, K., Malte, H., Wang, T., Baatrup, E., 2010. Silver nanoparticles and silver nitrate cause respiratory stress in

Eurasian perch (Perca fluviatilis). Aquat. Toxicol. 96, 159–165. doi:10.1016/j.aquatox.2009.10.019

Blaser, S.A., Scheringer, M., MacLeod, M., Hungerbühler, K., 2008. Estimation of cumulative aquatic exposure and

risk due to silver: Contribution of nano-functionalized plastics and textiles. Sci. Total Environ. 390, 396–409.

doi:10.1016/j.scitotenv.2007.10.010

Blinova, I., Niskanen, J., Kajankari, P., Kanarbik, L., Käkinen, A., Tenhu, H., Penttinen, O.-P., Kahru, A., 2012. Toxicity

of two types of silver nanoparticles to aquatic crustaceans Daphnia magna and Thamnocephalus platyurus.

Environ. Sci. Pollut. Res. 1–8. doi:10.1007/s11356-012-1290-5

Böhme, S., Baccaro, M., Schmidt, M., Potthoff, A., Stärk, H.-J., Reemtsma, T., Kühnel, D., 2017. Metal uptake and

distribution in the zebrafish (Danio rerio) embryo: differences between nanoparticles and metal ions. Environ. Sci.

Nano 4, 1005–1015. doi:10.1039/C6EN00440G

Bone, A.J., Colman, B.P., Gondikas, A.P., Newton, K.M., Harrold, K.H., Cory, R.M., Unrine, J.M., Klaine, S.J., Matson,

C.W., Di Giulio, R.T., 2012. Biotic and Abiotic Interactions in Aquatic Microcosms Determine Fate and Toxicity of

Ag Nanoparticles: Part 2–Toxicity and Ag Speciation. Environ. Sci. Technol. 46, 6925–6933.

doi:10.1021/es204683m

Borowsky, B., 1984. The Use of the Males' Gnathopods During Precopulation in Some Gammaridean

Amphipods. Crustaceana 47, 245–250. doi:10.1163/156854084X00504



Bour, A., Mouchet, F., Silvestre, J., Gauthier, L., Pinelli, E., 2015. Environmentally relevant approaches to assess

nanoparticles ecotoxicity: a review. J. Hazard. Mater. 283, 764–777. doi:10.1016/j.jhazmat.2014.10.021

Bousfield, E.L., 1973. shallow-water-gammaridean-amphipoda-of-new-england-handbooks-of-american.pdf

[WWW Document]. URL http://www.thebookishblog.com/shallow-water-gammaridean-amphipoda-of-new-

england-handbooks-of-american.pdf (accessed 8.20.17).

Bowman, C.R., Bailey, F.C., Elrod-Erickson, M., Neigh, A.M., Otter, R.R., 2012. Effects of silver nanoparticles on

zebrafish (Danio rerio) and Escherichia coli (ATCC 25922): a comparison of toxicity based on total surface area

versus mass concentration of particles in a model eukaryotic and prokaryotic system. Environ. Toxicol. Chem. 31,

1793–1800. doi:10.1002/etc.1881

216

Bozich, J.S., Lohse, S.E., Torelli, M.D., Murphy, C.J., Hamers, R.J., Klaper, R.D., 2014. Surface chemistry, charge and

ligand type impact the toxicity of gold nanoparticles to Daphnia magna. Environ. Sci. Nano 1, 260–270.

doi:10.1039/C4EN00006D

Brigelius-Flohé, R., Maiorino, M., 2013. Glutathione peroxidases. Biochim. Biophys. Acta BBA - Gen. Subj., Cellular

functions of glutathione 1830, 3289–3303. doi:10.1016/j.bbagen.2012.11.020

Brown, J.K., Frohlich, D.R., Rosell, R.C., 1995. The Sweetpotato or Silverleaf Whiteflies: Biotypes of Bemisia tabaci

or a Species Complex? Annu. Rev. Entomol. 40, 511–534. doi:10.1146/annurev.en.40.010195.002455

Brulle, F., Bernard, F., Vandenbulcke, F., Cuny, D., Dumez, S., 2014. Identification of suitable qPCR reference genes

in leaves of Brassica oleracea under abiotic stresses. Ecotoxicology 23, 459–471. doi:10.1007/s10646-014-1209-7



Buffet, P.-E., Pan, J.-F., Poirier, L., Amiard-Triquet, C., Amiard, J.-C., Gaudin, P., Faverney, C.R., Guibbolini, M.,

Gilliland, D., Valsami-Jones, E., Mouneyrac, C., 2013. Biochemical and behavioural responses of the endobenthic

bivalve Scrobicularia plana to silver nanoparticles in seawater and microalgal food. Ecotoxicol. Environ. Saf. 89,

117–124. doi:10.1016/j.ecoenv.2012.11.019

Burić, P., Jakšić, Ž., Štajner, L., Dutour Sikirić, M., Jurašin, D., Cascio, C., Calzolai, L., Lyons, D.M., 2015. Effect of

silver nanoparticles on Mediterranean sea urchin embryonal development is species specific and depends on

moment of first exposure. Mar. Environ. Res., Particles in the Oceans: Implication for a safe marine environment

111, 50–59. doi:10.1016/j.marenvres.2015.06.015

Bury, N.R., Shaw, J., Glover, C., Hogstrand, C., 2002. Derivation of a toxicity-based model to predict how water

chemistry influences silver toxicity to invertebrates. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol. 133, 259–

270. doi:10.1016/S1532-0456(02)00096-0

Bustin, S.A., Benes, V., Garson, J.A., Hellemans, J., Huggett, J., Kubista, M., Mueller, R., Nolan, T., Pfaffl, M.W.,

Shipley, G.L., Vandesompele, J., Wittwer, C.T., 2009. The MIQE Guidelines: Minimum Information for Publication

of Quantitative Real-Time PCR Experiments. Clin. Chem. 55, 611–622. doi:10.1373/clinchem.2008.112797

Cambier, S., Røgeberg, M., Georgantzopoulou, A., Serchi, T., Karlsson, C., Verhaegen, S., Iversen, T.-G., Guignard,

C., Kruszewski, M., Hoffmann, L., Audinot, J.-N., Ropstad, E., Gutleb, A.C., 2018. Fate and effects of silver

nanoparticles on early life-stage development of zebrafish (Danio rerio) in comparison to silver nitrate. Sci. Total


Canesi, L., Balbi, T., Fabbri, R., Salis, A., Damonte, G., Volland, M., Blasco, J., 2017. Biomolecular coronas in

invertebrate species: Implications in the environmental impact of nanoparticles. NanoImpact.

doi:10.1016/j.impact.2017.08.001

Canesi, L., Ciacci, C., Fabbri, R., Marcomini, A., Pojana, G., Gallo, G., 2012. Bivalve molluscs as a unique target group

for nanoparticle toxicity. Mar. Environ. Res., Emerging and persistent impacts on Marine Organisms: Detection

methods and action mechanisms 76, 16–21. doi:10.1016/j.marenvres.2011.06.005

Chamier, A.-C., Willoughby, L.G., 1986. The role of fungi in the diet of the amphipod Gammarus pulex (L.): an

enzymatic study. Freshw. Biol. 16, 197–208. doi:10.1111/j.1365-2427.1986.tb00964.x

Charron, L., 2014. Biomarqueurs énergétiques chez u amphipode d’eau douce Gammarus fossarum" Développement, lien avec le succés reproducteur et application in situ. Université de Reims-Champagne-Ardenne.

217

Chen, Y., Si, Y., Zhou, D., Dang, F., 2017. Differential bioaccumulation patterns of nanosized and dissolved silver in

a land snail Achatina fulica. Environ. Pollut. 222, 50–57. doi:10.1016/j.envpol.2017.01.007

Chevreux, É., Fage, L., Faunistique, F. des S.F. de S.N.O.C. de, 1970. Amphipodes. Kraus.



doi:10.1016/j.aquatox.2009.12.012

Clemente, Z., Castro, V.L.S.S., Moura, M.A.M., Jonsson, C.M., Fraceto, L.F., 2014. Toxicity assessment of TiO2

nanoparticles in zebrafish embryos under different exposure conditions. Aquat. Toxicol. 147, 129–139.

doi:10.1016/j.aquatox.2013.12.024

Cleveland, D., Long, S.E., Pennington, P.L., Cooper, E., Fulton, M.H., Scott, G.I., Brewer, T., Davis, J., Petersen, E.J.,

Wood, L., 2012. Pilot estuarine mesocosm study on the environmental fate of Silver nanomaterials leached from

consumer products. Sci. Total Environ., Special Section: Reviews of Trace Metal Pollution in China 421–422, 267–

272. doi:10.1016/j.scitotenv.2012.01.025

Connolly, M., Fernandez-Cruz, M.-L., Quesada-Garcia, A., Alte, L., Segner, H., Navas, J.M., 2015. Comparative

Cytotoxicity Study of Silver Nanoparticles (AgNPs) in a Variety of Rainbow Trout Cell Lines (RTL-W1, RTH-149, RTG-

2) and Primary Hepatocytes. Int. J. Environ. Res. Public. Health 12, 5386–5405. doi:10.3390/ijerph120505386

Copilaş-Ciocianu, D., Rutová, T., Pařil, P., Petrusek, A., 2017. Epigean gammarids survived millions of years of severe

climatic fluctuations in high latitude refugia throughout the Western Carpathians. Mol. Phylogenet. Evol. 112, 218–

229. doi:10.1016/j.ympev.2017.04.027

Correia, A.D., Pereira, A.L., Costa, M.H., Carrapi??o, F., 2002. Functional anatomy of the midgut gland of Gammarus

locusta (Crustacea: Amphipoda). J. Mar. Biol. Assoc. UK 82, 201–204.

Coulaud, R., Geffard, O., Xuereb, B., Lacaze, E., Quéau, H., Garric, J., Charles, S., Chaumot, A., 2011. In situ feeding

assay with Gammarus fossarum (Crustacea): Modelling the influence of confounding factors to improve water

quality biomonitoring. Water Res. 45, 6417–6429. doi:10.1016/j.watres.2011.09.035

Cristescu, M.E., Hebert, P.D., 2005. The “Crustacean Seas” — an evolutionary perspective on the Ponto–Caspian

peracarids. Can. J. Fish. Aquat. Sci. 62, 505–517. doi:10.1139/f04-210

Croteau, M.-N., Misra, S.K., Luoma, S.N., Valsami-Jones, E., 2011. Silver Bioaccumulation Dynamics in a Freshwater

Invertebrate after Aqueous and Dietary Exposures to Nanosized and Ionic Ag. Environ. Sci. Technol. 45, 6600–

6607. doi:10.1021/es200880c

Cumberland, S.A., Lead, J.R., 2009. Particle size distributions of silver nanoparticles at environmentally relevant

conditions. J. Chromatogr. A 1216, 9099–9105. doi:10.1016/j.chroma.2009.07.021

Cupi, D., Hartmann, N.B., Baun, A., 2015. The influence of natural organic matter and aging on suspension stability

in guideline toxicity testing of silver, zinc oxide, and titanium dioxide nanoparticles with Daphnia magna. Environ.


Dale, A.L., Lowry, G.V., Casman, E.A., 2013. Modeling Nanosilver Transformations in Freshwater Sediments.

Environ. Sci. Technol. 47, 12920–12928. doi:10.1021/es402341t

Dangles, O., Guérold, F., 2001. Linking Shredders and Leaf Litter Processing: Insights from an Acidic Stream Study.

Int. Rev. Hydrobiol. 86, 395–406. doi:10.1002/1522-2632(200107)86:4/5<395::AID-IROH395>3.0.CO;2-1

Dayal, N., Thakur, M., Patil, P., Singh, D., Vanage, G., Joshi, D.S., 2016. Histological and genotoxic evaluation of gold

nanoparticles in ovarian cells of zebrafish (Danio rerio). J. Nanoparticle Res. 18, 291. doi:10.1007/s11051-016-

218

3549-0

De Coen, W.M.D., Janssen, C.R., 1997. The use of biomarkers in Daphnia magna toxicity testing. IV. Cellular Energy

Allocation: a new methodology to assess the energy budget of toxicant-stressed Daphnia populations. J. Aquat.

Ecosyst. Stress Recovery 6, 43–55. doi:10.1023/A:1008228517955

De Spiegelaere, W., Dern-Wieloch, J., Weigel, R., Schumacher, V., Schorle, H., Nettersheim, D., Bergmann, M.,

Brehm, R., Kliesch, S., Vandekerckhove, L., Fink, C., 2015. Reference Gene Validation for RT-qPCR, a Note on

Different Available Software Packages. PLoS ONE 10. doi:10.1371/journal.pone.0122515

Dedeh, A., Ciutat, A., Treguer-Delapierre, M., Bourdineaud, J.-P., 2015. Impact of gold nanoparticles on zebrafish

exposed to a spiked sediment. Nanotoxicology 9, 71–80. doi:10.3109/17435390.2014.889238

Dedourge-Geffard, O., Charron, L., Hofbauer, C., Gaillet, V., Palais, F., Lacaze, E., Geffard, A., Geffard, O., 2013.

Temporal patterns of digestive enzyme activities and feeding rate in gammarids (Gammarus fossarum) exposed to

inland polluted waters. Ecotoxicol. Environ. Saf. 97, 139–146. doi:10.1016/j.ecoenv.2013.07.016




Deponte, M., 2013. Glutathione catalysis and the reaction mechanisms of glutathione-dependent enzymes.

Biochim. Biophys. Acta BBA - Gen. Subj., Cellular functions of glutathione 1830, 3217–3266.

doi:10.1016/j.bbagen.2012.09.018



Biol. 58, 419–472. doi:10.1163/157075608X383719

Domingos, R.F., Baalousha, M.A., Ju-Nam, Y., Reid, M.M., Tufenkji, N., Lead, J.R., Leppard, G.G., Wilkinson, K.J.,

2009. Characterizing Manufactured Nanoparticles in the Environment: Multimethod Determination of Particle

Sizes. Environ. Sci. Technol. 43, 7277–7284. doi:10.1021/es900249m

Domingos, R.F., Franco, C., Pinheiro, J.P., 2015a. The role of charged polymer coatings of nanoparticles on the

speciation and fate of metal ions in the environment. Environ. Sci. Pollut. Res. Int. 22, 2900–2906.

doi:10.1007/s11356-014-3546-8

Domingos, R.F., Gélabert, A., Carreira, S., Cordeiro, A., Sivry, Y., Benedetti, M.F., 2015b. Metals in the Aquatic

Environment—Interactions and Implications for the Speciation and Bioavailability: A Critical Overview. Aquat.

Geochem. 21, 231–257. doi:10.1007/s10498-014-9251-x

Domingos, R.F., Peyrot, C., Wilkinson, K.J., 2010. Aggregation of titanium dioxide nanoparticles: role of calcium

and phosphate. Environ. Chem.

Domingos, R.F., Rafiei, Z., Monteiro, C.E., Khan, M.A.K., Wilkinson, K.J., 2013. Agglomeration and dissolution of

zinc oxide nanoparticles: role of pH, ionic strength and fulvic acid. Environ. Chem. 10, 306–312.

doi:10.1071/EN12202

Dong, M., Zhang, X., Chi, X., Mou, S., Xu, J., Xu, D., Wang, W., Ye, N., 2012. The validity of a reference gene is highly

dependent on the experimental conditions in green alga Ulva linza. Curr. Genet. 58, 13–20. doi:10.1007/s00294-

011-0361-3

Dowsett, D., Audinot, J.-N., Vollnhals, F., Eswara, S., Wirtz, T., 2016. High Resolution Chemical Imaging on the

Helium Ion Microscope, in: European Microscopy Congress 2016: Proceedings. Wiley-VCH Verlag GmbH & Co.

219

KGaA. doi:10.1002/9783527808465.EMC2016.6559

Dunel-Erb, S., Barradas, C., Lignon, J., 1997. Morphological Evidence for the Existence of Two Distinct Types of

Mitochondria Rich Cells in the Gill of the Crayfish Astacus leptodactylusEschscholtz. Acta Zool. 78, 195–203.

doi:10.1111/j.1463-6395.1997.tb01006.x

Ellis, L.-J.A., Valsami-Jones, E., Lead, J.R., Baalousha, M., 2016. Impact of surface coating and environmental

conditions on the fate and transport of silver nanoparticles in the aquatic environment. Sci. Total Environ. 568,

95–106. doi:10.1016/j.scitotenv.2016.05.199

Erel, O., 2004. A novel automated direct measurement method for total antioxidant capacity using a new

generation, more stable ABTS radical cation. Clin. Biochem. 37, 277–285. doi:10.1016/j.clinbiochem.2003.11.015

Eybe, T., Bohn, T., Audinot, J.N., Udelhoven, T., Cauchie, H.M., Migeon, H.N., Hoffmann, L., 2009. Uptake

visualization of deltamethrin by NanoSIMS and acute toxicity to the water flea Daphnia magna. Chemosphere 76,

134–140. doi:10.1016/j.chemosphere.2009.02.005



Farkas, A., Salánki, J., Varanka, I., 2003. Crustaceans as biological indicators of heavy metal pollution in Lake

Balaton (Hungary). Hydrobiologia 506–509, 359–364. doi:10.1023/B:HYDR.0000008615.86330.1d

Farkas, J., Christian, P., Gallego-Urrea, J.A., Roos, N., Hassellöv, M., Tollefsen, K.E., Thomas, K.V., 2011. Uptake and

effects of manufactured silver nanoparticles in rainbow trout (Oncorhynchus mykiss) gill cells. Aquat. Toxicol. 101,

117–125. doi:10.1016/j.aquatox.2010.09.010

Farmen, E., Mikkelsen, H.N., Evensen, Ø., Einset, J., Heier, L.S., Rosseland, B.O., Salbu, B., Tollefsen, K.E., Oughton,

D.H., 2012. Acute and sub-lethal effects in juvenile Atlantic salmon exposed to low μg/L concentrations of Ag

nanoparticles. Aquat. Toxicol., Proceedings from the 16th International Symposium on Pollutant Responses in

Marine Organisms (PRIMO16) 108, 78–84. doi:10.1016/j.aquatox.2011.07.007

Farré, M., Gajda-Schrantz, K., Kantiani, L., Barceló, D., 2008. Ecotoxicity and analysis of nanomaterials in the aquatic

environment. Anal. Bioanal. Chem. 393, 81–95. doi:10.1007/s00216-008-2458-1

Feckler, A., Thielsch, A., Schwenk, K., Schulz, R., Bundschuh, M., 2012. Differences in the sensitivity among cryptic

lineages of the Gammarus fossarum complex. Sci. Total Environ. 439, 158–164.

doi:10.1016/j.scitotenv.2012.09.003

Feckler, A., Zubrod, J.P., Thielsch, A., Schwenk, K., Schulz, R., Bundschuh, M., 2014. Cryptic species diversity: an

overlooked factor in environmental management? J. Appl. Ecol. 51, 958–967. doi:10.1111/1365-2664.12246

Felten, V., 2003. Effets de l’acidification des ruisseaux vosgiens sur la biologie, l’écologie et l’écop[hysiologie de Hammarus fossarum Koch, 1835 (Crustacea Amphipoda): Approche integrée à différentsniveaux d’organisation. Université de Metz. Felten, V., Baudoin, J.M., Guérold, F., 2006. Physiological recovery from episodic acid stress does not mean

population recovery of Gammarus fossarum. Chemosphere 65, 988–998. doi:10.1016/j.chemosphere.2006.03.059


Physiological and behavioural responses of Gammarus pulex (Crustacea: Amphipoda) exposed to cadmium. Aquat.


Felten, V., Guérold, F., 2006. Short-term physiological responses to a severe acid stress in three macroinvertebrate

220

species: A comparative study. Chemosphere 63, 1427–1435. doi:10.1016/j.chemosphere.2005.10.003

Fielding, N.J., MacNeil, C., Dick, J.T.A., Elwood, R.W., Riddell, G.E., Dunn, A.M., 2003. Effects of the

acanthocephalan parasite Echinorhynchus truttae on the feeding ecology of Gammarus pulex (Crustacea:

Amphipoda). J. Zool. 261, 321–325. doi:10.1017/S0952836903004230



doi:10.1002/etc.5620190820

Fu, P.P., Xia, Q., Hwang, H.-M., Ray, P.C., Yu, H., 2014. Mechanisms of nanotoxicity: Generation of reactive oxygen

species. J. Food Drug Anal., Nanomaterials - Toxicology and Medical Applications 22, 64–75.

doi:10.1016/j.jfda.2014.01.005

Gagné, F., André, C., Skirrow, R., Gélinas, M., Auclair, J., van Aggelen, G., Turcotte, P., Gagnon, C., 2012. Toxicity of

silver nanoparticles to rainbow trout: A toxicogenomic approach. Chemosphere 89, 615–622.


Gagné, F., Blaise, C., Pellerin, J., 2005. Altered exoskeleton composition and vitellogenesis in the crustacean

Gammarus sp. collected at polluted sites in the Saguenay Fjord, Quebec, Canada. Environ. Res. 98, 89–99.

doi:10.1016/j.envres.2004.09.008

Gaiser, B.K., Biswas, A., Rosenkranz, P., Jepson, M.A., Lead, J.R., Stone, V., Tyler, C.R., Fernandes, T.F., 2011. Effects

of silver and cerium dioxide micro- and nano-sized particles on Daphnia magna. J. Environ. Monit. 13, 1227–1235.

doi:10.1039/C1EM10060B

Gallego-Urrea, J.A., Tuoriniemi, J., Hassellöv, M., 2011. Applications of particle-tracking analysis to the

determination of size distributions and concentrations of nanoparticles in environmental, biological and food

samples. TrAC Trends Anal. Chem., Characterization, Analysis and Risks of Nanomaterials in Environmental and

Food Samples II 30, 473–483. doi:10.1016/j.trac.2011.01.005

Gandar, A., Laffaille, P., Canlet, C., Tremblay, M., Gautier, R., Perrault, A., Gress, L., Mormède, P., Tapie, N.,

Budzinski, H., Jean, S., 2017. Adaptive response under multiple stress exposure in fish: From the molecular to

individual level. Chemosphere 188. doi:10.1016/j.chemosphere.2017.08.089

Gao, J., Powers, K., Wang, Y., Zhou, H., Roberts, S.M., Moudgil, B.M., Koopman, B., Barber, D.S., 2012. Influence of

Suwannee River humic acid on particle properties and toxicity of silver nanoparticles. Chemosphere 89, 96–101.


Garaud, M., Auffan, M., Devin, S., Felten, V., Pagnout, C., Pain-Devin, S., Proux, O., Rodius, F., Sohm, B., Giamberini,

L., 2016. Integrated assessment of ceria nanoparticle impacts on the freshwater bivalve Dreissena polymorpha.

Nanotoxicology 10, 935–944. doi:10.3109/17435390.2016.1146363

Garaud, M., Auffan, M., Devin, S., Felten, V., Pagnout, C., Pain-Devin, S., Proux, O., Rodius, F., Sohm, B., Giambérini,

L., 2015. Fate and integrated assessment of ceria nanoparticle impacts on the freshwater bivalve Dreissena

polymorpha: a mesocosm approach. Nanotoxicology.

Garaud, Trapp, J., Devin, S., Cossu-Leguille, C., Pain-Devin, S., Felten, V., Giamberini, L., 2015. Multibiomarker

assessment of cerium dioxide nanoparticle (nCeO2) sublethal effects on two freshwater invertebrates, Dreissena

polymorpha and Gammarus roeseli. Aquat. Toxicol. 158, 63–74. doi:10.1016/j.aquatox.2014.11.004

García-Alonso, J., Rodriguez-Sanchez, N., Misra, S.K., Valsami-Jones, E., Croteau, M.-N., Luoma, S.N., Rainbow, P.S.,

2014. Toxicity and accumulation of silver nanoparticles during development of the marine polychaete Platynereis

dumerilii. Sci. Total Environ. 476–477, 688–695. doi:10.1016/j.scitotenv.2014.01.039

221

García-Cambero, J.P., Núñez García, M., López, G.D., Herranz, A.L., Cuevas, L., Pérez-Pastrana, E., Cuadal, J.S.,

Castelltort, M.R., Calvo, A.C., 2013. Converging hazard assessment of gold nanoparticles to aquatic organisms.


García-Negrete, C.A., Blasco, J., Volland, M., Rojas, T.C., Hampel, M., Lapresta-Fernández, A., Jiménez de Haro,

M.C., Soto, M., Fernández, A., 2013. Behaviour of Au-citrate nanoparticles in seawater and accumulation in

bivalves at environmentally relevant concentrations. Environ. Pollut. 174, 134–141.

doi:10.1016/j.envpol.2012.11.014

Georgantzopoulou, A., 2015. Effects of silver nanoparticles and ions and interaction with first line of defense.

Wageningen University, Wageningen Netherlands.




doi:10.3109/17435390.2012.715312

Georgantzopoulou, A., Cambier, S., Serchi, T., Kruszewski, M., Balachandran, Y.L., Grysan, P., Audinot, J.-N., Ziebel,

J., Guignard, C., Gutleb, A.C., Murk, A.J., 2016. Inhibition of multixenobiotic resistance transporters (MXR) by silver

nanoparticles and ions in vitro and in Daphnia magna. Sci. Total Environ. 569–570, 681–689.

doi:10.1016/j.scitotenv.2016.06.157

Gismondi, E., Beisel, J.-N., Cossu-Leguille, C., 2012a. Influence of gender and season on reduced glutathione

concentration and energy reserves of Gammarus roeseli. Environ. Res. 118, 47–52.

doi:10.1016/j.envres.2012.06.004

Gismondi, E., Cossu-Leguille, C., Beisel, J.-N., 2012b. Acanthocephalan parasites: help or burden in gammarid


Gismondi, E., Rigaud, T., Beisel, J.-N., Cossu-Leguille, C., 2012c. Microsporidia parasites disrupt the responses to


Gomes, S.I.L., Soares, A.M.V.M., Scott-Fordsmand, J.J., Amorim, M.J.B., 2013. Mechanisms of response to silver

nanoparticles on Enchytraeus albidus (Oligochaeta): Survival, reproduction and gene expression profile. J. Hazard.

Mater. 254–255, 336–344. doi:10.1016/j.jhazmat.2013.04.005

Gomes, T., Araújo, O., Pereira, R., Almeida, A.C., Cravo, A., Bebianno, M.J., 2013a. Genotoxicity of copper oxide

and silver nanoparticles in the mussel Mytilus galloprovincialis. Mar. Environ. Res. 84, 51–59.

doi:10.1016/j.marenvres.2012.11.009

Gomes, T., Chora, S., Pereira, C.G., Cardoso, C., Bebianno, M.J., 2014a. Proteomic response of mussels Mytilus

galloprovincialis exposed to CuO NPs and Cu2+: An exploratory biomarker discovery. Aquat. Toxicol. 155, 327–

336. doi:10.1016/j.aquatox.2014.07.015

Gomes, T., Pereira, C.G., Cardoso, C., Bebianno, M.J., 2013b. Differential protein expression in mussels Mytilus

galloprovincialis exposed to nano and ionic Ag. Aquat. Toxicol. 136–137, 79–90.

doi:10.1016/j.aquatox.2013.03.021

Gomes, T., Pereira, C.G., Cardoso, C., Pinheiro, J.P., Cancio, I., Bebianno, M.J., 2012. Accumulation and toxicity of

copper oxide nanoparticles in the digestive gland of Mytilus galloprovincialis. Aquat. Toxicol. 118–119, 72–79.

doi:10.1016/j.aquatox.2012.03.017

Gomes, T., Pereira, C.G., Cardoso, C., Sousa, V.S., Teixeira, M.R., Pinheiro, J.P., Bebianno, M.J., 2014b. Effects of

silver nanoparticles exposure in the mussel Mytilus galloprovincialis. Mar. Environ. Res.

222

doi:10.1016/j.marenvres.2014.07.004

Gottschalk, F., Sonderer, T., Scholz, R.W., Nowack, B., 2009. Modeled Environmental Concentrations of Engineered

Nanomaterials (TiO2, ZnO, Ag, CNT, Fullerenes) for Different Regions. Environ. Sci. Technol. 43, 9216–9222.

doi:10.1021/es9015553



Graça, M. a. S., Maltby, L., Calow, P., 1994. Comparative ecology of Gammarus pulex (L.) and Asellus aquaticus (L.)

I: population dynamics and microdistribution. Hydrobiologia 281, 155–162. doi:10.1007/BF00028678

Grassé, P.-P., 1961. Traité de zoologie: anatomie, systématique, biologie. Tome IV, Premier fascicule Tome IV,

Premier fascicule. Masson, Paris.



Grosell, M., Nielsen, C., Bianchini, A., 2002. Sodium turnover rate determines sensitivity to acute copper and silver

exposure in freshwater animals. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol. 133, 287–303.

doi:10.1016/S1532-0456(02)00085-6

Gross, M.Y., Maycock, D.S., Thorndyke, M.C., Morritt, D., Crane, M., 2001. Abnormalities in sexual development of

the amphipod Gammarus pulex (L.) found below sewage treatment works. Environ. Toxicol. Chem. 20, 1792–1797.

doi:10.1002/etc.5620200824

Guerriero, G., Legay, S., Hausman, J.-F., 2014. Alfalfa Cellulose Synthase Gene Expression under Abiotic Stress: A

Hitchhiker’s Guide to RT-qPCR Normalization. PLOS ONE 9, e103808. doi:10.1371/journal.pone.0103808

Gündel, U., Benndorf, D., von Bergen, M., Altenburger, R., Küster, E., 2007. Vitellogenin cleavage products as

indicators for toxic stress in zebra fish embryos: A proteomic approach. PROTEOMICS 7, 4541–4554.

doi:10.1002/pmic.200700381

Hansen, S.F., Roverskov Heggelund, L., Besora, P.R., Mackevica, A., Boldrin, A., Baun, A., 2016. Nanoproducts –

what is actually available to European consumers? Environ. Sci. Nano 3, 169–180. doi:10.1039/C5EN00182J

Heckmann, L.-H., Connon, R., Hutchinson, T.H., Maund, S.J., Sibly, R.M., Callaghan, A., 2006. Expression of target

and reference genes in Daphnia magna exposed to ibuprofen. BMC Genomics 7, 1–8. doi:10.1186/1471-2164-7-

175

Heinlaan, M., Ivask, A., Blinova, I., Dubourguier, H.-C., Kahru, A., 2008. Toxicity of nanosized and bulk ZnO, CuO

and TiO2 to bacteria Vibrio fischeri and crustaceans Daphnia magna and Thamnocephalus platyurus. Chemosphere

71, 1308–1316. doi:10.1016/j.chemosphere.2007.11.047

Heinlaan, M., Kahru, A., Kasemets, K., Arbeille, B., Prensier, G., Dubourguier, H.-C., 2011. Changes in the Daphnia

magna midgut upon ingestion of copper oxide nanoparticles: a transmission electron microscopy study. Water

Res. 45, 179–190. doi:10.1016/j.watres.2010.08.026



doi:10.3389/fphys.2012.00431

Henry, Y., Piscart, C., Charles, S., Colinet, H., 2017. Combined effect of temperature and ammonia on molecular

response and survival of the freshwater crustacean Gammarus pulex. Ecotoxicol. Environ. Saf. 137, 42–48.

doi:10.1016/j.ecoenv.2016.11.011

223

Hole, P., Sillence, K., Hannell, C., Maguire, C.M., Roesslein, M., Suarez, G., Capracotta, S., Magdolenova, Z., Horev-

Azaria, L., Dybowska, A., Cooke, L., Haase, A., Contal, S., Manø, S., Vennemann, A., Sauvain, J.-J., Staunton, K.C.,

Anguissola, S., Luch, A., Dusinska, M., Korenstein, R., Gutleb, A.C., Wiemann, M., Prina-Mello, A., Riediker, M.,

Wick, P., 2013. Interlaboratory comparison of size measurements on nanoparticles using nanoparticle tracking

analysis (NTA). J. Nanoparticle Res. 15, 1–12. doi:10.1007/s11051-013-2101-8

Hood, E., 2004. Nanotechnology: Looking As We Leap. Environ. Health Perspect. 112, A740–A749.

Hotze, E.M., Bottero, J.-Y., Wiesner, M.R., 2010. Theoretical framework for nanoparticle reactivity as a function of

aggregation state. Langmuir 26, 11170–11175. doi:10.1021/1a9046963

Hu, W., Culloty, S., Darmody, G., Lynch, S., Davenport, J., Ramirez-Garcia, S., Dawson, K.A., Lynch, I., Blasco, J.,

Sheehan, D., 2014. Toxicity of copper oxide nanoparticles in the blue mussel, Mytilus edulis: A redox proteomic

investigation. Chemosphere 108, 289–299. doi:10.1016/j.chemosphere.2014.01.054

Isnard, P., Flammarion, P., Roman, G., Babut, M., Bastien, P., Bintein, S., Esserméant, L., Férard, J.F., Gallotti-

Schmitt, S., Saouter, E., Saroli, M., Thiébaud, H., Tomassone, R., Vindimian, E., 2001. Statistical analysis of

regulatory ecotoxicity tests. Chemosphere 45, 659–669. doi:10.1016/S0045-6535(00)00600-7

Ivask, A., ElBadawy, A., Kaweeteerawat, C., Boren, D., Fischer, H., Ji, Z., Chang, C.H., Liu, R., Tolaymat, T., Telesca,

D., Zink, J.I., Cohen, Y., Holden, P.A., Godwin, H.A., 2014. Toxicity Mechanisms in Escherichia coli Vary for Silver

Nanoparticles and Differ from Ionic Silver. ACS Nano 8, 374–386. doi:10.1021/nn4044047



Jensen, L.H.S., Skjolding, L.M., Thit, A., Sørensen, S.N., Købler, C., Mølhave, K., Baun, A., 2017. Not all that glitters

is gold—Electron microscopy study on uptake of gold nanoparticles in Daphnia magna and related artifacts.

Environ. Toxicol. Chem. 36, 1503–1509. doi:10.1002/etc.3697

Jovanović, B., Ji, T., Palić, D., 2011. Gene expression of zebrafish embryos exposed to titanium dioxide

nanoparticles and hydroxylated fullerenes. Ecotoxicol. Environ. Saf. 74, 1518–1525.

doi:10.1016/j.ecoenv.2011.04.012



Kaegi, R., Sinnet, B., Zuleeg, S., Hagendorfer, H., Mueller, E., Vonbank, R., Boller, M., Burkhardt, M., 2010. Release

of silver nanoparticles from outdoor facades. Environ. Pollut. 158, 2900–2905. doi:10.1016/j.envpol.2010.06.009

Kaegi, R., Voegelin, A., Ort, C., Sinnet, B., Thalmann, B., Krismer, J., Hagendorfer, H., Elumelu, M., Mueller, E., 2013.

Fate and transformation of silver nanoparticles in urban wastewater systems. Water Res. 47, 3866–3877.

doi:10.1016/j.watres.2012.11.060

Karaman, G., 1977. freshwater Gammarus species from europe, North Africa and adjacent regions of Asia.

Gammarus pulex group and related species.pdf [WWW Document]. URL

http://www.nativefishlab.net/library/textpdf/17884.pdf (accessed 2.27.14).

Keller, A.A., McFerran, S., Lazareva, A., Suh, S., 2013. Global life cycle releases of engineered nanomaterials. J.

Nanoparticle Res. 15, 1692. doi:10.1007/s11051-013-1692-4

Keller, A.A., Wang, H., Zhou, D., Lenihan, H.S., Cherr, G., Cardinale, B.J., Miller, R., Ji, Z., 2010. Stability and

Aggregation of Metal Oxide Nanoparticles in Natural Aqueous Matrices. Environ. Sci. Technol. 44, 1962–1967.

doi:10.1021/es902987d

224



Khan, F.R., Kennaway, G.M., Croteau, M.-N., Dybowska, A., Smith, B.D., Nogueira, A.J.A., Rainbow, P.S., Luoma,

S.N., Valsami-Jones, E., 2014. In vivo retention of ingested Au NPs by Daphnia magna: No evidence for trans-

epithelial alimentary uptake. Chemosphere. doi:10.1016/j.chemosphere.2013.12.051

Khan, F.R., Misra, S.K., Bury, N.R., Smith, B.D., Rainbow, P.S., Luoma, S.N., Valsami-Jones, E., 2015. Inhibition of

potential uptake pathways for silver nanoparticles in the estuarine snail Peringia ulvae. Nanotoxicology 9, 493–

501. doi:10.3109/17435390.2014.948519



Chem. 27, 1825–1851. doi:10.1897/08-090.1

Kozera, B., Rapacz, M., 2013. Reference genes in real-time PCR. J. Appl. Genet. 54, 391–406. doi:10.1007/s13353-

013-0173-x

Kumar, D., Kumari, J., Pakrashi, S., Dalai, S., Raichur, A.M., Sastry, T.P., Mandal, A.B., Chandrasekaran, N.,

Mukherjee, A., 2014. Qualitative toxicity assessment of silver nanoparticles on the fresh water bacterial isolates

and consortium at low level of exposure concentration. Ecotoxicol. Environ. Saf. 108, 152–160.

doi:10.1016/j.ecoenv.2014.05.033




Kutlu, M., Düzen, A., Bayçu, C., Özata, A., 2002. A transmission electron microscope investigation of the effect of

lead acetate on the hepatopancreatic ceca of Gammarus pulex. Environ. Toxicol. Pharmacol. 12, 181–187.

doi:10.1016/S1382-6689(02)00036-4

Kuznetsova, G.P., Larina, O.V., Petushkova, N.A., Kisrieva, Y.S., Samenkova, N.F., Trifonova, O.P., Karuzina, I.I.,

Ipatova, O.M., Zolotaryov, K.V., Romashova, Y.A., Lisitsa, A.V., 2014. Effects of Fullerene C60 on Proteomic Profile

of Danio Rerio Fish Embryos. Bull. Exp. Biol. Med. 156, 694–698. doi:10.1007/s10517-014-2427-y

Kvítek, L., Panáček, A., Soukupová, J., Kolář, M., Večeřová, R., Prucek, R., Holecová, M., Zbořil, R., 2008. Effect of

Surfactants and Polymers on Stability and Antibacterial Activity of Silver Nanoparticles (NPs). J. Phys. Chem. C 112,

5825–5834. doi:10.1021/jp711616v

Kwok, K.W.H., Auffan, M., Badireddy, A.R., Nelson, C.M., Wiesner, M.R., Chilkoti, A., Liu, J., Marinakos, S.M.,

Hinton, D.E., 2012. Uptake of silver nanoparticles and toxicity to early life stages of Japanese medaka (Oryzias

latipes): Effect of coating materials. Aquat. Toxicol. 120–121, 59–66. doi:10.1016/j.aquatox.2012.04.012

Lacave, J.M., Fanjul, Á., Bilbao, E., Gutierrez, N., Barrio, I., Arostegui, I., Cajaraville, M.P., Orbea, A., 2017. Acute

toxicity, bioaccumulation and effects of dietary transfer of silver from brine shrimp exposed to PVP/PEI-coated

silver nanoparticles to zebrafish. Comp. Biochem. Physiol. Part C Toxicol. Pharmacol.

doi:10.1016/j.cbpc.2017.03.008



doi:10.1016/j.mrgentox.2010.04.025

Lacroix, C., Coquillé, V., Guyomarch, J., Auffret, M., Moraga, D., 2014. A selection of reference genes and early-

warning mRNA biomarkers for environmental monitoring using Mytilus spp. as sentinel species. Mar. Pollut. Bull.

225

86, 304–313. doi:10.1016/j.marpolbul.2014.06.049

Ladewig, V., Jungmann, D., Köhler, H.-R., Schirling, M., Triebskorn, R., Nagel, R., 2006. Population Structure and

Dynamics of Gammarus fossarum (Amphipoda) Upstream and Downstream from Effluents of Sewage Treatment

Plants. Arch. Environ. Contam. Toxicol. 50, 370–383. doi:10.1007/s00244-005-7039-0

Lagadic, L., 2002. Biomarkers: Useful tools for the monitoring of aquatic environments. Rev. Med. Veterinaire 153,

581–588.

Lagadic, L., Caquet, T., Ramade, F., 1994. The role of biomarkers in environmental assessment (5). Invertebrate

populations and communities. Ecotoxicology 3, 193–208. doi:10.1007/BF00117084

Lagrue, C., Wattier, R., Galipaud, M., Gauthey, Z., Rullmann, J.-P., Dubreuil, C., Rigaud, T., Bollache, L., 2014.

Confrontation of cryptic diversity and mate discrimination within Gammarus pulex and Gammarus fossarum

species complexes. Freshw. Biol. 59, 2555–2570. doi:10.1111/fwb.12453



Larkin, P., Folmar, L.C., Hemmer, M.J., Poston, A.J., Denslow, N.D., 2003. Expression profiling of estrogenic

compounds using a sheepshead minnow cDNA macroarray. EHP Toxicogenomics J. Natl. Inst. Environ. Health Sci.

111, 29–36.

Lebrun, J.D., Geffard, O., Urien, N., François, A., Uher, E., Fechner, L.C., 2015. Seasonal variability and inter-species

comparison of metal bioaccumulation in caged gammarids under urban diffuse contamination gradient:

Implications for biomonitoring investigations. Sci. Total Environ. 511, 501–508.

doi:10.1016/j.scitotenv.2014.12.078

Lebrun, J.D., Perret, M., Geffard, A., Gourlay-Francé, C., 2012. Modelling copper bioaccumulation in Gammarus

pulex and alterations of digestive metabolism. Ecotoxicology 21, 2022–2030. doi:10.1007/s10646-012-0955-7

Lee, B., Duong, C.N., Cho, J., Lee, J., Kim, K., Seo, Y., Kim, P., Choi, K., Yoon, J., 2012. Toxicity of Citrate-Capped

Silver Nanoparticles in Common Carp (Cyprinus carpio). J. Biomed. Biotechnol. 2012. doi:10.1155/2012/262670

Lee, B.-T., Ranville, J.F., 2012. The effect of hardness on the stability of citrate-stabilized gold nanoparticles and

their uptake by Daphnia magna. J. Hazard. Mater. 213–214, 434–439. doi:10.1016/j.jhazmat.2012.02.025

Lee, S.Y., Nam, Y.K., 2016. Evaluation of reference genes for RT-qPCR study in abalone Haliotis discus hannai during

heavy metal overload stress. Fish. Aquat. Sci. 19. doi:10.1186/s41240-016-0022-z

Legay, S., Guerriero, G., André, C., Guignard, C., Cocco, E., Charton, S., Boutry, M., Rowland, O., Hausman, J.-F.,

2016. MdMyb93 is a regulator of suberin deposition in russeted apple fruit skins. New Phytol. 212, 977–991.

doi:10.1111/nph.14170

Legay, S., Guerriero, G., Deleruelle, A., Lateur, M., Evers, D., André, C.M., Hausman, J.-F., 2015. Apple russeting as

seen through the RNA-seq lens: strong alterations in the exocarp cell wall. Plant Mol. Biol. 88, 21–40.

doi:10.1007/s11103-015-0303-4

Leroy, D., Haubruge, E., De Pauw, E., Thomé, J.P., Francis, F., 2010. Development of ecotoxicoproteomics on the

freshwater amphipod Gammarus pulex: identification of PCB biomarkers in glycolysis and glutamate pathways.

Ecotoxicol. Environ. Saf. 73, 343–352. doi:10.1016/j.ecoenv.2009.11.006



226

Libralato, G., 2014. The case of Artemia spp. in nanoecotoxicology. Mar. Environ. Res. 101, 38–43.

doi:10.1016/j.marenvres.2014.08.002

Lignot, J.-H., Spanings-Pierrot, C., Charmantier, G., 2000. Osmoregulatory capacity as a tool in monitoring the

physiological condition and the effect of stress in crustaceans. Aquaculture 191, 209–245. doi:10.1016/S0044-

8486(00)00429-4

Lincoln, R.J., 1979. British Marine Amphipoda: Gammaridea. - -With 290 figs., 658 pp. London: British Museum

(Natural History) 1979. ISBN 0 565 00818 8. £ 50.00. Int. Rev. Gesamten Hydrobiol. Hydrogr. 66, 446–447.

doi:10.1002/iroh.19810660336

Lipinski, B., 2011. Hydroxyl Radical and Its Scavengers in Health and Disease [WWW Document]. Oxid. Med. Cell.

Longev. doi:10.1155/2011/809696

Lockwood, A.P.M., Inman, C.B.E., 1973. The blood volume of some amphipod crustaceans in relation to the salinity

of the environment they inhabit. Comp. Biochem. Physiol. A Physiol. 44, 935–941. doi:10.1016/0300-

9629(73)90158-8

López-Landavery, E.A., Portillo-López, A., Gallardo-Escárate, C., Del Río-Portilla, M.A., 2014. Selection of reference

genes as internal controls for gene expression in tissues of red abalone Haliotis rufescens (Mollusca,

Vetigastropoda; Swainson, 1822). Gene 549, 258–265. doi:10.1016/j.gene.2014.08.002



Lowry, G.V., Gregory, K.B., Apte, S.C., Lead, J.R., 2012. Transformations of Nanomaterials in the Environment.





Luoma, S.N., Rainbow, P.S., 2005. Why Is Metal Bioaccumulation So Variable? Biodynamics as a Unifying Concept.


Lynch, I., Dawson, K.A., 2008. Protein-nanoparticle interactions. Nano Today 3, 40–47. doi:10.1016/S1748-

0132(08)70014-8

Ma, H., Williams, P.L., Diamond, S.A., 2013. Ecotoxicity of manufactured ZnO nanoparticles – A review. Environ.


Mackevica, A., Skjolding, L.M., Gergs, A., Palmqvist, A., Baun, A., 2015. Chronic toxicity of silver nanoparticles to

Daphnia magna under different feeding conditions. Aquat. Toxicol. 161, 10–16.

doi:10.1016/j.aquatox.2015.01.023



72, 349–364. doi:10.1111/j.1469-185X.1997.tb00017.x

Makama, S., Piella, J., Undas, A., Dimmers, W.J., Peters, R., Puntes, V.F., van den Brink, N.W., 2016. Properties of

silver nanoparticles influencing their uptake in and toxicity to the earthworm Lumbricus rubellus following

exposure in soil. Environ. Pollut. doi:10.1016/j.envpol.2016.08.016

Mangeot-Peter, L., Legay, S., Hausman, J.-F., Esposito, S., Guerriero, G., 2016. Identification of Reference Genes

for RT-qPCR Data Normalization in Cannabis sativa Stem Tissues. Int. J. Mol. Sci. 17, 1556.

227

doi:10.3390/ijms17091556

Manke, A., Wang, L., Rojanasakul, Y., 2013. Mechanisms of Nanoparticle-Induced Oxidative Stress and Toxicity

[WWW Document]. BioMed Res. Int. doi:10.1155/2013/942916

Mann, R.M., Grosell, M., Bianchini, A., Wood, C.M., 2004. Biologically incorporated dietary silver has no

ionoregulatory effects in American red crayfish (Procambarus clarkii). Environ. Toxicol. Chem. 23, 388–395.

doi:10.1897/02-572

Mantel, L.H., Farmer, L.L., 1983. 2 - Osmotic and Ionic Regulation, in: Linda Mantel (Ed.), Internal Anatomy and

Physiological Regulation. Academic Press, pp. 53–161.

Manzo, S., Miglietta, M.L., Rametta, G., Buono, S., Di Francia, G., 2013. Toxic effects of ZnO nanoparticles towards

marine algae Dunaliella tertiolecta. Sci. Total Environ. 445–446, 371–376. doi:10.1016/j.scitotenv.2012.12.051

Martinez, A.-S., Charmantier, G., Compère, P., Charmantier-Daures, M., 2005. Branchial chamber tissues in two

caridean shrimps: the epibenthic Palaemon adspersus and the deep-sea hydrothermal Rimicaris exoculata. Tissue

Cell 37, 153–165. doi:10.1016/j.tice.2004.12.004

McCahon, C.P., Pascoe, D., 1988. Use of Gammarus pulex (L.) in safety evaluation tests: Culture and selection of a

sensitive life stage. Ecotoxicol. Environ. Saf. 15, 245–252. doi:10.1016/0147-6513(88)90078-4





Guérold, F., Audinot, J.-N., Giambérini, L., Gutleb, A.C., 2016a. Gammarus fossarum (Crustacea, Amphipoda) as a

model organism to study the effects of silver nanoparticles. Sci. Total Environ. doi:10.1016/j.scitotenv.2016.06.068


Guérold, F., Audinot, J.-N., Giambérini, L., Gutleb, A.C., 2016b. Gammarus fossarum (Crustacea, Amphipoda) as a

model organism to study the effects of silver nanoparticles. Sci. Total Environ. 566–567, 1649–1659.

doi:10.1016/j.scitotenv.2016.06.068

Meijering, M.P.D., 1991. Lack of oxygen and low pH as limiting factors for Gammarus in Hessian brooks and rivers.

Hydrobiologia 223, 159–169. doi:10.1007/BF00047637

Michalec, F.-G., Holzner, M., Barras, A., Lacoste, A.-S., Brunet, L., Lee, J.-S., Slomianny, C., Boukherroub, R., Souissi,

S., 2017. Short-term exposure to gold nanoparticle suspension impairs swimming behavior in a widespread

calanoid copepod. Environ. Pollut. 228, 102–110. doi:10.1016/j.envpol.2017.04.084

Moore, M.N., 2006. Do nanoparticles present ecotoxicological risks for the health of the aquatic environment?

Environ. Int., Environmental Risk Management - the State of the Art 32, 967–976.

doi:10.1016/j.envint.2006.06.014

Moreno-Garrido, I., Pérez, S., Blasco, J., 2015. Toxicity of silver and gold nanoparticles on marine microalgae. Mar.

Environ. Res., Particles in the Oceans: Implication for a safe marine environment 111, 60–73.

doi:10.1016/j.marenvres.2015.05.008

Morgan, I.J., Henry, R.P., Wood, C.M., 1997. The mechanism of acute silver nitrate toxicity in freshwater rainbow

trout (Oncorhynchus mykiss) is inhibition of gill Na+ and Cl−1 transport. Aquat. Toxicol. 38, 145–163.

doi:10.1016/S0166-445X(96)00835-1

Mueller, N.C., Nowack, B., 2008. Exposure Modeling of Engineered Nanoparticles in the Environment. Environ. Sci.

228

Technol. 42, 4447–4453. doi:10.1021/es7029637

Müller, J., 2000. Mitochondrial DNA Variation and the Evolutionary History of Cryptic Gammarus fossarum Types.

Mol. Phylogenet. Evol. 15, 260–268. doi:10.1006/mpev.1999.0740

Müller, J., 1998. Genetic population structure of two cryptic Gammarus fossarum types across a contact zone. J.

Evol. Biol. 11, 79–101. doi:10.1046/j.1420-9101.1998.11010079.x

Nair, P.M.G., Park, S.Y., Choi, J., 2013. Evaluation of the effect of silver nanoparticles and silver ions using stress

responsive gene expression in Chironomus riparius. Chemosphere 92, 592–599.




Nebeker, A.V., Dominguez, S.E., Chapman, G.A., Onjukka, S.T., Stevens, D.G., 1992. Effects of low dissolved oxygen

on survival, growth and reproduction of Daphnia, Hyalella and Gammarus. Environ. Toxicol. Chem. 11, 373–379.

doi:10.1002/etc.5620110311

Nel, A., Xia, T., Mädler, L., Li, N., 2006. Toxic Potential of Materials at the Nanolevel. Science 311, 622–627.

doi:10.1126/science.1114397

Newton, K.M., Puppala, H.L., Kitchens, C.L., Colvin, V.L., Klaine, S.J., 2013. Silver nanoparticle toxicity to Daphnia

magna is a function of dissolved silver concentration. Environ. Toxicol. Chem. 32, 2356–2364.

doi:10.1002/etc.2300

Nowack, B., Ranville, J.F., Diamond, S., Gallego-Urrea, J.A., Metcalfe, C., Rose, J., Horne, N., Koelmans, A.A., Klaine,

S.J., 2012. Potential scenarios for nanomaterial release and subsequent alteration in the environment. Environ.


Oberdörster, E., Zhu, S., Blickley, T.M., McClellan-Green, P., Haasch, M.L., 2006. Ecotoxicology of carbon-based

engineered nanoparticles: Effects of fullerene (C60) on aquatic organisms. Carbon 44, 1112–1120.

doi:10.1016/j.carbon.2005.11.008

Oliver, A.L.-S., Croteau, M.-N., Stoiber, T.L., Tejamaya, M., Römer, I., Lead, J.R., Luoma, S.N., 2014. Does water

chemistry affect the dietary uptake and toxicity of silver nanoparticles by the freshwater snail Lymnaea stagnalis?


Omondi, B.A., Latorre-Estivalis, J.M., Rocha Oliveira, I.H., Ignell, R., Lorenzo, M.G., 2015. Evaluation of reference

genes for insect olfaction studies. Parasit. Vectors 8, 243. doi:10.1186/s13071-015-0862-x

Oukarroum, A., Bras, S., Perreault, F., Popovic, R., 2012. Inhibitory effects of silver nanoparticles in two green algae,

Chlorella vulgaris and Dunaliella tertiolecta. Ecotoxicol. Environ. Saf. 78, 80–85. doi:10.1016/j.ecoenv.2011.11.012

Owens, T.G., King, F.D., 1975. The measurement of respiratory electron-transport-system activity in marine

zooplankton. Mar. Biol. 30, 27–36. doi:10.1007/BF00393750

Paglia, D.E., Valentine, W.N., 1967. Studies on the quantitative and qualitative characterization of erythrocyte

glutathione peroxidase. J. Lab. Clin. Med. 70, 158–169.

Pan, J.-F., Buffet, P.-E., Poirier, L., Amiard-Triquet, C., Gilliland, D., Joubert, Y., Pilet, P., Guibbolini, M., Risso de

Faverney, C., Roméo, M., Valsami-Jones, E., Mouneyrac, C., 2012. Size dependent bioaccumulation and ecotoxicity

of gold nanoparticles in an endobenthic invertebrate: The Tellinid clam Scrobicularia plana. Environ. Pollut. 168,

37–43. doi:10.1016/j.envpol.2012.03.051

229

Park, Chung, J., Colman, B.P., Matson, C.W., Kim, Y., Lee, B.-C., Kim, P.-J., Choi, K., Choi, J., 2015a. Ecotoxicity of

bare and coated silver nanoparticles in the aquatic midge, Chironomus riparius. Environ. Toxicol. Chem. 34, 2023–

2032. doi:10.1002/etc.3019

Park, Woodhall, J., Ma, G., Veinot, J.G.., Boxall, A.B.A., 2015b. Do particle size and surface functionality affect

uptake and depuration of gold nanoparticles by aquatic invertebrates? Environ. Toxicol. Chem. 34, 850–859.

doi:10.1002/etc.2868

Pequeux, A., 1995. OSMOTIC REGULATION IN CRUSTACEANS. J. Crustac. Biol. 15, 1–60.

doi:10.1163/193724095X00578

Petersen, E.J., Henry, T.B., Zhao, J., MacCuspie, R.I., Kirschling, T.L., Dobrovolskaia, M.A., Hackley, V., Xing, B.,

White, J.C., 2014. Identification and Avoidance of Potential Artifacts and Misinterpretations in Nanomaterial

Ecotoxicity Measurements. Environ. Sci. Technol. 48, 4226–4246. doi:10.1021/es4052999

Pfaffl, M.W., Tichopad, A., Prgomet, C., Neuvians, T.P., 2004. Determination of stable housekeeping genes,

differentially regulated target genes and sample integrity: BestKeeper – Excel-based tool using pair-wise

correlations. Biotechnol. Lett. 26, 509–515. doi:10.1023/B:BILE.0000019559.84305.47

Pham, C.H., Yi, J., Gu, M.B., 2012. Biomarker gene response in male Medaka (Oryzias latipes) chronically exposed

to silver nanoparticle. Ecotoxicol. Environ. Saf. 78, 239–245. doi:10.1016/j.ecoenv.2011.11.034

Piccinno, F., Gottschalk, F., Seeger, S., Nowack, B., 2012. Industrial production quantities and uses of ten

engineered nanomaterials in Europe and the world. J. Nanoparticle Res. 14, 1–11. doi:10.1007/s11051-012-1109-

9

Piscart, C., Bollache, L., 2012. Crustacés amphipodes de surface : gammares d’eau douce., Introduction pratique à

la systématique des organismes des eaux continentales de France. Association Française de Limnologie.

Plusquin, M., DeGheselle, O., Cuypers, A., Geerdens, E., Roten, A.V., Artois, T., Smeets, K., 2012. Reference genes

for qPCR assays in toxic metal and salinity stress in two flatworm model organisms. Ecotoxicology 21, 475–484.

doi:10.1007/s10646-011-0809-8

Pöckl, M., Webb, B.W., Sutcliffe, D.W., 2003. Life history and reproductive capacity of Gammarus fossarum and G.

roeseli (Crustacea: Amphipoda) under naturally fluctuating water temperatures: a simulation study. Freshw. Biol.

48, 53–66. doi:10.1046/j.1365-2427.2003.00967.x

Poynton, H.C., Lazorchak, J.M., Impellitteri, C.A., Blalock, B.J., Rogers, K., Allen, H.J., Loguinov, A., Heckman, J.L.,

Govindasmawy, S., 2012. Toxicogenomic Responses of Nanotoxicity in Daphnia magna Exposed to Silver Nitrate

and Coated Silver Nanoparticles. Environ. Sci. Technol. 46, 6288–6296. doi:10.1021/es3001618

Rainbow, P.S., 1997. Ecophysiology of Trace Metal Uptake in Crustaceans. Estuar. Coast. Shelf Sci. 44, 169–175.

doi:10.1006/ecss.1996.0208

Rainbow, P.S., 1995. Physiology, physicochemistry and metal uptake—A crustacean perspective. Mar. Pollut. Bull.

31, 55–59. doi:10.1016/0025-326X(95)00005-8

Rainville, L.-C., Carolan, D., Varela, A.C., Doyle, H., Sheehan, D., 2014. Proteomic evaluation of citrate-coated silver

nanoparticles toxicity in Daphnia magna. Analyst 139, 1678–1686. doi:10.1039/C3AN02160B

Rauscher, H., Roebben, G., Boix Sanfeliu, A., Emmons, H., Gibson, N., Koeber, R., Linsinger, T., Rasmussen, K., Riego

Sintes, J., Sokull-Klüttgen, B., Stamm, H., 2015. Towards a review of the EC Recommendation for a definition of the

term “nanomaterial” Part 3 Scientific-technical evaluation of options to clarify the definition and to facilitate its

implementation. doi:10.2788/678452

230

Regoli, F., Gorbi, S., Frenzilli, G., Nigro, M., Corsi, I., Focardi, S., Winston, G.., 2002. Oxidative stress in

ecotoxicology: from the analysis of individual antioxidants to a more integrated approach. Mar. Environ. Res. 54,

419–423. doi:10.1016/S0141-1136(02)00146-0




Revel, M., Châtel, A., Mouneyrac, C., 2017. Omics tools: New challenges in aquatic nanotoxicology? Aquat. Toxicol.

doi:10.1016/j.aquatox.2017.10.005

Ribeiro, F., Van Gestel, C.A.M., Pavlaki, M.D., Azevedo, S., Soares, A.M.V.M., Loureiro, S., 2017. Bioaccumulation

of silver in Daphnia magna: Waterborne and dietary exposure to nanoparticles and dissolved silver. Sci. Total


Römer, I., White, T.A., Baalousha, M., Chipman, K., Viant, M.R., Lead, J.R., 2011. Aggregation and dispersion of

silver nanoparticles in exposure media for aquatic toxicity tests. J. Chromatogr. A 1218, 4226–4233.

doi:10.1016/j.chroma.2011.03.034

Rosas, C., Sanchez, A., Escobar, E., Soto, L., Bolongaro-Crevenna, A., 1992. Daily variations of oxygen consumption

and glucose hemolymph level related to morphophysiological and ecological adaptations of crustacea. Comp.

Biochem. Physiol. A Physiol. 101, 323–328. doi:10.1016/0300-9629(92)90540-7

Sakka, Y., Skjolding, L.M., Mackevica, A., Filser, J., Baun, A., 2016. Behavior and chronic toxicity of two differently

stabilized silver nanoparticles to Daphnia magna. Aquat. Toxicol. 177, 526–535.

doi:10.1016/j.aquatox.2016.06.025

Sasidharan, A., Riviere, J.E., Monteiro-Riviere, N.A., 2015. Gold and silver nanoparticle interactions with human

proteins: impact and implications in biocorona formation. J. Mater. Chem. B 3, 2075–2082.

doi:10.1039/C4TB01926A

Sattler, T., Bontadina, F., Hirzel, A.H., Arlettaz, R., 2007. Ecological niche modelling of two cryptic bat species calls

for a reassessment of their conservation status. J. Appl. Ecol. 44, 1188–1199. doi:10.1111/j.1365-

2664.2007.01328.x

Savolainen, K., Backman, U., Brouwer, D., Fadeel, B., Fernandes, T., Kuhlbusch, T.A.J., Landsiedel, R., Lynch, I.,

Pylkkanen, L., Ahluwalia, A., Jukka, A., Aitken, R., Alenius, H., Asbach, C., Athersuch, T., Bergamaschi, E., Berges,

M., Bessems, J., Bleeker, E.A.J., Özgüz, V., 2013. Nanosafety in Europe 2015-2025: Towards Safe and Sustainable

Nanomaterials and Nanotechnology Innovations. doi:10.13140/2.1.3084.8969

Scheepmaker, M., van Dalfsen, 1989. Genetic differentiation in Gammarus fossarum and G. caparti (Crustacea,

Amphipoda) with reference to G. pulex pulex in northwestern Europe. Bijdr. Tot Dierkd. 59, 127–139.

Schmitz, E.H., Scherrey, P.M., 1983. Digestive anatomy of Halella azteca (Crustacea, Amphipoda). J. Morphol. 175,

91–100. doi:10.1002/jmor.1051750109

Schopf, F.H., Biebl, M.M., Buchner, J., 2017. The HSP90 chaperone machinery. Nat. Rev. Mol. Cell Biol. 18, 345–

360. doi:10.1038/nrm.2017.20

Schultz, A.G., Boyle, D., Chamot, D., Ong, K.J., Wilkinson, K.J., McGeer, J.C., Sunahara, G., Goss, G.G., 2014. Aquatic

toxicity of manufactured nanomaterials: challenges and recommendations for future toxicity testing. Environ.

Chem. 11, 207–226. doi:10.1071/EN13221

Schultz, A.G., Ong, K.J., MacCormack, T., Ma, G., Veinot, J.G.C., Goss, G.G., 2012. Silver Nanoparticles Inhibit

231

Sodium Uptake in Juvenile Rainbow Trout (Oncorhynchus mykiss). Environ. Sci. Technol. 46, 10295–10301.

doi:10.1021/es3017717

Scott, G.R., Sloman, K.A., 2004. The effects of environmental pollutants on complex fish behaviour: integrating

behavioural and physiological indicators of toxicity. Aquat. Toxicol. 68, 369–392.

doi:10.1016/j.aquatox.2004.03.016




Sharma, N., Rather, M.A., Ajima, M.N.O., Gireesh-Babu, P., Kumar, K., Sharma, R., 2016. Assessment of DNA

damage and molecular responses in Labeo rohita (Hamilton, 1822) following short-term exposure to silver

nanoparticles. Food Chem. Toxicol. 96, 122–132. doi:10.1016/j.fct.2016.06.020

Shaw, J.R., Colbourne, J.K., Davey, J.C., Glaholt, S.P., Hampton, T.H., Chen, C.Y., Folt, C.L., Hamilton, J.W., 2007.

Gene response profiles for Daphnia pulex exposed to the environmental stressor cadmium reveals novel

crustacean metallothioneins. BMC Genomics 8, 1–20. doi:10.1186/1471-2164-8-477

Shepard, J.L., Olsson, B., Tedengren, M., Bradley, B.P., 2000. Protein expression signatures identified in Mytilus

edulis exposed to PCBs, copper and salinity stress. Mar. Environ. Res. 50, 337–340. doi:10.1016/S0141-

1136(00)00065-9

Shi, J., Xu, B., Sun, X., Ma, C., Yu, C., Zhang, H., 2013. Light induced toxicity reduction of silver nanoparticles to

Tetrahymena Pyriformis: Effect of particle size. Aquat. Toxicol. 132–133, 53–60.

doi:10.1016/j.aquatox.2013.02.001

Shimek, R.L., 2008. Amphipods [WWW Document]. URL http://reefkeeping.com/issues/2004-09/rs/index.php

(accessed 8.25.17).

Silva, T., Pokhrel, L.R., Dubey, B., Tolaymat, T.M., Maier, K.J., Liu, X., 2014. Particle size, surface charge and

concentration dependent ecotoxicity of three organo-coated silver nanoparticles: Comparison between general

linear model-predicted and observed toxicity. Sci. Total Environ. 468–469, 968–976.

doi:10.1016/j.scitotenv.2013.09.006

Silver, N., Best, S., Jiang, J., Thein, S.L., 2006. Selection of housekeeping genes for gene expression studies in human

reticulocytes using real-time PCR. BMC Mol. Biol. 7, 33. doi:10.1186/1471-2199-7-33

Skjolding, L.M., Ašmonaitė, G., Jølck, R.I., Andresen, T.L., Selck, H., Baun, A., Sturve, J., 2017. An assessment of the

importance of exposure routes to the uptake and internal localisation of fluorescent nanoparticles in zebrafish

(Danio rerio), using light sheet microscopy. Nanotoxicology 11, 351–359. doi:10.1080/17435390.2017.1306128

Skjolding, L.M., Kern, K., Hjorth, R., Hartmann, N., Overgaard, S., Ma, G., Veinot, J.G.C., Baun, A., 2014. Uptake and

depuration of gold nanoparticles in Daphnia magna. Ecotoxicology 23, 1172–1183. doi:10.1007/s10646-014-1259-

x



doi:10.1016/j.aquatox.2016.10.004



158, 1288–1295. doi:10.1016/j.envpol.2010.01.022

232




128.

Sutcliffe, D.W., 1984. Quantitative aspects of oxygen uptake by Gammarus (Crustacea, Amphipoda): a critical

review. Freshw. Biol. 14, 443–489. doi:10.1111/j.1365-2427.1984.tb00168.x

Tachet, H., 2000. Invertébrés d’eau douce: systématique, biologie, écologie. CNRS Editions.

Tedesco, S., Doyle, H., Blasco, J., Redmond, G., Sheehan, D., 2010. Oxidative stress and toxicity of gold

nanoparticles in Mytilus edulis. Aquat. Toxicol. 100, 178–186. doi:10.1016/j.aquatox.2010.03.001

Tejamaya, M., Römer, I., Merrifield, R.C., Lead, J.R., 2012. Stability of Citrate, PVP, and PEG Coated Silver

Nanoparticles in Ecotoxicology Media. Environ. Sci. Technol. 46, 7011–7017. doi:10.1021/es2038596

The European Commision, 2011. Commission Recommendation of 18 October 2011 on the definition of

nanomaterialText with EEA relevance - commission-recommendation-on-the-definition-of-nanomater-

18102011_en.pdf [WWW Document]. URL

https://ec.europa.eu/research/industrial_technologies/pdf/policy/commission-recommendation-on-the-

definition-of-nanomater-18102011_en.pdf (accessed 8.30.17).

Thompson, L.B., Carfagno, G.L.F., Andresen, K., Sitton, A.J., Bury, T., Lee, L.L., Lerner, K.T., Fong, P.P., 2017.

Differential uptake of gold nanoparticles by 2 species of tadpole, the wood frog (Lithobates sylvaticus) and the

bullfrog (Lithobates catesbeianus). Environ. Toxicol. Chem. n/a-n/a. doi:10.1002/etc.3909

Tiede, K., Hassellöv, M., Breitbarth, E., Chaudhry, Q., Boxall, A.B.A., 2009. Considerations for environmental fate

and ecotoxicity testing to support environmental risk assessments for engineered nanoparticles. J. Chromatogr. A,

Tools for the REACH Programme - analytical methods for the evaluation of industrial contaminants 1216, 503–509.

doi:10.1016/j.chroma.2008.09.008

Tolaymat, T.M., El Badawy, A.M., Genaidy, A., Scheckel, K.G., Luxton, T.P., Suidan, M., 2010. An evidence-based

environmental perspective of manufactured silver nanoparticle in syntheses and applications: A systematic review

and critical appraisal of peer-reviewed scientific papers. Sci. Total Environ. 408, 999–1006.

doi:10.1016/j.scitotenv.2009.11.003

Trapp, J., Armengaud, J., Pible, O., Gaillard, J.-C., Abbaci, K., Habtoul, Y., Chaumot, A., Geffard, O., 2014. Proteomic

Investigation of Male Gammarus fossarum, a Freshwater Crustacean, in Response to Endocrine Disruptors. J.

Proteome Res. doi:10.1021/pr500984z

Trapp, J., 2015. Approche protéomiques pour le développement de biomarqurs chez l’amphipode d’eau douce Gammarus fossarum: découverte et caractérisation de protéines impliquées dans la fonction reproductrice. Université Claude Bernard Lyon 1.

US Geological Survey, 2016. Mineral Commodity Summaries 2016 - mcs-2016-silve.pdf [WWW Document]. URL

http://minerals.usgs.gov/minerals/pubs/commodity/silver/mcs-2016-silve.pdf (accessed 3.7.16).

Väinölä, R., Witt, J.D.S., Grabowski, M., Bradbury, J.H., Jazdzewski, K., Sket, B., 2008. Global diversity of amphipods

(Amphipoda; Crustacea) in freshwater. SpringerLink 241–255. doi:10.1007/978-1-4020-8259-7_27

Vale, G., Franco, C., Diniz, M.S., Santos, M.M.C. dos, Domingos, R.F., 2014. Bioavailability of cadmium and

biochemical responses on the freshwater bivalve Corbicula fluminea – the role of TiO2 nanoparticles. Ecotoxicol.

233


Vale, G., Mehennaoui, K., Cambier, S., Libralato, G., Jomini, S., Domingos, R.F., 2016. Manufactured nanoparticles

in the aquatic environment-biochemical responses on freshwater organisms: A critical overview. Aquat. Toxicol.

170, 162–174. doi:10.1016/j.aquatox.2015.11.019

Vance, M.E., Kuiken, T., Vejerano, E.P., McGinnis, S.P., Hochella, M.F., Rejeski, D., Hull, M.S., 2015. Nanotechnology

in the real world: Redeveloping the nanomaterial consumer products inventory. Beilstein J. Nanotechnol. 6, 1769–

1780. doi:10.3762/bjnano.6.181

Vandesompele, J., De Preter, K., Pattyn, F., Poppe, B., Van Roy, N., De Paepe, A., Speleman, F., 2002. Accurate

normalization of real-time quantitative RT-PCR data by geometric averaging of multiple internal control genes.

Genome Biol. 3, research0034. doi:10.1186/gb-2002-3-7-research0034



Vellinger, C., Felten, V., Sornom, P., Rousselle, P., Beisel, J.-N., Usseglio-Polatera, P., 2012a. Behavioural and






doi:10.1016/j.aquatox.2013.05.010

Vellinger, C., Parant, M., Rousselle, P., Immel, F., Wagner, P., Usseglio-Polatera, P., 2012b. Comparison of arsenate

and cadmium toxicity in a freshwater amphipod (Gammarus pulex). Environ. Pollut. 160, 66–73.

doi:10.1016/j.envpol.2011.09.002

Vindimian, É., Garric, J., Flammarion, P., Thybaud, É., Babut, M., 1999. An index of effluent aquatic toxicity designed

by partial least squares regression, using acute and chronic tests and expert judgements. Environ. Toxicol. Chem.

18, 2386–2391. doi:10.1002/etc.5620181037

Vindimian, E., Robaut, C., Fillion, G., 1983. A method for cooperative or noncooperative binding studies using

nonlinear regression analysis on a microcomputer. J. Appl. Biochem. 5, 261–268.

Vioque-Fernández, A., Alves de Almeida, E., López-Barea, J., 2009. Assessment of Doñana National Park

contamination in Procambarus clarkii: Integration of conventional biomarkers and proteomic approaches. Sci.


Voellmy, R., Boellmann, F., 2007. Chaperone Regulation of the Heat Shock Protein Response 89–99.

doi:10.1007/978-0-387-39975-1_9

Völker, C., Kämpken, I., Boedicker, C., Oehlmann, J., Oetken, M., 2015. Toxicity of silver nanoparticles and ionic

silver: Comparison of adverse effects and potential toxicity mechanisms in the freshwater clam Sphaerium

corneum. Nanotoxicology 9, 677–685. doi:10.3109/17435390.2014.963723

Volland, M., Blasco, J., Hampel, M., 2017. Validation of reference genes for RT-qPCR in marine bivalve

ecotoxicology: Systematic review and case study using copper treated primary Ruditapes philippinarum

hemocytes. Aquat. Toxicol. 185, 86–94. doi:10.1016/j.aquatox.2017.01.003

Volland, M., Hampel, M., Martos-Sitcha, J.A., Trombini, C., Martínez-Rodríguez, G., Blasco, J., 2015. Citrate gold

nanoparticle exposure in the marine bivalve Ruditapes philippinarum: uptake, elimination and oxidative stress

234

response. Environ. Sci. Pollut. Res. 22, 17414–17424. doi:10.1007/s11356-015-4718-x

Wang, L.-F., Habibul, N., He, D.-Q., Li, W.-W., Zhang, X., Jiang, H., Yu, H.-Q., 2015. Copper release from copper

nanoparticles in the presence of natural organic matter. Water Res. 68, 12–23. doi:10.1016/j.watres.2014.09.031

Wang, Z., von dem Bussche, A., Kabadi, P.K., Kane, A.B., Hurt, R.H., 2013. Biological and environmental

transformations of copper-based nanomaterials. ACS Nano 7, 8715–8727. doi:10.1021/nn403080y

Webb, Bartlein, and P.J., 1992. Global Changes During the Last 3 Million Years: Climatic Controls and Biotic

Responses. Annu. Rev. Ecol. Syst. 23, 141–173. doi:10.1146/annurev.es.23.110192.001041

Webb, N.A., Wood, C.M., 1998. Physiological analysis of the stress response associated with acute silver nitrate

exposure in freshwater rainbow trout (Oncorhynchus mykiss). Environ. Toxicol. Chem. 17, 579–588.

doi:10.1002/etc.5620170408

Weiss, M., Macher, J.N., Seefeldt, M.A., Leese, F., 2014. Molecular evidence for further overlooked species within

the Gammarus fossarum complex (Crustacea: Amphipoda). Hydrobiologia 721, 165–184. doi:10.1007/s10750-013-

1658-7

Wepener, V., van Dyk, C., Bervoets, L., O’Brien, G., Covaci, A., Cloete, Y., 2011. An assessment of the influence of

multiple stressors on the Vaal River, South Africa. Phys. Chem. Earth Parts ABC, 11th WaterNet/WARFSA/GWP-SA

Symposium: IWRM for National and Regional Integration through Science, Policy and Practice 36, 949–962.

doi:10.1016/j.pce.2011.07.075

Westram, A.M., Jokela, J., Baumgartner, C., Keller, I., 2011. Spatial Distribution of Cryptic Species Diversity in

European Freshwater Amphipods (Gammarus fossarum) as Revealed by Pyrosequencing. PLoS ONE 6, e23879.

doi:10.1371/journal.pone.0023879

Westram, A.M., Jokela, J., Keller, I., 2010. Isolation and characterization of ten polymorphic microsatellite markers

for three cryptic Gammarus fossarum (Amphipoda) species. Conserv. Genet. Resour. 2, 401–404.

doi:10.1007/s12686-010-9287-1

Whitehurst, I.T., Lindsey, B.I., 1990. The impact of organic enrichment on the benthic macroinvertebrate

communities of a lowland river. Water Res. 24, 625–630. doi:10.1016/0043-1354(90)90195-C



Xie, F., Xiao, P., Chen, D., Xu, L., Zhang, B., 2012. miRDeepFinder: a miRNA analysis tool for deep sequencing of

plant small RNAs. Plant Mol. Biol. doi:10.1007/s11103-012-9885-2

Xuereb, B., Bezin, L., Chaumot, A., Budzinski, H., Augagneur, S., Tutundjian, R., Garric, J., Geffard, O., 2011.

Vitellogenin-like gene expression in freshwater amphipod Gammarus fossarum (Koch, 1835): functional

characterization in females and potential for use as an endocrine disruption biomarker in males. Ecotoxicology 20,

1286–1299. doi:10.1007/s10646-011-0685-2



doi:10.1016/j.scitotenv.2015.12.150

Zavala, E., Reyes, D., Deerenberg, R., Vidal, R., 2017. Selection of reference genes for microRNA analysis associated

to early stress response to handling and confinement in Salmo salar. Sci. Rep. 7, 1756. doi:10.1038/s41598-017-

01970-3


235


Zhang, M.F., Liu, Q., Jia, G.X., 2016. Reference gene selection for gene expression studies in lily using quantitative

real-time PCR. Genet. Mol. Res. 15. doi:10.4238/gmr.15027982

Zhang, P., Selck, H., Tangaa, S.R., Pang, C., Zhao, B., 2017. Bioaccumulation and effects of sediment-associated

gold- and graphene oxide nanoparticles on Tubifex tubifex. J. Environ. Sci. 51, 138–145.

doi:10.1016/j.jes.2016.08.015

Zhao, C.-M., Wang, W.-X., 2012. Importance of surface coatings and soluble silver in silver nanoparticles toxicity

to Daphnia magna. Nanotoxicology 6, 361–370. doi:10.3109/17435390.2011.579632

Zhao, C.-M., Wang, W.-X., 2010. Biokinetic Uptake and Efflux of Silver Nanoparticles in Daphnia magna. Environ.

Sci. Technol. 44, 7699–7704. doi:10.1021/es101484s

Zimmermann, S., Ruchter, N., Loza, K., Epple, M., Sures, B., 2017. Nanoparticulate versus ionic silver: Behavior in

the tank water, bioaccumulation, elimination and subcellular distribution in the freshwater mussel Dreissena

polymorpha. Environ. Pollut. 222, 251–260. doi:10.1016/j.envpol.2016.12.048

Živić, I., Marković, Z., 2007. Distribution of the Species Gammarus balcanicus and Gammarus fossarum on the

Territory of Serbia (Central Part of the Balkan Peninsula). Crustaceana 80, 67–76. doi:10.2307/20107783

Aquatic Toxicology 170 (2016) 162–174

Contents lists available at ScienceDirect

Aquatic Toxicology

j o ur na l ho me pag e: www.elsev ier .com/ locate /aquatox

Manufactured nanoparticles in the aquatic environment-biochemicalresponses on freshwater organisms: A critical overview

Gonc alo Vale a,b,∗, Kahina Mehennaoui c, Sebastien Cambier c, Giovanni Libralatod,Stéphane Jomini e, Rute F. Domingos a,f

a Centro de Química Estrutural, Instituto Superior Técnico, Universidade de Lisboa, Torre Sul Av. Rovisco Pais, 1049-001 Lisboa, Portugalb Department of Molecular Genetics, University of Texas Southwestern Medical Center, Harry Dallas, TX 75390, USAc Luxembourg Institute of Science and Technology, Environmental Research and Innovation (ERIN) Department, Belvaux, Luxembourgd Department of Environmental Sciences, Informatics and Statistics, University Ca’ Foscari Venice, Via Torino 155, 30172, Mestre, Venice, Italye Laboratoire Interdisciplinaire des Environements Continentaux (LIEC), Université de Lorraine, UMR 7360, Campus Bridoux rue du Général Delestraint,

57070 Metz, Francef Institut de Physique du Globe de Paris, Sorbonne Paris Cité, UMR CNRS 7154, Université Paris Diderot, 75205 Paris Cedex 05, France

a r t i c l e i n f o

Article history:

Received 26 August 2015

Received in revised form

16 November 2015

Accepted 19 November 2015

Available online 23 November 2015

Keywords:

Nanoparticles

Transformations

Freshwater organisms

Nanotoxicity

Reactive oxygen species (ROS)

Anti-oxidant enzymes

a b s t r a c t

The enormous investments in nanotechnology have led to an exponential increase of new manufactured

nano-enabled materials whose impact in the aquatic systems is still largely unknown. Ecotoxicity and

nanosafety studies mostly resulted in contradictory results and generally failed to clearly identify biologi-

cal patterns that could be related specifically to nanotoxicity. Generation of reactive oxygen species (ROS)

is one of the most discussed nanotoxicity mechanism in literature. ROS can induce oxidative stress (OS),

resulting in cyto- and genotoxicity. The ROS overproduction can trigger the induction of anti-oxidant

enzymes such as catalase (CAT), superoxide dismutase (SOD) and glutathione peroxidases (GPx), which

are used as biomarkers of response. A critical overview of the biochemical responses induced by the pres-

ence of NPs on freshwater organisms is performed with a strong interest on indicators of ROS and general

stress. A special focus will be given to the NPs transformations, including aggregation, and dissolution,

in the exposure media and the produced biochemical endpoints.

© 2015 Elsevier B.V. All rights reserved.

Contents

1. Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163

2. NPs transformations in aquatic systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 163

3. Nanotoxicity toward aquatic organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 164

3.1. Generation of ROS. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .164

3.2. Omics endpoints . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165

4. NPs toxicity on freshwater organisms . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165

4.1. Silver NPs (nAg) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 165

4.2. Titanium dioxide NPs (nTiO2) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 166

4.3. Zinc oxide NPs (nZnO) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 167

4.4. Copper oxide NPs (nCuO) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 170

5. General remarks and conclusions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 172

∗ Corresponding author.

E-mail addresses: [email protected] (G. Vale), [email protected] (K. Mehennaoui), [email protected] (S. Cambier), [email protected]

(G. Libralato), [email protected] (S. Jomini), [email protected] (R.F. Domingos).


0166-445X/© 2015 Elsevier B.V. All rights reserved.

dx.doi.org/10.1016/j.aquatox.2015.11.019

http://www.sciencedirect.com/science/journal/0166445X

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G. Vale et al. / Aquatic Toxicology 170 (2016) 162–174 163

1. Introduction

Nanotechnology has emerged as a fast growing sector impactingkey economical fields and providing new engineered nano-enabledproducts, constituted by nanoparticles (NPs), with novel andunique functions that reach the market every day (Bour et al.,2015). NPs are defined as materials with a size between 1 and100 nm on at least one dimension, having unique physicochemi-cal properties differing from their bulk forms due to their greatersurface area to volume ratio. This size related-properties results inlarger reactivity and higher mobility (Rauscher et al., 2014), lead-ing to numerous applications in medical diagnostics, electronics,computers, cosmetics and environmental remediation. The world-wide consumption of NPs is expected to grow from 225,060 metrictons in 2014 to nearly 584,984 metric tons in 2019 represent-ing an annual growth rate of 21.1% (Research, 2015). Althoughimpressive, these numbers are in fact “expected” values obtainedby estimation or modeling. The lack of legislation for nanotech-nologies gives the manufacturers no onus to reveal the real figures,thus, indeed, these predicted values are most probably signifi-cantly higher. The absence of real numbers hinders the predictionof the NPs amount that are actually being released into the envi-ronment (Piccinno et al., 2012). Even though several studies havebeen performed with the goal of modeling NPs environmentalconcentrations (Gottschalk et al., 2013), they should only be con-sidered as guidelines, since they derive from uncertain data aboutthe NPs production (often obtained by surveys to the producers)and extrapolations used to scale up regional to worldwide amounts(Piccinno et al., 2012; Gottschalk et al., 2011; Keller et al., 2013).

When released in natural media NPs will be subjected to adynamic physical and chemical environment that consequentlyresults in different and unknown endpoints far from their pris-tine or as released state. Therefore, environments and humans arenot facing pristine manufactured NPs but rather transformed nano-enabled products, which is factually accepted but so far neglected.In fact, the large majority of the physicochemical and toxicity dataobtained so far was focused on simple nanoscale particles andnot on relevant nano-enabled products. This includes not onlythe NP embedded in the manufactured matrix but also the mate-rials resulting from the interaction with biotic and abiotic (bio)molecules composing the natural systems. To further complicatethe interpretation of the NPs studies, there are two distinct mecha-nisms that should be considered but are not easily differentiated: (i)chemical toxicity by the release of possible ions and/or formationof reactive oxygen species (ROS) (Fu et al., 2014), and (ii) physicalstress or stimuli caused by NPs size, shape and surface properties(Vale et al., 2014; Libralato et al., 2013). These materials are gener-ally associated with cellular perturbations such as ROS generation,and gene expression and proteome profiles alterations. For thesereasons, the NPs escalating production and applications has raisedconcerns about their environmental and human safety, which haveled to large investments in nanosafety-related projects resultingin a considerable amount of data assessing their potential hazard(Savolainen et al., 2013). However, the establishment of relation-ships between bioavailable NP-containing species and the specificbioadverse or biocompatible endpoints is still lacking, mainly since,the effects are NP-dependent and also specie-dependent (Buricet al., 2015).

This work provides an overview of the latest studies on theimpact of NPs onto freshwater ecosystems, considered by many asthe ultimate sink of these particles, with a special focus on (i) NPstransformations and characterization in the different test media,and (ii) toxicological effects such as generation of ROS, genotoxic-ity, metallomic and proteomic changes. This survey is focused onmetallic NPs including nAg, nTiO2, nZnO and nCuO, mostly due tothe great number of studies dedicated to these particles.

2. NPs transformations in aquatic systems

NPs can enter in an aquatic compartment from (i) wastewa-ter treatment plants effluents, (ii) direct use (e.g., application ofNPs-containing paintings on boats), and (iii) deposition from theair compartment. When entering aquatic compartment, NPs willbe exposed to a highly dynamic physical and chemical environ-ment that leads to several transformations that will change theirpristine or as released physicochemical properties (Fig. 1). Thesetransformations, including dissolution, aggregation and sedimen-tation, are dependent on both physicochemical properties of the NP(and nanoforms thereof) and those of the environment into whichthey were released.

Colloidal particles, including organic and inorganic matter,are ubiquitous in the aquatic environment and can be origi-nated from both natural and anthropic sources. These colloidscan strongly interact with NPs, thereby determining their formsover space and time (dynamic speciation), and greatly affectingtheir bioavailability. Thus, the NP will have a specific speciationin each environmental compartment, and this speciation is alwaysdynamic with reaction rates that depend upon the chemical natureand physical sizes of the engineered and natural colloids. Althoughit is clear that dynamic speciation must be considered in order tomake relevant predictions of NPs fate, toxicity and risk, until nowthis critical issue, was mostly neglected (see detailed explanationon Pinheiro and Domingos, 2015).

Dissolution, which is one of the main transformations of metallicNPs such as nZnO, nCuO, and nAg, is mainly due to (i) the formationof partially soluble metal-oxide (Heinlaan et al., 2008; Domingoset al., 2013a; Wang et al., 2015), (ii) the oxidation of the parti-cle constituents (Ma et al., 2014; Wang et al., 2013a; Dale et al.,2013; Lok et al., 2007; Derfus et al., 2004), and (iii) the complexa-tion of the particle constituent metal by complexants present inthe environmental compartment or even in the NPs embeddedmatrix (including the manufactured stabilizers) (Domingos et al.,2013b; Domingos et al., 2014). The sulfidation of the metallic NPscan retard their oxidation and, thus, their dissolution (Ma et al.,2014; Wang et al., 2013a; Dale et al., 2013; Thalmann et al., 2014).This dissolution mechanism results in the release of toxic cations,such that their persistence is reduced but the toxicity is increased.Evidently, complete dissolution of the NPs allows the prediction oftheir impact using already existing models for metal speciation andtoxicity.

Photoreactions can also be important transformations affectingthe NPs coatings, oxidation state, generation of ROS, and persis-tence, which is the case of the innately photoactive TiO2 and ZnOparticles (Hund-Rinke and Simon, 2006; Zhang et al., 2007a).

Aggregation is other critical transformation, which mainly byinteraction with naturally occurring bio- or geomacromoleculesaffect NPs size and surface chemistry. For example, organic mat-ter (OM) provides both charge and steric stabilization (Mohammedet al., 2008; Domingos et al., 2009a) of the NPs, although they mayalso result in bridging flocculation when in presence of multiplecharged cations and anions (Domingos et al., 2010). OM effects arecomplex and difficult to predict, however, is of extremely impor-tance to explore these interactions since, OM concentrations aretypically orders of magnitude higher in concentration than engi-neered NPs, and, thus, likely to substantially modify their propertiesand behaviors. Despite the significance of these interactions, withboth organic and inorganic matter, and to the best of our knowl-edge, no relevant toxicity studies are available.

Dissolution and aggregation are dynamic processes that candecrease the NPs available surface area, thereby decreasing theirreactivity. However, this decrease is dependent on the surface prop-erties, particle number, size distribution, and the fractal dimensionsof the aggregate (Hotze et al., 2010). The NP size will affect its

164 G. Vale et al. / Aquatic Toxicology 170 (2016) 162–174

Fig. 1. Representative chemical and physical transformations of NPs when entering in natural aquatic systems: dissolution, phosphatization, sulfidation, homo- and heteroag-

gregation, and sedimentation. Important constituents with which NPs can interact governing their fate and transport includes hardness cations (e.g., Ca2+ , Mg2+), alkalinity,

phosphate and sulfide anions, pH, dissolved organic carbon (DOC), organic matter (OM) and mineral surfaces (such as iron and manganese oxides, and clays). Legend: blue

circles: engineered NPs; yellow circles: humic substances (HM); brown circles: natural inorganic colloids; blue lines: rigid biopolymers; gray surroundings: representing

sulfidation; Mz+: free metal ion. Adapted from Domingos et al. (2015). For interpretation of the references to color in this figure legend, the reader is referred to the web

version of this article.

bioavailability to the organisms; when aggregates become too largefor direct transport across the cell wall and/or membrane, uptakemay be prevented, whereas partial dissolution, which will lead tosmaller sizes, would facilitate this cellular transport. Since thesetransformations are most often not in equilibrium, they requirereal-time kinetic measurements, limiting the methodology to beused: (i) the storage of whole unfractionated samples for ion anal-ysis may not be possible since the dissolution rate may be fastor not attain the equilibrium during the experimental time, and(ii) the aggregation rate can be fast or the aggregates size distri-bution may not reach equilibrium within the experimental timewindow.

Despite the large number of studies focused on nanotoxicol-ogy, most of them disregard the particles kinetic physicochemicalcharacterization under the exposure conditions, hindering theestablishment of crucial predictive structure-activity relationshipsthat can be used afterwards in the categorization and function forrisk assessment studies. In absence of these realistic studies, twoless constrain conditions were used to select the literature studiesthat will be discussed in this critical overview:

i) studies where the NPs characterization was performed in thesame media as the bioassays.

ii) quantification of the NPs dissolution for studies using metallicNPs with propensity for dissolution such as nAg and nZnO.

3. Nanotoxicity toward aquatic organisms

3.1. Generation of ROS

Despite the large number of studies on NPs toxicity both in cellline systems and organisms, a complete understanding about themechanisms behind is still lacking (see reviews (Manke et al., 2013;Fu et al., 2014; Bour et al., 2015; Schultz et al., 2014)). ROS genera-tion, whose overproduction can lead to oxidative stress (OS) in theorganism tissues, is unquestionably the most studied nanotoxicitymechanism.

Molecular oxygen is used as an oxidizing agent for the pro-duction of adenosine triphosphate (ATP) in the organism cells,being afterwards reduced to water. The non reduced oxygen resultsin the formation of superoxides

(

O−

2•)

that can be further con-verted to hydroxyl radicals (HO•), which has the highest reductionpotential of all the physiological relevant ROS. When under con-trol, these species are easily scavenged by (i) antioxidant agentssuch as polyphenols (Fu et al., 2014; Lipinski, 2011) (e.g., elim-ination of HO•), and (ii) enzymes such as superoxide dismutase(SOD), catalase (CAT) and glutathione peroxidase (GPx). The SODenzymes catalyze the dismutation of O−

2• into oxygen or hydro-

gen peroxide (H2O2), which is decomposed by CAT into water andoxygen. Even though H2O2 is less reactive than the radical species,is still a strong oxidant that needs further elimination. The GPx,also plays a role in the detoxification of H2O2 by using glutathione


(GSH) as a reductant. During the process, GSH is oxidized and con-verted to glutathione disulfide (GSSG) being latter reduced back toGSH by glutathione s-transferase enzymes (GST), thus completingthe cycle (Brigelius-Flohe and Maiorino, 2013; Deponte, 2013). Theantioxidant-enzymes activity is considered a reflection of the redoxstate of the cells and is frequently studied as a biomarker of OS.

When ROS is overproduced beyond the organism antioxidantresponse capacity, it leads to several deleterious effects on the cellscomponents such as lipids, proteins and DNA, possibly resultingin lipid peroxidation, apoptosis and/or cancer initiation processes,respectively. The production of ROS can be enhanced by the pres-ence of NPs, depending mainly on their size, aggregation, solubilityand coating. It is commonly accepted that smaller particles can eas-ily penetrate cell membranes, and thus induce cytotoxicity (Sakaiet al., 2011; Buzea et al., 2007). However, this correlation betweensize and toxicity is still controversial. Shi et al. (2013) reportedthat 5–10 nm nAg had higher toxicity to Tetrahymena pyriformis

when compared to slightly larger nAg (15–25 nm), while no size-dependent response on Danio rerio was obtained when exposedto 20, 50 and 110 nm (Bowman et al., 2012). Most studies showthat toxicity increases with decreasing particle size. However, oth-ers reported that either the size has no role on toxicity or thatsmaller NPs are less toxic (Ivask et al., 2014). A consensus aboutthe size effect is still lacking, and, most probably, will be unlikelyto be attained since the effects seem to be NP- and even specie-dependent (Buric et al., 2015).

The dissolution of metallic NPs such as nCuO and nAg resultsin the release of Cu and Ag ions, which are known to catalyzeFenton, Fenton-like and Haber–Weiss reactions, leading to the for-mation of ROS (Fu et al., 2014; Lipinski, 2011; He et al., 2012; Wanget al., 2013b,c). Moreover, the highly reactive surface of NPs andthe presence of manufactured and/or natural coatings can lead tothe adsorption/complexation of trace metals present in the envi-ronmental compartment also acting as a catalyzer platform to theabove-mentioned reactions, and thus increasing the concentrationof ROS in the system. Photoactive NPs such as nTiO2 and nZnO,can also induce the formation of ROS. When exposed to visible orUV light, these NPs can be photo excited resulting in the forma-tion of electron–holes, which are powerful oxidants that can reactwith surface bounded molecules forming radicals (Clemente et al.,2014). All these processes are schematized in Fig. 2.

3.2. Omics endpoints

The omics tools, such as toxicogenomic, metallomic and pro-teomic, are very useful on the establishment of toxic endpoints.A toxicogenomics approach allows the identification of gene andprotein activities in the organisms cells induced when in the pres-ence of a certain xenobiotic. A central assumption is that chemicalsgenerating toxicity by the same mechanism will produce similargene expression responses under a given set of conditions, bring-ing new insights about their mode of action that can be linked totheir specific physicochemical properties. A metallomic and pro-teomic approach will allow a complete analysis on the metal andmetalloid species composition within a cell or tissue and the estab-lishment of metalloproteins profiles leading to the identification ofnew biomarkers (e.g., proteins expressed by the NP itself (Shepardet al., 2000)). This will allow a better understanding and profil-ing of NPs toxic mechanisms discriminating them from their bulkcomponents.

Despite the large number of studies about nanotoxicology, onlya few number have reported the use of omics tools to evaluatethe NPs toxic effects at the molecular level on freshwater organ-isms (Rainville et al., 2014; Kuznetsova et al., 2014). Indeed, the“omics” approach generates a huge amount of data whose inter-pretation is not always straightforward, being, most probably, the

main reason moving nanotoxicologists away from using these tools.This large amount of data together with the fact that usually anappropriate physicochemical characterization prior to the biolog-ical assays is not performed, results in an escalating number ofunknown variables impeding a comprehensible understanding ofthe biochemical responses.

4. NPs toxicity on freshwater organisms

4.1. Silver NPs (nAg)

nAg are known for their antifungal and antimicrobial proper-ties, being extensively used in several products such as clothing,cosmetics, medical devices, paints, humidifiers and refrigerators(Reidy et al., 2013; Fabrega et al., 2011), leading to predicted con-centrations of 0.088–2.16 ng L−1 in European and North Americansurface waters (Gottschalk et al., 2009).

Despite nAg is one of the most studied NPs, the toxicity mech-anisms are still not fully clear; some assign the nAg toxicity to therelease of Ag+ ions to the media, while others assume that intactnAg particles are responsible for the induction of toxic responsesin the organisms (Schultz et al., 2014). It is well known that Ag+

ions has a great propensity to bioaccumulate in the tissues leadingto ROS generation, genotoxicity and inhibition of Na+/K+-ATPaseactivity by blocking the Na+ uptake by the cells (Luoma, 2008;Morgan et al., 1997). Similar toxic effects were also observed injuvenile Oncorhynchus mykiss fishes exposed to citrate coated nAg,with claimed low propensity for dissolution (Schultz et al., 2012).However, it was not possible to confidently establish if these inhi-bitions were caused by the nAg per se or by dissolved Ag+. In fact,it is very difficult to distinguish between the toxic effects inducedby particulate or ionic Ag, and, thus, is crucial not only to evaluatethe size distribution but also to quantify the nAg dissolution in thebioassay (Table 1).

It was possible to identify two distinct routes that could inducebiochemical responses in freshwater organisms (Table 1) (1) pres-ence of the NPs per se, and (2) presence of both nAg and dissolvedAg+. For the first route the identification of responses that areuniquely due to the presence of nAg and do not occur in matchedAg+ exposures is crucial. This is possible by (i) using nAg with lowdissolution rates, so, that the leached Ag+ in the media is insufficientto induce toxicity to the organisms, or (ii) identifying endpointsspecific to nAg, such as internalization of NPs, cytotoxicity andgenotoxicity. The work performed by Kumar et al. (2014) is anexample on the application of these strategies. They reported a sig-nificant increase of ROS and SOD and morphological alterations onfreshwater bacteria exposed to nAg coated with PVP. Since the NPsdissolution was very low (leached Ag+ < 1 �g L−1), it was concludedthat the results were related to the silver particulate form. Mor-phological changes and alteration in genes profiles, related to thepresence of nAg itself, were also observed in Cyprinus carpio (Leeet al., 2012), zebrafish (Griffitt et al., 2009; Choi et al., 2010), Daph-

nia magna (Poynton et al., 2012), medaka fish (Pham et al., 2012)and rainbow trout (Gagne et al., 2012). The exposure of 7 day oldD. magna to citrate-coated nAg (size 10 nm, 30 �g L−1) and AgNO3

(2.5 �g L−1) during 24 h showed that both Ag forms act throughdifferent pathways; both forms increased proteins thiol content,but only particles increased proteins carbonyl levels (Gundel et al.,2007).

For the second route, both forms, particulate and ionic Ag, caninduce OS and genotoxicity being the distinction between theseeffects a truly challenge. For example, in Chlorella vulgaris a positivecorrelation between ROS production and LPO on the tissues after24 h of exposure to uncoated nAg (1 and 10 mg L−1) (Oukarroumet al., 2012) was found, but with no possibility to establish a positive


Fig. 2. Potential routes for the generation of ROS due to the presence of NPs. (1) Internalization of NPs–ROS generation could occur due to the NPs dissolution inside the

cells and/or due to the NPs photocatalytic activity. (2) Dissolution of the NPs leads to an increase concentration of metal ions in the media; some of these metals can also be

uptake by the organisms. (3) NPs and/or their surrounding coatings can adsorb/complex other metals present in the media, being taken up by the cells. (4) Photocatalytic

activity of the NPs in the presence of UV and/or natural light.

correlation with solely the particulate or ionic Ag. Similar difficul-ties were also observed in the snail Lymnaea luteola (Ali et al., 2014),where the exposure to nAg resulted in DNA damage and inductionof OS but without the possibility to identify which Ag forms has thedominant role on the observed effects. In these situations, a com-plete physicochemical characterization of the NP in the exposuremedia along with the use of controls containing ionic Ag are cru-cial to understand which Ag form (or both) is responsible for theobserved biochemical responses.

Although in the majority of the studies is not possible to estab-lish if the nAg potential toxicity is due to its particulate or ionicform, it is clear that the presence of nAg in freshwater systemspresents a high risk to aquatic life.

4.2. Titanium dioxide NPs (nTiO2)

nTiO2 is one of the most produced NPs in the world, withan expected production of 201,000 tons during 2015 (Epa, 2011;Markets, 2015). Some studies have reported evidences that thisNP have a low toxicity toward aquatic organisms even at concen-trations higher than the ones expected to occur in the freshwatersystems (3 ng L−1 to only 1.6 �g L−1) (Gottschalk et al., 2013).Federici et al. (2007) have showed that even at 0.1–1 mg L−1 ofnTiO2 the rainbow trout, defense system can naturally scavengeROS species avoiding OS. For this reason, most of the nanotoxicol-ogy studies (if not all) uses much higher concentrations with theobjective of obtain more straightforward acute toxicity responses.Evidently, as used NPs concentrations are largely higher than theabove predicted values, the environmental impact of these stud-ies decreases significantly. Moreover, higher nTiO2 concentrationsimpacts on their own undergone transformations; the presence ofa higher number of particles can lead to homoaggregation (seeSection 2), most probably, resulting in a misinterpretation of theobtained results. Dalai et al. (2013) showed that for nTiO2 concen-trations larger than 16 mg L−1, ROS levels in the Ceriodaphnia dubia

decreased significantly due to the agglomeration of the NPs becom-ing less bioavailable to the organisms. The exposure of O. mykiss

(rainbow trout) to fairly low concentrations of nTiO2 (0.1–1 mg L−1)resulted in biochemical disturbances, respiratory distress and sev-eral organ pathologies (Federici et al., 2007; Boyle et al., 2013).However, the same did not occur, when other organisms wereexposed to higher concentrations (Dalai et al., 2013; Hao et al.,2009). Table 2 resumes the biochemical responses of freshwaterorganisms, mostly daphnids and fishes exposed to nTiO2. In mostof these studies, the nTiO2 characterization was solely based on theevaluation of the NPs size distribution since aggregation is the mostimportant transformation for this particle.

Three main routes leading to nTiO2 toxicity were identified: (i)physical stress, associated with the NPs size and surface properties(cytotoxicity), (ii) photocatalytic activity (phototoxicity), and (iii)NPs capacity do adsorb xenobiotics in the media.

The internalization of nTiO2 by an organism may result in itsaccumulation in different organs leading to physical stress and tis-sues damage. For instance, accumulation of nTiO2 in daphnid’s guts(Dalai et al., 2013; Tan and Wang, 2014; Zhu et al., 2010), bivalve’sdigestive gland (Vale et al., 2014) and fish’s gills (Federici et al.,2007; Boyle et al., 2013; Hao et al., 2009) constricts their alimentarycanal affecting their breathing capacity. The possible mechanismsinvolved in the NPs internalization in the organisms were alreadydescribed (Reidy et al., 2013; Chen and Bothun, 2014; Shang et al.,2014), and will not be further discussed here.

Due to their photocatalytic activity, nTiO2 can generate radi-cal species when exposed to natural and/or UV light radiation. Alinear relationship between illumination and the hydroxyl radi-cal generation in the media was established in presence of nTiO2,with the extracellular radical species generated inducing oxida-tive damage on the gill tissue cells of D. rerio (Xiong et al., 2011).Similar effects were also observed in other freshwater organismssuch as zebrafish (Bar-Ilan et al., 2013), rainbow trout (Boyle et al.,2013) and daphnids (Dalai et al., 2013). Moreover, nTiO2 can leadto toxicity assumed to be promoted by OS under dark conditions inlaboratory and in microcosm study (Jomini et al., 2012).

The nTiO2 can adsorb or interact with different elements, (e.g.,Cd, Zn, Pb, Cu, Ni and As (Vale et al., 2014; Engates and Shipley,


Table 1

Impact of nAg in freshwater organisms, including physicochemical characterization of the NPs in the media and biochemical responses. The biomarkers of response expressed

as ↑ and ↓, indicates an increase or decrease on the biomarkers activity or concentration, respectively, when compared with the control groups (organism not exposed to

nAg). The biomarkers here listed are: superoxide dismutase (SOD), catalase (CAT), glutathione (GSH), glutathione-s-transferase (GST), total glutathione peroxidase (GPx),

metallothioneins (MTs), lipid peroxidation (LPO) and reactive oxygen species (ROS).

Supplier information

(supplier; coating)

nAg characterization

(Size (nm);

concentration

(mg L−1); medium)

Bioassay

Organism Medium Biomarkers of response

(nAg concentration

(mg L−1))

Ref.

Sigma–Aldrich

uncoated

260 ± 26 (DLS)a;

0.001–0.08; TWc

32 (TEM)b; 0.001–0.08;

TWc

Lymnaea luteola L. TWc After 24 h exposure:

↓GSH (0.012; 0.024;

0.036); ↑CAT (0.012;

0.024; 0.036); ↓GPx

(0.024; 0.036); ↓GST

(0.036); ↑LPO (0.012;

0.024; 0.036); DNA

damage

After 96 h exposure:

↓GSH (0.012; 0.024;

0.036); ↑CAT (0.012;

0.024; 0.036); ↓GPx

(0.024; 0.036); ↓GST

(0.024; 0.036); ↑LPO

(0.004;0.012; 0.024;

0.036); DNA damage

Ali et al. (2014)

MTI Corporation

uncoated

307 (DLS)a; 1; BG-11d

50 (TEM)b BG-11d

Chlorella vulgaris BG-11d ↑ROS (1 and 10); ↑LPO

(1 and 10)

Oukarroum et al.

(2012)

ABC Nanotech

citrate-capped

70 (DLS)a; 5; FW e

90 (DLS)a; 10; FW e

12 (TEM)b; 5 and 10;

Freshwater e

Cyprinus carpio FWe Brain: ↓GST (0.1 and

0.2)

Liver: ↑GST (0.05; 0.1

and 0.2); ↓CAT (0.025;

0.05 and 0.1)

Lee et al. (2012)

Sigma– Aldrich

PVP coated

443 ± 15 (DLS)a; 1;

FSLWf

20-100 (TEM)b: 1;

FSLWf

B. thuringiensis and

B. aquimaris

FSLWf ↑ROS (1)

↑SOD (1)

Kumar et al. (2014)

Nanopoly

uncoated

5–20 (TEM)b D. rerio BWg ↑LPO (60,120)

↑GSH (120)

↓CAT (60,120)

↓GPx (120)

Choi et al. (2010)

a Dynamic light scattering.b Transmission electron microscopy.c Tap water.d Culture medium for Cyanobacteria (Rippka et al., 1979).e Freshwater.f Filtered and sterile lake water.g Bottled water.

2011; Gao et al., 2004; Sun et al., 2006)), changing their specia-tion in the media and affecting their bioavailability and toxicityto organisms. For instances, it was shown a correlation betweenthe decrease on Cd toxicity to the green algae Pseudokirchneriella

subcapita and the presence of nTiO2 not internalized by the algae.But on the other hand, if nTiO2 is already present in the organ-ism tissues, an increased bioaccumulation of xenobiotics due toadsorption onto the internalized NPs can occur (Tan and Wang,2014). Dispersed nTiO2 can also act as a carrier of xenobiotics inthe media increasing the metal uptake rate (Zhang et al., 2007b).

Independently of the route taken, nTiO2 toxicity is mostly asso-ciated with ROS generation, and the most common biochemicalendpoints are related with the anti-oxidant enzymes activity (seeTable 2). In D. magna, an increase of CAT, GST and GPx activity wasobserved when animals were exposed to 5–10 mg L−1 of nTiO2 (Kimet al., 2010), whereas in C. dubia an increase in SOD activity was alsoobserved after exposure to 1–64 mg L−1 nTiO2 (Dalai et al., 2013).Similar responses were also found in fishes, D. rerio (Clemente et al.,2014; Xiong et al., 2011), C. carpio (Hao et al., 2009) and O. mykiss

(Federici et al., 2007).

4.3. Zinc oxide NPs (nZnO)

As for nTiO2 also, nZnO are among the most used NPs withan estimated production of 30,000 metric tons per year (2015),originating predicted environmental concentrations in surfacewaters ranging from 0.008 to 0.055 �g L−1 in Europe and 0.001 to0.003 �g L−1 in US (Gottschalk et al., 2013, 2009). However, and asfor the other NPs, the concentrations used in ecotoxicology studiesare in general far higher than the ones predicted (Table 3).

Besides being an essential microelement, when at higher con-centrations Zn2+ is known to be toxic to aquatic organisms (Brunet al., 2014; Mortimer et al., 2010a). Being dissolution one of themain transformations of this particle is necessary to consider thattoxicity effects may be provoked by the NP per se and/or by the ionicfraction (Bondarenko et al., 2013; Franklin et al., 2007). Whereasdissolution of nZnO is hindered by the increase of the particle sizeor agglomerates size (Brun et al., 2014), the presence of proteinscan enhance it due to the proteins binding ability toward Zn (Reedet al., 2012). The presence of manufactured stabilizers can alsogreatly affect the NPs dissolution, with largest dissolved Zn usuallyobserved for uncoated nZnO followed by polymer-stabilized par-


Table 2

Impact of nTiO2 in freshwater organisms, including physicochemical characterization of the NPs in the media and biochemical responses. The biomarkers of response expressed


nTiO2). The biomarkers here listed are: superoxide dismutase (SOD), catalase (CAT), glutathione (GSH), glutathione-s-transferase (GST), total glutathione peroxidase (GPx),

metallothioneins (MTs), lipid peroxidation (LPO) and reactive oxygen species (ROS).


(supplier; chrystal

phase)

nTiO2 characterization

(Size (nm);

concentration

(mg L−1); medium)

Bioassay


(nTiO2 concentration

(mg L−1))

Ref.

Sigma–Aldrich

30% rutile + 70%

anatase

400–800 (DLS)a; 5;

MHWb

D. magna MHWb ↑CAT (10); ↑GST (5 and

10); ↑GPx (5 and 10)

Kim et al. (2010)

Sigma–Aldrich

100% anatase

400–700 (DLS)a; 100;

EEMc

D. rerio EEMc Visible light: ↓CAT (1

and 10); ↑GST (10)

UV + visible light: ↓GST

(10)

Clemente et al. (2014)

Sigma–Aldrich

100% anatase

20.9 ± 2.86 (DLS)a; 0.1;

SM7d

218 ± 47.3 (DLS)a; 1;

SM7d

21 (TEM)d;; 0.1 and 1;

SM7d

D. magna SM7d No significant effects

on ROS and MTs

Tan and Wang (2014)

Sigma–Aldrich

100% anatase

248–293(1); FSLWe

517–925 (16); FSLWe

697–1090 (64); FSLWe

C. dubia FSLWe Photoperiod (16 h

light: 8 h dark): ↑SOD

(1–64); ↑ROS (1–32); ↓

ROS (>32)

Dark period:

↑SOD(1–64); ↑ROS

(1–64)

Dalai et al. (2013)

Degussa Evonik

80% rutile + 20%

anatase

>1000 (DLS)a; 100;

EEMc

D. rerio EEMc UV + visible light: ↓CAT

(1 and 10); ↓GST (1)

Clemente et al. (2014)

Degussa Evonik

75% rutile + 25%

anatase

24 (TEM)f; 0.1;0.5 TWg O. mykiss TWg Liver tissues: ↓GSH (0.5

and 1)

Gill tissues: ↑ GSH (1);

↑LPO (0.1 and 5)

Brain tissues: ↑LPO

(0.1; 0.5 and 1)

Intestine tissues: ↑LPO

(0.1; 0.5 and 1)

Federici et al. (2007)

Nanjing University

100% anatase

249–488 (DLS)a; 1;

DWh

270–535 (DLS)a; 10;

DWh

251–630 (DLS)a; 50;

DWh

402–633 (DLS)a;100;

DWh

245–617 (DLS)a; 300;

DWh

20–70 (TEM)f; DWh

D. rerio DWh Liver tissues: ↓CAT

(50), ↓SOD (50), and

↓GSH (50)

Gut tissues: ↑SOD (50);

↑GSH (50); ↑LPO (50)

Gill tissues: ↑LPO (50)

Xiong et al. (2011)

a Dynamic light scattering.b Moderately hard water.c Embryo exposure medium.d Elendt simplified M7 medium.e Filtered and sterile lake water.f Transmission electron microscopy.g Tap water.h Distillated water.

ticles (Merdzan et al., 2014). These NPs can also dissolve after itsinternalization in the extra- or intracellular compartments (Fig. 2),dramatically changing the organism’s metallome profile.

The exposure of the algae P. subcapitata to nZnO, ZnSO4, ZnCl2and bulk ZnO resulted in similar toxic effects indicating that ionic Znwas the responsible for the observed toxicity (Franklin et al., 2007;Aruoja et al., 2009a; Neale et al., 2015). Dose–dependent adverseeffects of nZnO on D. rerio embryos and eleuthero-embryos wereobserved (Brun et al., 2014), resulting in hatching and inflammationreactions (Yu et al., 2011; Xia et al., 2011). Zn may act as a competi-tor with Ca influx in skin chloride cells by blocking its transportthrough the pore canals and affecting a large surface of the skin. The

observed toxic effects were related to the ionic Zn, since the NPssize distribution was too large to pass through the chorion pores.In contrast, zebrafish embryos showed a greater acute toxicity tonZnO than when in presence of ionic Zn (Zhu et al., 2008).

nZnO can establish hydrogen bonds and ligand exchanges withthe bacterial cell wall, modifying the protein structure and conse-quently changing its function (Jiang et al., 2010). These particlescan also bind to the cytoplasmic bacterial membrane possibly dis-rupting its integrity, and interrupt the fundamental role of electrontransport phosphorylation and energy transduction process (Lyonet al., 2007). nZnO can generate ROS due to its photocatalyticalproperties (Suresh et al., 2015), but can also suffer photocorrosion


Table 3

Impact of nZnO in freshwater organisms including physicochemical characterization of the NPs in the media and biochemical responses. The biomarkers of response expressed


nZnO). The biomarkers here listed are: superoxide dismutase (SOD), catalase (CAT), glutathione (GSH), glutathione-s-transferase (GST), total glutathione peroxidase (GPx),

lipid peroxidation (LPO), nitric oxide (NO), reactive oxygen species (ROS), tumor necrosis factor � (TNF�), myxovirus-resistance protein A (MxA), interleukin-1 beta (IL-1�),

mitochondrial uncoupling protein 2 (Ucp-2), heat shock proteins (HSP) and macrophage scavenger receptor (MSR).


(supplier; chrystal

phase)

nZnO characterization

(Size (nm);

concentration

(mg L−1); medium)

Bioassay



(mg L−1))

Ref.

Genes’Ink 218–316 (NTA)a; 5;

HM b

196–211 (NTA)a; 0.2;

HMAc

214–236 (NTA)a; 1;

HMA c

223–242 (NTA)a; 5;

HMAc

D. rerio

(embryos)

HMAc ↑ CAT (0.2, 1 and 5)

48 hpf; ↓ CAT (5) 96 hpf

↑ Cu/Zn-SOD (5)

48 hpf; ↓ Cu/Zn-SOD

(5) at 96 hpf

↑ mt2 gene (0.2, 1 and

5) 48 and 96 hpf

↓ TNF� (5) 96 hpf

↑ c-jun protein (0.2, 1

and 5) 96 hpf

↓ MxA (1 and 5) at

48 hpf; ↓ MxA (5) 96

hpf

↓ IL-1� (5) 96 hpf

Brun et al. (2014)

Genes’Ink 218–316 (NTA)a; 5;

HMWAb

196–211 (NTA)a; 0.2;

HMAc

214–236 (NTA)a; 1;

HMAc

223–242 (NTA)a; 5;

HMAc

D. rerio

(eleuthero-embryos)

HMA c ↑ CAT (0.2 and 1)

120 hpf

↑ mt2 gene (5) 120 and

168 hpf

↑ TNF� (1 and 5)

120 hpf

↓ Cu/Zn-SOD (5)

120 hpf

↓ c-jun protein (1 and

5) 120 and 168 hpf,

↓ MxA (5) 120 and

168 hpf,

↓ IL-1� (1 and 5) 120

and 168 hpf

Brun et al. (2014)

Nanjing High

Technology

2196–3144 (DLS)d; 10;

ZCMe

D. rerio (embryos) ZCMe ↑ ROS (1 and 10) at

96 hpf

↓ GSTp2 (10) at 96 hpf

↓ Nqo1 gene (10) at 24

and 96 hpf

↑ GSTp2 and Nqo1

gene (sediment + nZnO

10) at 96 hpf

Zhu et al. (2009)

Sigma–Aldrich 50–100 (DLS)d; 50;

ZCMe

D. rerio (embryos)

at 144 hpf

ZCMe ↑ SOD (1–100,

dose–dependent from

1 to 50)

↑ MDA (20–100); ↑ GPx

(20–100); ↑ DNA

damages (100)

↑ ROS (1 and 10; ↑

Ucp-2 (1–100);

↓ CAT (100); ↓ GSTp2

(50–100); ↓ Nqo1 gene

(100);

↓ Bcl-2 genes (50–100)

Zhao et al. (2013)

BASF UK

Z-COTE©

150 ± 60 (TEM)f; 100;

NHg

E. coli NHg ↑ SoxS (100) McQuillan and Shaw

(2014)

Nanostructured

& Amorphous

Material, Inc.

24–72 (FCS)h; 1; EPi

10–13 (AUC)j; 1; EPi

C. reinhardtii EPi Moderate Visible light:

↑ GSTs1 (1); ↑ HSP22C

(1);

↑ HSP70A (1); ↑

MSR1(1); ↑ MSR2(1);

and ↑ HSP90(1)

Simon et al. (2013)

nCuO; Nanostructured

& Amorphous

Materials, Inc.

∼130 (DLS)a; 10; MESb

∼140 (DLS)a:10;

MOPSc

∼140 (DLS)a;10; TAPd

∼140 (DLS)a;10; OECDe

∼140 (DLS)a; 10; LWf

Chlamydomonas

reinhardtii

MESb; MOPSc; TAPd;

OECDe; LWf

↑ROS showing

OECDe > MOPSc > MESb > TAPd > LWf

after 24 h

von Moos et al. (2015)

MTI Corporation 148(DLS)a; 40; HSMg C. reinhardtii HSMg ↑ROS (0.004 g L−1) Perreault et al. (2012)

Nanostructured

& Amorphous

Material, Inc.

27.2 ± 6.7 (TEM)f; 22;

MHRWk

D. magna MHRWk ↓ Ferritin 3 (2.2 and

9.0); ↓ C1q proteins

(2.2 and 9.0)

Poynton et al. (2011)


Table 3 (Continued)


(supplier; chrystal

phase)

nZnO characterization

(Size (nm);

concentration

(mg L−1); medium)

Bioassay



(mg L−1))

Ref.

Sigma–Aldrich 264.8 (DLS)d; 1000;

DWl

L. luteola L. DWl Digestive gland cells

↑ DNA Damage (10–32)

24 h and 96 h

↑ LPO (21–32) 24 h; ↑

LPO (10–32) 96 h

↑ CAT (21–32) 24 h and

96 h

↓ GPx (21–32) 24 h; ↓

GPx (10–32) 96 h

↓ GST (21–32) 24 h; ↓

GST (10-32) 96 h

↓ GSH (21–32) 24 h and

96 h

Ali et al. (2012)

Sigma–Aldrich 71.11 (DLS)d; 1000;

DWl

B. alexandrina DWl Hemolymph: ↓ CAT

(7–35); ↓ SOD (7) and ↑

SOD (35);

↓ GSH (7–35); ↓ GST

(7–35); ↑ NO (7–35)

↑ LPO (7–35)

Tissues: ↓ CAT (7) and ↑

CAT (35); ↓ SOD (7) and

↑ SOD (35); ↓ GSH

(7–35); ↓ GST (7–35); ↑

NO (7–35)

↑ LPO (7–35)

Fahmy et al. (2014)

a Nanoparticle tracking analysis.b Holtfreter’s medium.c Holtfreter’s medium with alginate.d Dynamic light scattering.e Zebrafish culture medium.f Transmission electron microscopy.g Neidhardt’s medium.h Fluorescence correlation spectroscopy.i Experimental media.j Analytical ultra centrifugation (Saison et al., 2010).k Moderately hard reconstituted water.l Destilated water.

when exposed to UV light, decreasing their photocatalytic activ-ity in aqueous suspensions (Hariharan, 2006). OS induced by bothnZnO and/or released Zn2+, has already been observed in severalfreshwater organisms, such as bacteria (Lyon et al., 2007; Cabiscolet al., 2000; Adams et al., 2006; Zhang et al., 2014; Gunawanet al., 2013), algae (Simon et al., 2013), crustacean (Mwaanga et al.,2014), mussel (Gagné et al., 2013), snail (Fahmy et al., 2014), frog(Bacchetta et al., 2012, 2013; Nations et al., 2011) and fish (Brunet al., 2014; Xia et al., 2011; Hao and Chen, 2012; Zhu et al., 2009;Bai et al., 2010; Zhao et al., 2013; Nel et al., 2006). In bacteria,the OS effect of nZnO may occur under dark conditions as wellas under natural or artificial light and affect both gram-positiveand negative bacteria (Barnes et al., 2013). After exposure to UVlight radiation, a loss of viability in Escherichia coli and ROS gener-ation results in the expression of genes encoding SoxS and CAT,which are part of the OS response mechanism (Gunawan et al.,2013; Kumar et al., 2011). In the algae Chlamydomonas reinhardtii,the presence of nZnO led to elevated levels of transcripts genes,GSTS1, HSP22C, and HSP70A, and the transcripts encoding sub unitsMSR1, MSR2, and HSP90 which are involved in the organism defenseresponse against ROS (Simon et al., 2013). In bivalves, the presenceof nZnO led to an increase of LPO in the digestive gland (Gagnéet al., 2013). Similar findings were also observed in snails (Fahmyet al., 2014), with a decrease in GSH content, a significant inhi-bition of SOD and CAT activity and a decrease in total protein

and albumin contents. A significant increase of LPO levels and adecrease of GSH activity in the gills, liver and brain of carps werealso reported after a 14 days exposure to 50 mg nZnO L−1 (Haoand Chen, 2012). A concentration-time exposure effect on SOD,CAT, GPx, and Ucp-2 expression with an increase in MDA contentwas shown for zebrafish embryos (Brun et al., 2014; Zhao et al.,2013).

As found for nTiO2, nZnO can also adsorb several elementssuch as As, Al, Mo, Hg, Pb, Cu, Ni and Cd (Gagné et al., 2013;Ghiloufi, 2013; Hua et al., 2012; Mahdavi et al., 2012; Sheelaet al., 2012), changing their speciation in the media and therebytheir bioavailability. When internalized, these NP-metal complexesmay undergo dissociation followed by NPs dissolution due to theacidic conditions found internally resulting in the release of theadsorbed xenobiotics, and drastically increasing their concentra-tion in organism tissues.

Overall, these studies highlight the fact that, despite the vastliterature, more investigation is needed to fully elucidates themechanisms of nZnO toxicity from the cellular level to the organ-isms or population level.

4.4. Copper oxide NPs (nCuO)

nCuO is a widely used metal oxide presenting biological activ-ity (Moschini et al., 2013; Blinova et al., 2010; Chen et al., 2008;


Table 4

Impact of nCuO in freshwater organisms including physicochemical characterization of the NPs in the media and biochemical responses. The biomarkers of response expressed


nCuO). Lipoprotein (LP) and reactive oxygen species (ROS).


(supplier)

nCuO characterization

(Size (nm);

concentration

(mg L−1); medium)

Bioassay


(nCuO concentration

(mg L−1))

Ref.

Nanostructured &

Amorphous

Materials, Inc.

∼130 (DLS)a; 10; MESb

∼140 (DLS)a:10;

MOPSc

∼140 (DLS)a;10; TAPd

∼140 (DLS)a;10; OECDe

∼140 (DLS)a; 10; LWf

Chlamydomonas

reinhardtii

MESb; MOPSc; TAPd;

OECDe; LWf

↑ROS showing

OECDe > MOPSc

>MESb > TAPd > LWf

after 24 h

von Moos et al. (2015)

MTI Corporation 148(DLS)a; 40; HSMg C. reinhardtii HSMg ↑ROS (0.004 g L−1) Perreault et al. (2012)

Sigma–Aldrich 209 ± 10 (DLS)a; 100;

MQWh

1230 ± 200 (DLS)a;

100; OMMi

25.5 ± 0.8 m2 g−1 (BET)j

T. thermophila OMMi ↑ROS (80); ↑LP (60) Mortimer et al. (2011)

Sigma–Aldrich 30 (SEM)k T. thermophila

(strain BIII)

OMMi Cytotoxicity:

fluorescence (4 h: 127

(124–144); 24 h: 97.9

(80.4–138) mg Cu L−1);

ATP (4 h: 129

(111–149); 24 h: 101

(91.1–190a mg Cu L−1)

Mortimer et al. (2010b)

MTI Corporation 523-800 (DLS)a; NRm ,

SFWn

−39.7 ± 3.8 mV (EPM)o;

NRm; SFWn

Lemna gibba SFW 150 ↑ROS (0.7 g L−1 of total

Cu) > esterase activity

Perreault et al. (2014)

a Dynamic light scattering.b 2-(N-morpholino) ethanesulfonic acid.c 3-(N-morpholino) propanesulfonic acid.d TAP x4 algae growth media.e OECD algae growth media.f Lake water (filtered and sterile).g High salt medium (Saison et al., 2010).h MilliQ water.i Osterhout’s mineral medium.j Brunauer–Emmet–Teller analysis.k Scanning electron microscopy.

m Not reported.n Synthetic freshwater (Frankart et al., 2002).o Electrophoretic mobility.

Midander et al., 2009), and producing DNA damages and cell deathcompared to micro-sized particles, and nTiO2 and nZnO (Midanderet al., 2009; Ahamed et al., 2010; Nel et al., 2009). However, itsenvironmental hazard has been poorly investigated (Kahru andDubourguier, 2010) as can be observed in Table 4.

The presence of nCuO (0.1 mg Cu L−1) induced formation ofsuperoxide anions, hydrogen peroxide and single-stranded DNAin different recombinant luminescent E. coli (Bondarenko et al.,2012). However, the dissolution of these particles was the key fac-tor triggering ROS and DNA damage. In other study is was alsoobserved that the presence of nCuO and released Cu2+ induced anapproximately 5-fold increase in ROS in E. coli compared to thebacteria-only control (Gunawan et al., 2011). The levels of non-viable cells exposed to nCuO and released Cu2+ were very similarsuggesting that adverse effects were originated by the ionic form asfound by (Ivask et al., 2010). Also Hu et al. (2009) showed that par-ticles significantly inhibited viable count of bacteria (36.8 − 81.9%)when using 25–200 mg nCuO L−1.

A low mutagenic potential to Salmonella typhimurium TA 97aand TA100 (marginal effects between 100 and 1600 �g/plate) wasdisplayed by nCuO (Pan et al., 2010). The OS provoked by nCuOon diverse Saccharomyces cerevisiae strains was investigated andcom pared with CuSO4 and bulk CuO (Kasemets et al., 2013). The

cup2� (Cu stress response-deficient strain) was the most sensi-tive strain (approximately 16-fold than the wild type), suggestingthat nCuO effect proceeds also via dissolved Cu-ions. EC50s of nCuO(16–19 mg Cu L−1) and CuSO4 (10–12 mg Cu L−1) differed from thebulk CuO (918–1082 mg Cu L−1). Effects on yeast growth were alsoreported (Kasemets et al., 2009a).

The paradigm of nCuO OS (10 mg L−1) was also investigatedon C. reinhardtii (von Moos et al., 2015), with results (24 h expo-sure) showing an immediate cell size increase, OS, and chlorophyllbleaching, while membrane permeabilization was observed after5 h. Agglomerated nCuO was toxic and the exposure media wasdecisive in whether or not particles or ionic Cu act as the main tox-icity mediators. C. reinhardtii was also used to show the effect ofthe presence of a manufactured coating on nCuO (polystyrene-co-butyl acrylate) comparing the outcomes with bare nCuO (Perreaultet al., 2012). Higher toxicity was obtained in presence of coatednCuO mainly due to its internalization in the cytosolic membranestructures. ROS formation was observed at 4 mg L−1 of coated nCuOreaching (392 ± 12)% at 40 mg L−1, whereas 4 mg L−1 of bare nCuOformed ROS that increased only up to (160 ± 15)% compared to thecontrol (Perreault et al., 2012). Coated nCuO was also 10-fold moretoxic for Lemna gibba than bare nCuO (Perreault et al., 2014); the48 h exposure of 0.4 g L−1 of coated-nCuO led to a 50% growth inhi-


bition, while 4.5 g L−1 of bare nCuO were required to reproduce thesame effect. But particles dissolution played the major role in thetoxicity.

Tetrahymena thermophila was used to assess nCuO, bulk CuOand Cu salt effects on fatty acid profiling, ROS and LPO (Mortimeret al., 2011). All Cu forms induced ROS, but a larger induction wasobtained in presence of nCuO, which could not be explained solelyby the released Cu. Protozoa exposure (24 h) to 80 mg L−1 of nCuOsignificantly decreased the proportion of the two major unsatu-rated fatty acids, increasing the relative amount of two saturatedfatty acids. Analogous effects were not observed with other Cuforms. Several studies confirmed that ionic Cu contributed onlypartially to nCuO toxicity (Shi et al., 2011; Kasemets et al., 2009b;Aruoja et al., 2009b; Heinlaan et al., 2011). However, Mortimeret al. (2010b) showed that nCuO is 10–20-fold more toxic for T.

thermophila than bulk CuO mainly due to the released Cu.The D. magna exposed (96 h) to nCuO (<50 nm; 1.97 mg L−1)

resulted in significant alterations in (i) encoding protein acting inmetal ion binding, (ii) OS, and (iii) exoskeleton (Adam et al., 2015).In all cases, the alteration up-regulated the transcription level ofthe transcripts genes. Glycogen, lipid and protein concentrationin exposed daphnids was lower than unexposed organisms, butwhen comparing nCuO and metal salt exposures, the microarrayevidenced no significant differences in transcribed gene fragments.Thus, the toxicity of nCuO to D. magna was caused by Cu ions(Adam et al., 2015). nCuO (26.7 nm; 100 �g L−1) exposure (48 h)led to significant changes in gills transcriptome of adults D. rerio,with induction of genes involved in apoptosis, mitogenesis and cellproliferation (Griffitt et al., 2009).

Despite the very limited available data about nCuO ROS gen-eration, cytotoxicity and genotoxicity in for freshwater organisms,it can be concluded that nCuO provoke ROS toxicity mediated byionic Cu.

5. General remarks and conclusions

Most of the biochemical responses reported are related to theorganism’s ROS defense mechanisms, mainly through gene expres-sion or changes on anti-oxidant enzymes activities. As mentionedbefore (Section 3.2), data related with changes in freshwater organ-ism genome and proteome due to the presence of NPs is very scarce.This is indeed surprisingly since omics techniques have alreadyproved a great potential on the recognition of signatures related tospecific stress, eventually leading to the discover of new biomark-ers. The few data available suggest that the interaction with theorganisms is NP-specific. For example, it seems that both nTiO2

and nAg have similar targets in the organism physiology affectingboth protein synthesis and circadian regulation, while nZnO andnCuO effects are generally related with OS.

Photoactive NPs such as nTiO2 and nZnO, when under naturalor UV light, can generate ROS inducing OS. However, NPs composi-tion can also play an important role on ROS generation, since somemetals constituting the NP can instigate Fenton and Weiss-typereactions releasing ROS in the intra or extracellular media. Evi-dently, and as usually performed, the bioassays should contain acontrol group exposed to the salt form of the metal constitutingthe NP, allowing the distinction between the effects provoked bythe NP per se and/or by the NP dissolution products.

Despite the large number of studies dealing with ecotoxicologyof NPs, it is evident that is still not possible to establish cru-cial predictive structure-activity relationships. This is mainly dueto the fact that the majority of the available studies have criti-cal deficiencies on their experimental designs; a comprehensivephysicochemical characterization of the particles under the expo-sure conditions is mostly miscarried or restricted to a secondary

task. Despite the scientific community is already aware about theimportance of the NPs physicochemical characterization prior andduring the bioassay, this is still frequently neglected in the mostrecent studies giving rise to more confusing and contradictory data.Clearly, this pushes back the possibility to establish a proper envi-ronmental risk assessment plan for these current early generationson ENPs (1st and 2nd generation passive and active nanostructures,respectively), while advanced generations of ENPs (3rd and 4thgeneration nanosystems and molecular nanosystems) may not befar away, bringing additional challenges that require further novelapproaches.

The dynamic speciation of the NPs should be assessed in thesame exposure media of the bioassay by following key NPs transfor-mations: (i) dissolution, (ii) homo- and heteroaggregation, and (iii)sedimentation. Several analytical tools are nowadays available forthe quantification of these physicochemical transformations, eachof them having their specific advantages and limitations being ableto provide different information on ENPs properties (see reviewsPinheiro and Domingos, 2015; Tiede et al., 2008; Domingos et al.,2009b). This physicochemical characterization approach allows(i) to assess the bioavailable NP-containing species to which theorganisms will be exposed, and (ii) to relate the biocompatibleor bioadverse effects with the NP-containing species permitting aNP categorization and function. Nanotoxicology is indeed a mul-tidisciplinary field where the study of the NPs physic, chemistryand biological impact is crucial for a complete toxicological assess-ment. Unfortunately, there is a lack of legislation controlling theproduction, use and release of these materials to the environ-ment, and new NPs are commercialized every day without anappropriate assessment about their impact in environment andhuman health. The establishment of national and international lawsregulating the production of these materials is mandatory. Further-more, it is also urgent to increase the number of comprehensivenano(eco) toxicology studies under natural more environmentally-realistic conditions implying the co-presence of ENPs (at low andenvironmentally-realistic doses) and environmental constituentssuch as natural organic and inorganic dissolved and colloidal mat-ter. Only with these approaches a comprehensive risk assessmentwill be possible with production of environmentally safe-by-designENPs.

Acknowledgments

All the authors acknowledge to the European COST actionES1205, Engineered Nanomaterials—From Wastewater Treatment& Stormwater to Rivers (ENTER). G. Vale and R.F. Domingosacknowledges to Fundac ão para a Ciência e Tecnologia (FCT,Portugal) the post—doctoral grant SFRH/BPD/73117/2010 and theproject PTDC/AAC-AMB/111998/2009, respectively. The contribu-tion of K. Mehennaoui and S. Cambier were in part possible withinNanoGAM (AFP-PhD-9229040) and NANION (FNR/12/SR/4009651)respectively, both founded by Fonds National de la Recherche Lux-embourg.

References

Adam, N., Vergauwen, L., Blust, R., Knapen, D., 2015. Environ. Res. 138, 82–92.Adams, L.K., Lyon, D.Y., Alvarez, P.J.J., 2006. Water Res. 40, 3527–3532.Ahamed, M., Siddiqui, M.A., Akhtar, M.J., Ahmad, I., Pant, A.B., Alhadlaq, H.A., 2010.

Biochem. Biophys. Res. Commun. 396, 578–583.Ali, D., Alarifi, S., Kumar, S., Ahamed, M., Siddiqui, M.A., 2012. Aquat. Toxicol.

124–125, 83–90.Ali, D., Yadav, P.G., Kumar, S., Ali, H., Alarifi, S., Harrath, A.H., 2014. Chemosphere

104, 134–140.Aruoja, V., Dubourguier, H.-C., Kasemets, K., Kahru, A., 2009a. Sci. Total Environ.

407, 1461–1468.Aruoja, V., Dubourguier, H.C., Kasemets, K., Kahru, A., 2009b. Sci. Total Environ.

407, 1461–1468.

http://refhub.elsevier.com/S0166-445X(15)30102-8/sbref0005
















































Bacchetta, R., Santo, N., Fascio, U., Moschini, E., Freddi, S., Chirico, G., Camatini, M.,Mantecca, P., 2012. Nanotoxicology 6, 381–398.

Bacchetta, R., Moschini, E., Santo, N., Fascio, U., Del Giacco, L., Freddi, S., Camatini,M., Mantecca, P., 2013. Nanotoxicology 8, 728–744.

Bai, W., Zhang, Z., Tian, W., He, X., Ma, Y., Zhao, Y., Chai, Z., 2010. J. Nanopart. Res.12, 1645–1654.

Bar-Ilan, O., Chuang, C.C., Schwahn, D.J., Yang, S., Joshi, S., Pedersen, J.A., Hamers,R.J., Peterson, R.E., Heideman, W., 2013. Environ. Sci. Technol. 47, 4726–4733.

Barnes, R., Molina, R., Xu, J., Dobson, P., Thompson, I., 2013. J. Nanopart. Res. 15,1–11.

Blinova, I., Ivask, A., Heinlaan, M., Mortimer, M., Kahru, A., 2010. Environ. Pollut.158, 41–47.

Bondarenko, O., Ivask, A., Käkinen, A., Kahru, A., 2012. Environ. Pollut. 169, 81–89.Bondarenko, O., Juganson, K., Ivask, A., Kasemets, K., Mortimer, M., Kahru, A., 2013.

Arch. Toxicol. 87, 1181–1200.Bour, A., Mouchet, F., Silvestre, J., Gauthier, L., Pinelli, E., 2015. J. Hazard. Mater.

283C, 764–777.Bowman, C.R., Bailey, F.C., Elrod-Erickson, M., Neigh, A.M., Otter, R.R., 2012.

Environ. Toxicol. Chem. 31, 1793–1800.Boyle, D., Al-Bairuty, G.A., Ramsden, C.S., Sloman, K.A., Henry, T.B., Handy, R.D.,

2013. Aquat. Toxicol. 126, 116–127.Brigelius-Flohe, R., Maiorino, M., 2013. Biochim. Biophys. Acta 1830, 3289–3303.Brun, N.R., Lenz, M., Wehrli, B., Fent, K., 2014. Sci. Total Environ. 476–477, 657–666.Buric, P., Jaksic, Z., Stajner, L., Dutour Sikiric, M., Jurasin, D., Cascio, C., Calzolai, L.,

Lyons, D.M., 2015. Mar. Environ. Res. 111, 50–59.Buzea, C., Blandino, I.I.P., Robbie, K., 2007. Biointherphases 2, MR17–MR172.Cabiscol, E., Tamarit, J., Ros, J., 2000. Int. Microbiol. 3, 3–8.Chen, K.L., Bothun, G.D., 2014. Environ. Sci. Technol. 48, 873–880.Chen, X., Chen, J., Qiao, X., Wang, D., Cai, X., 2008. Appl. Catal. B: Environ. 80,

116–121.Choi, J.E., Kim, S., Ahn, J.H., Youn, P., Kang, J.S., Park, K., Yi, J., Ryu, D.Y., 2010. Aquat.

Toxicol. 100, 151–159.Clemente, Z., Castro, V.L., Moura, M.A., Jonsson, C.M., Fraceto, L.F., 2014. Aquat.

Toxicol. 147, 129–139.Dalai, S., Pakrashi, S., Chandrasekaran, N., Mukherjee, A., 2013. PLoS One 8, e62970.Dale, A.L., Lowry, G.V., Casman, E.A., 2013. Environ. Sci. Technol. 47, 12920–12928.Deponte, M., 2013. Biochim. Biophys. Acta 1830, 3217–3266.Derfus, A.M., Chan, W.C.W., Bhatia, S.N., 2004. Nano Lett. 4, 11–18.Domingos, R.F., Tufenkji, N., Wilkinson, K.J., 2009a. Environ. Sci. Technol. 43,

1282–1286.Domingos, R.F., Baalousha, M.A., Ju-Nam, Y., Reid, M.M., Tufenkji, N., Lead, J.R.,

Leppard, G.G., Wilkinson, K.J., 2009b. Environ. Sci. Technol. 43, 7277–7284.Domingos, R.F., Peyrot, C., Wilkinson, K.J., 2010. Environ. Chem. 7, 61.Domingos, R.F., Rafiei, Z., Monteiro, C.E., Khan, M.A.K., Wilkinson, K.J., 2013a.

Environ. Chem. 10, 306–312.Domingos, R.F., Franco, C., Pinheiro, J.P., 2013b. Environ. Sci. Pollut. Res. 20,

4872–4880.Domingos, R.F., Franco, C., Pinheiro, J.P., 2014. Environ. Sci. Pollut. Res. 22,

2900–2906.Domingos, R.F., Gélabert, A., Carreira, S., Cordeiro, A., Sivry, Y., Benedetti, M.F.,

2015. Aquat. Geochem., http://dx.doi.org/10.1007/s10498-014-9251-x.Engates, K.E., Shipley, H.J., 2011. Environ. Sci. Pollut. Res. Int. 18, 386–395.Epa U.S., Laser detection of nanoparticles in the environment, 2011.Fabrega, J., Luoma, S.N., Tyler, C.R., Galloway, T.S., Lead, J.R., 2011. Environ. Int. 37,

517–531.Fahmy, S., Abdel-Ghaffar, F., Bakry, F., Sayed, D., 2014. Arch. Environ. Contam.

Toxicol. 67, 192–202.Federici, G., Shaw, B.J., Handy, R.D., 2007. Aquat. Toxicol. 84, 415–430.Frankart, C., Eullaffroy, P., Vernet, G., 2002. Ecotoxicol. Environ. Saf. 53, 439–445.Franklin, N.M., Rogers, N.J., Apte, S.C., Batley, G.E., Gadd, G.E., Casey, P.S., 2007.

Environ. Sci. Technol. 41, 8484–8490.Fu, P.P., Xia, Q., Hwang, H.-M., Ray, P.C., Yu, H., 2014. J. Food Drug Anal. 22, 64–75.Gagné, F., Turcotte, P., Auclair, J., Gagnon, C., 2013. Comp. Biochem. Physiol. Part C:

Toxicol. Pharmacol. 158, 22–28.Gagne, F., Andre, C., Skirrow, R., Gelinas, M., Auclair, J., van Aggelen, G., Turcotte, P.,

Gagnon, C., 2012. Chemosphere 89, 615–622.Gao, Y., Wahi, R., Kan, A.T., Falkner, J.C., Colvin, V.L., Tomson, M.B., 2004. Langmuir

20, 9585–9593.Ghiloufi I., Recent Advances in Circuits Communications and Signal Processing,

2013, 321-326.Gottschalk, F., Sonderer, T., Scholz, R.W., Nowack, B., 2009. Environ. Sci. Technol.

43, 9216–9222.Gottschalk, F., Ort, C., Scholz, R.W., Nowack, B., 2011. Environ. Pollut. 159,

3439–3445.Gottschalk, F., Sun, T., Nowack, B., 2013. Environ. Pollut. 181, 287–300.Griffitt, R.J., Hyndman, K., Denslow, N.D., Barber, D.S., 2009. Toxicol. Sci. 107,

404–415.Gunawan, C., Teoh, W.Y., Marquis, C.P., Amal, R., 2011. ACS Nano 5, 7214–7225.Gunawan, C., Teoh, W.Y., Ricardo, Marquis, C.P., Amal, R., 2013. Part. Part. Syst.

Charact. 30, 375–380.Gundel, U., Benndorf, D., von Bergen, M., Altenburger, R., Kuster, E., 2007.

Proteomics 7, 4541–4554.Hao, L., Chen, L., 2012. Ecotoxicol. Environ. Saf. 80, 103–110.Hao, L., Wang, Z., Xing, B., 2009. J. Environ. Sci. 21, 1459–1466.Hariharan, C., 2006. Appl. Cata. A: Gen. 304, 55–61.

He, W., Zhou, Y.T., Wamer, W.G., Boudreau, M.D., Yin, J.J., 2012. Biomaterials 33,7547–7555.

Heinlaan, M., Ivask, A., Blinova, I., Dubourguier, H.-C., Kahru, A., 2008.Chemosphere 71, 1308–1316.

Heinlaan, M., Kahru, A., Kasemets, K., Arbeille, B., Prensier, G., Dubourguier, H.-C.,2011. Water Res. 45, 179–190.

Hotze, E.M., Bottero, J.Y., Wiesner, M.R., 2010. Langmuir 26, 11170–11175.Hu, X., Cook, S., Wang, P., Hwang, H.m., 2009. Sci. Total Environ. 407, 3070–3072.Hua, M., Zhang, S., Pan, B., Zhang, W., Lv, L., Zhang, Q., 2012. J. Hazard. Mater.

211–212, 317–331.Hund-Rinke, K., Simon, M., 2006. Environ. Sci. Pollut. Res. Int. 13, 225–232.Ivask, A., Bondarenko, O., Jepihhina, N., Kahru, A., 2010. Anal. Bioanal. Chem. 398,

701–716.Ivask, A., Kurvet, I., Kasemets, K., Blinova, I., Aruoja, V., Suppi, S., Vija, H., Kakinen,

A., Titma, T., Heinlaan, M., Visnapuu, M., Koller, D., Kisand, V., Kahru, A., 2014.PLoS One 9, e102108.

Jiang, W., Yang, K., Vachet, R.W., Xing, B., 2010. Langmuir 26, 18071–18077.Jomini, S., Labille, J., Bauda, P., Pagnout, C., 2012. Toxicol. Lett. 215, 54–61.Kahru, A., Dubourguier, H.C., 2010. Toxicology 269, 105–119.Kasemets, K., Ivask, A., Dubourguier, H.C., Kahru, A., 2009a. Toxicol. In Vitro 23,

1116–1122.Kasemets, K., Ivask, A., Dubourguier, H.-C., Kahru, A., 2009b. Toxicol. In Vitro 23,

1116–1122.Kasemets, K., Suppi, S., Künnis-Beres, K., Kahru, A., 2013. Chem. Res. Toxicol. 26,

356–367.Keller, A.A., McFerran, S., Lazareva, A., Suh, S., 2013. J. Nanopart. Res. 15.Kim, K.T., Klaine, S.J., Cho, J., Kim, S.H., Kim, S.D., 2010. Sci. Total Environ. 408,

2268–2272.Kumar, A., Pandey, A.K., Singh, S.S., Shanker, R., Dhawan, A., 2011. Free Radic. Biol.

Med. 51, 1872–1881.Kumar, D., Kumari, J., Pakrashi, S., Dalai, S., Raichur, A.M., Sastry, T.P., Mandal, A.B.,

Chandrasekaran, N., Mukherjee, A., 2014. Ecotoxicol. Environ. Saf. 108,152–160.

Kuznetsova, G.P., Larina, O.V., Petushkova, N.A., Kisrieva, Y.S., Samenkova, N.F.,Trifonova, O.P., Karuzina II, Ipatova, O.M., Zolotaryov, K.V., Romashova, Y.A.,Lisitsa, A.V., 2014. Bull. Exp. Biol. Med. 156, 694–698.

Lee, B., Duong, C.N., Cho, J., Lee, J., Kim, K., Seo, Y., Kim, P., Choi, K., Yoon, J., 2012. J.Biomed. Biotechnol. 2012, 262670.

Libralato, G., Minetto, D., Totaro, S., Micetic, I., Pigozzo, A., Sabbioni, E., Marcomini,A., Volpi Ghirardini, A., 2013. Mar. Environ. Res. 92, 71–78.

Lipinski, B., 2011. Oxid. Med. Cell. Longev. 2011, 809696.Lok, C.N., Ho, C.M., Chen, R., He, Q.Y., Yu, W.Y., Sun, H., Tam, P.K.H., Chiu, J.F., Che,

C.M., 2007. J. Biol. Inorg. Chem. 12, 527–534.Luoma, S.N., 2008. Silver Nanotechnologies and the Environment: Old Problems or

New Chalenges? Woodrow Wilson international center for scholars or thePEW charitable trusts, Washington DC.

Lyon, D.Y., Thill, A., Rose, J., Alvarez, P.J.J., 2007. Environmental Nanotechnology:Applications and Impacts of Nanomaterials. In: Wiesner, M., Bottero, J.-Y.(Eds.). McGraw Hill Publishing, New York, NY, pp. 445–480.

Ma, R., Stegemeier, J., Levard, C., Dale, J.G., Noack, C.W., Yang, T., Brown Jr., G.E.,Lowry, G.V., 2014. Environ. Sci. : Nano 1, 347–357.

Mahdavi, S., Jalali, M., Afkhami, A., 2012. J. Nanopart. Res. 14, 1–18.Manke, A., Wang, L., Rojanasakul, Y., 2013. BioMed Res. Int. 2013, 942916.R. Markets, The world market for nanoparticle titanium dioxide (TiO2), <http://

www.researchandmarkets.com/reports/1651709/>.McQuillan, J.S., Shaw, A.M., 2014. Biosens. Bioelectron. 51, 274–279.Merdzan, V., Domingos, R.F., Monteiro, C.E., Hadioui, M., Wilkinson, K.J., 2014. Sci.

Total Environ. 488–489, 316–324.Midander, K., Cronholm, P., Karlsson, H.L., Elihn, K., Möller, L., Leygraf, C.,

Wallinder, I.O., 2009. Small 5, 389–399.Baalousha, Mohammed, Manciulea, Adriana, Cumberland, Susan, Kendall, Kevin,

Lead, R. Jamie, 2008. Environ. Toxicol. Chem. 27, 1875–1882.Morgan, I.J., Henry, R.P., Wood, C.M., 1997. Aquat. Toxicol. 38, 147–163.Mortimer, M., Kasemets, K., Kahru, A., 2010a. Toxicology 269, 182–189.Mortimer, M., Kasemets, K., Kahru, A., 2010b. Toxicology 269, 182–189.Mortimer, M., Kasemets, K., Vodovnik, M., Marinsek-Logar, R., Kahru, A., 2011.

Environ. Sci. Technol. 45, 6617–6624.Moschini, E., Gualtieri, M., Colombo, M., Fascio, U., Camatini, M., Mantecca, P.,

2013. Toxicol. Lett. 222, 102–116.Mwaanga, P., Carraway, E.R., van den Hurk, P., 2014. Aquat. Toxicol. 150, 201–209.Nations S., Long M., Wages M., Canas J., Maul J.D., Theodorakis C., Cobb G.P.

SECOTOX Conference & Second International Conference on EnvironmentalManagement, Engineering, Planning and Economics (CEMPE 09) Pollutanteffects on aquatic and terrestrial organisms SECOTOX Conference & SecondInternational Conference on Environmental Management, Engineering,Planning and Economics, 74, 2011, 203-210.

Neale, P.A., Jamting, A.K., O’Malley, E., Herrmann, J., Escher, B.I., 2015. Environ. Sci.2, 86–93.

Nel, A., Xia, T., Mädler, L., Li, N., 2006. Science 311, 622–627.Nel, A.E., Madler, L., Velegol, D., Xia, T., Hoek, E.M.V., Somasundaran, P., Klaessig, F.,

Castranova, V., Thompson, M., 2009. Nat. Mater. 8, 543–557.Oukarroum, A., Bras, S., Perreault, F., Popovic, R., 2012. Ecotoxicol. Environ. Saf. 78,

80–85.Pan, X., Redding, J.E., Wiley, P.A., Wen, L., McConnell, J.S., Zhang, B., 2010.

Chemosphere 79, 113–116.



































































































































































































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Perreault, F., Oukarroum, A., Melegari, S.P., Matias, W.G., Popovic, R., 2012.Chemosphere 87, 1388–1394.

Perreault, F., Popovic, R., Dewez, D., 2014. Environ. Pollut. 185, 219–227.Pham, C.H., Yi, J., Gu, M.B., 2012. Ecotoxicol. Environ. Saf. 78, 239–245.Piccinno, F., Gottschalk, F., Seeger, S., Nowack, B., 2012. J. Nanopart. Res. 14.Pinheiro, J.P., Domingos, R.F., 2015. Nanomaterials for Environmental Protection.

John Wiley & Sons.Poynton, H.C., Lazorchak, J.M., Impellitteri, C.A., Smith, M.E., Rogers, K., Patra, M.,

Hammer, K.A., Allen, H.J., Vulpe, C.D., 2011. Environ. Sci. Technol. 45, 762–768.Poynton, H.C., Lazorchak, J.M., Impellitteri, C.A., Blalock, B.J., Rogers, K., Allen, H.J.,

Loguinov, A., Heckman, J.L., Govindasmawy, S., 2012. Environ. Sci. Technol. 46,6288–6296.

Rainville, L.C., Carolan, D., Varela, A.C., Doyle, H., Sheehan, D., 2014. Analyst 139,1678–1686.

Rauscher, H., Roebben, G., Amenta, V., Sanfeliu, A.B., Calzolai, L., Emons, H.,Gaillard, C., Gibson, N., Linsinger, T., Mech, A., Pesudo, L.Q., Rasmunssen, K.,Sintes, J.R., Sokull-Klüttgen, B., Stamm, H., 2014. Towards a review of the ECrecommendation for a definition of the term nanomaterial. In: Report1831–9424. Luxembourg Publication Office of the European Union.

Reed, R.B., Ladner, D.A., Higgins, C.P., Westerhoff, P., Ranville, J.F., 2012. Environ.Toxicol. Chem. 31, 93–99.

Reidy, B., Haase, A., Luch, A., Dawson, K., Lynch, I., 2013. Materials 6, 2295–2350.Research B. Global Markets for Nanocomposites, Nanoparticles, Nanoclays and

Nanotubes 2015; <http://www.bccresearch.com/market-research/nanotechnology/nanocomposites-market-nan021f.html?vsmaid=203/>.

Rippka, R., Deruelles, J.B., Herdman, M., Waterbury, B., Stanier, R.Y., 1979. J. Gen.Microbiol. 111, 1–61.

Saison, C., Perreault, F., Daigle, J.C., Fortin, C., Claverie, J., Morin, M., Popovic, R.,2010. Aquat. Toxicol. 96, 109–114.

Sakai, N., Matsui, Y., Nakayama, A., Tsuda, A., Yoneda, M., 2011. J. Phys.: Conf. Ser.304, 012049.

Savolainen, K., Backman, U., Brouwer, D., Fadeel, B., Fernandes, T., Kuhlbusch, T.,Landsiedel, R., Lynch, L., Pylkkänen, I., 2013. Nanosafety in Europe 2015–2025Towards Safe and Sustainable Nanomaterials and NanotechnologyInnovations. Finnish Institute of Occupational Health.

Schultz, A.G., Ong, K.J., MacCormack, T., Ma, G., Veinot, J.G., Goss, G.G., 2012.Environ. Sci. Technol. 46, 5–10301.

Schultz, A.G., Boyle, D., Chamot, D., Ong, K.J., Wilkinson, K.J., McGeer, J.C., Sunahara,G., Goss, G.G., 2014. Environ. Chem. 11, 207.

Shang, L., Nienhaus, K., Nienhaus, G.U., 2014. J. Nanobiotechnol. 5, 1–12.Sheela, T., Nayaka, Y.A., Viswanatha, R., Basavanna, S., Venkatesha, T.G., 2012.

Powder Technol. 217, 163–170.Shepard, J.L., Olsson, B., Tedengren, M., Bradley, B.P., 2000. Mar. Environ. Res. 50,

337–340.

Shi, J., Abid, A.D., Kennedy, I.M., Hristova, K.R., Silk, W.K., 2011. Environ. Pollut. 159,1277–1282.

Shi, J., Xu, B., Sun, X., Ma, C., Yu, C., Zhang, H., 2013. Aquat. Toxicol. 132–133, 53–60.Simon, D.F., Domingos, R.F., Hauser, C., Hutchins, C.M., Zerges, W., Wilkinson, K.J.,

2013. Appl. Environ. Microbiol. 79, 4774–4785.Sun, H., Zhang, X., Niu, Q., Chen, Y., Crittenden, J.C., 2006. Water Air Soil Pollut. 178,

245–254.Suresh, P., Vijaya, J.J., Kennedy, L.J., 2015. React. Kinet. Mech. Catal. 114, 767–780.Tan, C., Wang, W.X., 2014. Environ. Pollut. 186, 36–42.Thalmann, B., Voegelin, A., Sinnet, B., Morgenroth, R., Kaegi, E., 2014. Environ. Sci.

48, 4885–4892.Tiede, K., Boxall, A.B., Tear, S.P., Lewis, J., David, H., Hassellov, M., 2008. J. Hazard.

Mater. 25, 795–821.Vale, G., Franco, C., Diniz, M.S., Santos, M.M.C., Domingos, R.F., 2014. Ecotoxicol.

Environ. Saf. 109, 161–168.Wang, Z., Bussche, A.V.D., Kabadi, P.K., Kane, A.B., Hurt, R.H., 2013a. ACS Nano 7,

8715–8727.Wang, Z., Bussche, A.V.D., Kabadi, P.K., Kane, A.B., Hurt, R.H., 2013b. ACS Nano 10,

8715–8727.Wang, B., Yin, J.-J., Zhou, X., Kurash, I., Chai, Z., Zhao Feng, Y.W., 2013c. J. Phys.

Chem. C 117, 383–392.Wang, L.-F., Habibul, N., He, D.-Q., Li, W.-W., Zhang, X., Jiang, H., Yu, H.-Q., 2015.

Water Res. 68, 12–23.Xia, T., Zhao, Y., Sager, T., George, S., Pokhrel, S., Li, N., Schoenfeld, D., Meng, H., Lin,

S., Wang, X., Wang, M., Ji, Z., Zink, J.I., Mädler, L., Castranova, V., Lin, S., Nel, A.E.,2011. ACS Nano 5, 1223–1235.

Xiong, D., Fang, T., Yu, L., Sima, X., Zhu, W., 2011. Sci. Total Environ. 409,1444–1452.

Yu, L.-p., Fang, T., Xiong, D.-w, Zhu, W.-t, Sima, X.-f., 2011. J. Environ. Monit. 13,1975–1982.

Zhang, L., Jiang, Y., Povery, Y., York, D., 2007a. J. Nanopart. Res. 9, 479–489.Zhang, X., Sun, H., Zhang, Z., Niu, Q., Chen, Y., Crittenden, J.C., 2007b. Chemosphere

67, 160–166.Zhang, Y., Gao, X., Zhi, L., Liu, X., Jiang, W., Sun, Y., Yang, J., 2014. J. Inorg. Biochem.

130, 74–83.Zhao, X., Wang, S., Wu, Y., You, H., Lv, L., 2013. Aquat. Toxicol. 136–137, 49–59.Zhu, X., Zhu, L., Duan, Z., Qi, R., Li, Y., Lang, Y., 2008. J. Environ. Sci. Health Part A 43,

278–284.Zhu, X., Wang, J., Zhang, X., Chang, Y., Chen, Y., 2009. Nanotechnology 20, 195103.Zhu, X., Chang, Y., Chen, Y., 2010. Chemosphere 78, 209–215.von Moos, N., Maillard, L., Slaveykova, V.I., 2015. Aquat. Toxicol. 161, 267–275.
























































































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Annexe 2: Identification of Gammarus sp. in Luxembourg streams

1. Sampling sites:

Gammarus sp. were collected from different unpolluted sites in Luxembourg (Figure 1). Animals were

collected using a hand net. They were quickly transported to the laboratory where they were kept at 12 °C.

Only adults were selected. Ten individuals from each site were kept and frozen in liquid nitrogen and

conserved at – 80 °C until analysis.

Figure 1: Unpolluted rivers in Luxembourg. Stars indicates sites where Gammarus sp. were collected for

molecular identification

Schwaarzbaach

252

2. Gammarus sp. identification

2.1. Genomic DNA extraction and PCR

Gammarids were digested overnight at 56°C in lysis buffer supplemented with proteinase K (10% v/v).

Genomic DNA (gDNA) was extracted using QIAmp DNA mini kit® (QIAgen), following manufacturer’s

instructions. The extracted DNA was quantified using Nanodrop. Primers for Gammarus sp. were designed

based on a set of six pairs of universal primers: COI, 12S, 16S, 18S and 28S (Berschick, 1997; Hou et al., 2007).

Primers used for amplification are listed in Table. 1.

Table 1: primers used for amplification and sequencing

Gene Primer Sequence (5’ – 3’) Reference

COI * HCO2198_F TAAACTTCAGGGTGACCAAAAAATCA

Folmer et al., 1994 LCO1490_R GGTCAACAAATCATAAAGATATTGG

12S 12S-2-SSU _ F GTGGATCCATTAGATACCC Berschick, 1997 12S-2-SSU_ R ACTGGTACCTTGTTACGACTT Hou et al., 2007

16S 16ST_F GGTAWHYTRACYGTGCTAAG MacDonald et al., 2005 16SB_R CCGGTTTGAACTCAGATCATGT Palumbi, 1996

18S 18S_ F CCTAYCTGGTTGATCCTGCCAGT

Englisch et al., 2003) 18S_R TAATGATCCTTCCGCAGGTT

28S 28S _F TTAGTAGGGGCGACCGAACAGGGAT

Hou et al., 2007 28S_R GTCTTTCGCCCCTATGCCCAACTGA

* COI : cytochrome c oxidase subunit 1

Polymerase chain reactions (PCR) were performed in a reaction volume of 25 µL containing 12.5 µL of 2X Q5®

master mix, 1.25 µL of each primer (0.5 µM) and 10µL of gDNA (1ng). PCR settings were:

2.2. Agarose gel electrophoresis and DNA profile analysis

Electrophoresis of PCR products was performed on a 3% (w/v) agarose gel containing ethidium bromide and

they were visualised under UV light. The PCR product sizes were determined by using Mass ruler TM DNA

ladder Mix ready to use #SM0403.

Denaturation 98°C 5min

Denaturation 98°C 30s

Hybridization 55°C 60s

Elongation 72°C 60s

Final extension 72°C 5 min

Cooling 4°C ∞

35 cycles

253

2.3. PCR purification and sequencing

PCR products were purified by classical ethanol precipitation method (Cambier et al., 2010). The purified DNA

fragments were quantified using Nanodrop, then amplified and purified using BigDye Xterminator® kit

following manufacturer instructions. The obtained fragments were sequenced using ABI genetic Analyser

3500.

3. Results

3.1. DNA profile analysis

One Gammarus was used to test the primers. Among the five-targeted genes, 18S was not amplified, and

many PCR products were obtained for 28S (Figure 2). This could be linked to the low specificity of the primers

allowing the amplification of various gDNA regions. COI, 12S and 16S genes were successfully amplified. Low

intensity fragments were obtained for COI (800bp) and 12S (800bp). An amplified fragment of 400bp was

obtained for 16S gene (Figure 2). Therefore, these 3 genes were selected for further steps.

Figure 2: Agarose gel electrophoresis of amplification products generated with universal primers designed for

Gammarus sp. White arrows indicate the selected genes for sequencing analyses. Black arrows indicate the size

of the amplified fragments

Low intensity PCR products of about 800bp of COI (Figure 3) and 12S (Figure 4) were obtained for 10

Gammarus. This confirms the low efficiency of these two primers for the amplification of COI and 12S genes

of Gammarus sp.

L L 28S COI 12S 18S 16S

800 bp

400 bp

254

Figure 3: Agarose gel electrophoresis of amplification products generated with COI primers for Gammarus sp.

Arrows indicate the amplified fragments.

Figure 4: Agarose gel electrophoresis of amplification products generated with 12S primers for Gammarus sp.

Arrows indicate the amplified fragments

Fragment of about 400 bp with a high intensity were obtained for the amplification of 16S gene for all

individuals (Figure 5). This gene was kept for the sequencing analysis and identification step.

Figure 5: Agarose gel electrophoresis of amplification products generated with 16S primer pair for Gammarus

sp.

L L L G1 G2 G3 G4 G5 G6 G7 G8 G9 G10

L L L G1 G2 G3 G4 G5 G6 G7 G8 G9 G10

L L L G1 G2 G3 G4 G5 G6 G7 G8 G9 G10

255

3.2. Sequencing results

Sequencing analysis (Figure 6) showed that the obtained PCR products, using 16S primers, corresponded to

~400bp mitochondrial partial rRNA16S gene.

Figure 6: Gammarus fossarum 16S chromatogram obtained with ABI genetic analyser 3500.

The obtained sequences were analysed using the Blastn program available on the NCBI website

(http://blast.st-va.ncbi.nlm.nih.gov/Blast.cgi) using the the GenBank database. Results are showed in table

2.

Homology of 97% to 100% with very low e-value were obtained for all 10 Blastn analysis. These Blastn results

on these 10 DNA sequences allowed to identify the Gammarus sp. as Gammarus fossarum. Furthermore,

among these 10 animals 9 were identified as Gammarus fossarum type B, and only one (G2) was

characterised as a Gammarus fossarum type C. These results show that the Gammarus species present in The

Attert River at Colmar berg (Luxembourg) is Gammarus fossarum with a majority of the cryptic species B.

256

Table 2: Species identification and GenBanK Accession number

Individual Gene Description Query

cover E- value Identification

Accession

number

G1 16S Gammarus fossarum type B

mitochondrial partial rRNA 16S gene 84% 2e-155 100% AJ269607.1

G2 16S Gammarus fossarum type C


















257

References

Berschick, P. (1997). One primer pair amplifies small subunit ribosomal DNA from mitochondria, plastids and

bacteria. Mitochondria, plastids and bacteria. BioTechniques 23, 494–498.

Cambier, S., Gonzalez, P., Durrieu, G., and Bourdineaud, J.-P. (2010). Cadmium-induced genotoxicity in zebrafish

at environmentally relevant doses. Ecotoxicol. Environ. Saf. 73, 312–319.

Englisch, U., Coleman, C.O., and Wägele, J.W. (2003). First observations on the phylogeny of the families

Gammaridae, Crangonyctidae, Melitidae, Niphargidae, Megaluropidae and Oedicerotidae (Amphipoda,

Crustacea), using small subunit rDNA gene sequences. J. Nat. Hist. 37, 2461–2486.

Folmer, O., Black, M., Hoeh, W., Lutz, R., and Vrijenhoek, R. (1994). DNA primers for amplification of

mitochondrial cytochrome c oxidase subunit I from diverse metazoan invertebrates. Mol. Mar. Biol. Biotechnol.

3, 294–299.

Hou, Z., Fu, J., and Li, S. (2007). A molecular phylogeny of the genus Gammarus (Crustacea: Amphipoda) based

on mitochondrial and nuclear gene sequences. Mol. Phylogenet. Evol. 45, 596–611.

MacDonald, K.S., Yampolsky, L., and Duffy, J.E. (2005). Molecular and morphological evolution of the amphipod

radiation of Lake Baikal. Mol. Phylogenet. Evol. 35, 323–343.

Palumbi, S.R., (1996). Nucleic acids II: The polymerase chain reaction.

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