Marine Biofilms: Targets of Cyanobacterial

Home , Biofouling, Marine microorganisms

Marine biofilms: targets of cyanobacterial

compounds towards a natural D antifouling strategy

Jorge Tiago Vieira Ferreirinha Antunes Doutoramento em Biologia Departamento de Biologia Ano de 2019

Orientador Doutor Pedro Leão, CIIMAR Coorientador Prof. Doutor Vitor Vasconcelos, FCUP and CIIMAR

Marine biofilms: targets of cyanobacterial bioactive compounds

towards a natural antifouling strategy

SFRH/BD/99003/2013

Jorge Tiago Vieira Ferreirinha Antunes

Blue Biotechnology and Ecotoxicology (BBE) and Cyano Natural Products (CNP) research teams

Interdisciplinary Centre of Marine and Environmental Research (CIIMAR)

Supervisor: Pedro Leão, CIIMAR Co-supervisor: Professor Vítor Vasconcelos, FCUP and CIIMAR

Faculty of Sciences of University of Porto, 2019

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Jorge Antunes acknowledges a PhD scholarship SFRH/BD/99003/2013 from FCT

(Fundação para a Ciência e Tecnologia).

This work was supported by FCT - Foundation for Science and Technology - under project

UID/Multi/04423/2013 and by the Structured Program of R&D&I INNOVMAR - Innovation and Sustainability in the Management and Exploitation of Marine Resources (reference

NORTE-01-0145-FEDER-000035, Research Line NOVELMAR), which was funded by the

Northern Regional Operational Programme (NORTE2020) through the European Regional

Development Fund (ERDF) and by the project CVMAR+I (0302_CVMAR_I_1_P) funded by the program Interreg V Portugal-Spain (POCTEP) through the ERDF.

Funding was also provided by the project NASCEM PTDC/BTA-BTA/31422/2017 (POCI-

01-0145-FEDER-031422), financed by the FCT, COMPETE2020 and PORTUGAL2020.

Personal acknowledgements

Writing this dissertation was, for me, one of the most challenging tasks I ever endeavored, one I would surely fail if not for the help of a number of wonderful people I was lucky enough to meet.

In particular, I would like to sincerely thank my supervisor, Doutor Pedro Leão, the best guide I could ever hope for his insightful, good humor– and always illuminating – commentaries. I also thank him for his constant thoroughness and constant readiness to help. His willingness to give me enough scientific independence during this project of discovery, and to challenge me for always strive for the best, surely allowed me to grow as a researcher and as a scientist.

I would like also to heartily thank my co-supervisor, Professor Doutor Vitor Vasconcelos for accepting my proposal to conduct this thesis in a such an interesting topic, and for welcoming me again in such a dynamic, good-spirited and exceptional team. His helpfulness, generosity and scientific knowledge along these last years, made me joyful to work in such an outstanding institution such as CIIMAR.

I also thank all my colleagues particularly in the BBE and CNP teams, as well in other CIIMAR teams for all their support, and friendship who are unfortunately too many to mention all by name. I would like to mention, however some who helped me more directly both in the lab, as well in fruitful scientific discussions, and who helped me to carry on through this difficult process until the end. Therefore, a special mention to Anoop, Aldo, António, Tito, Cidália, Adriana and Teresa. An even more special mention to Joana, my closest teammate during these last years, for her friendship and help during our marine fouling scientific projects, the best colleague I could hope for.

Finally, I would like to thank my family for their unconditional support and patience, particularly my parents for their understanding and love, to whom I dedicate this thesis, and who are the true biologists.

Abstract

Marine biofouling remains an unresolved problem with serious economic impact for different industries. Marine biofilms are the initial step of the marine fouling process in artificial surfaces and are the preferential substrata for the settlement of macroinvertebrate larvae and macroalgae spores. In this thesis, an extensive and critical literature review was carried out to digest current information on the distribution of marine biofilms, the already known chemical cues involved in their formation, as well the overall importance of marine biofilms for the marine fouling process. In addition, to understand how the marine biofilm formation process evolves, we employed amplicon-based NGS (Next-generation sequencing) methodologies to characterize early-stage marine biofilms growing on steel surfaces in two distinct seasons in the port of Leixões in Northern Portugal. The 16S rRNA gene sequencing data analysis demonstrated distinct taxonomic and functional profiles of the marine biofilm communities in the sampled seasons, suggesting that seasonality is a major factor influencing the diversity of those prokaryotic communities. Furthermore, the sampled biofilms revealed the presence of pathogenic taxa which, accordingly to our knowledge was not reported previously, and whose abundance was significantly influenced by temporal variations. This knowledge should be important for future anti-fouling and anti-corrosion purposes. After years of application of toxic antifoulants to control biofouling, there is a strong necessity for the development of environmentally friendly antifouling products, and cyanobacteria are a particularly encouraging source of new natural antifouling compounds. An extensive screening was therefore performed with crude organic extracts of 61 cyanobacterial strains of the LEGE Culture Collection (LEGE CC) to test their anti-fouling potential. The obtained fractions of several cyanobacterial strains revealed significant activity against the growth of microfouling organisms involved in marine biofilm formation (bacteria and diatoms) and inhibited the quorum-sensing (QS) phenomenon, which is associated with biofilm formation.

Subsequent work resulted in a set of HPLC-PDA purified bioactive fractions from a marine cyanobacterial strain whose structure is being elucidated by spectroscopic methods.

Additionally, the already known cyanobacterial compounds portoamides, isolated from a marine cyanobacterium from the LEGE CC, were also tested for their antifouling potential.

A multi-bioassay approach revealed a wide spectrum antimicrofouling, anti- macroinvertebrate settlement, and anti-biofilm activity of those compounds, while displaying no toxicity to target and non-target organisms, suggesting that portoamides have a potential to be used as new environmentally friendly antifouling agents. We have also optimized an anti-biofilm assay protocol that should be of interest for the discovery of bioactive compounds with anti- marine biofilm activity which is indispensable for the discovery of efficient broad-spectrum antifouling compounds.

Keywords: anti-biofilm; antifouling; bioactive compounds; cyanobacteria compounds; marine biofilms; marine biofouling; next-generation-sequencing; quorum-sensing.

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Resumo

As camadas incrustantes causadas por organismos marinhos são um problema com grandes consequências económicas para diferentes indústrias. Os biofilmes marinhos são o primeiro passo no processo de colonização das camadas incrustantes em superfícies artificiais, e constituem superfícies preferenciais para a colonização e adesão de larvas de macroinvertebrados, e de esporos de macrogalgas. Neste projeto de doutoramento, um artigo de revisão foi escrito para discutir a distribuição de biofilmes marinhos, os compostos químicos envolvidos na sua formação, assim como a importância dos biofilmes no contexto das camadas incrustantes marinhas. A tecnologia de sequenciação de nova geração foi adotada para caracterizar os biofilmes marinhos que cresceram em superfícies de metal em duas estações distintas do ano no porto de Leixões, no Norte de Portugal. A análise dos dados de sequenciação de 16S RNA demonstrou um perfil taxonómico e funcional distinto nas duas estações do ano, sugerindo que o tempo e a estação do ano influenciam fortemente a composição taxonómica e funcional desses biofilmes. Adicionalmente, foram identificados nesses biofilmes grupos taxonómicos associados a situações patogénicas que, segundo o nosso conhecimento, não tinham sido descritos em estudos anteriores sobre biofilmes marinhos e cuja abundância foi significativamente influenciada pelas variações temporais testadas. Esta informação pode ter relevância para futuras estratégias eficientes de controlo de crescimento de camadas incrustantes e de biofilmes marinhos. Depois de anos de aplicação de produtos anti-incrustante tóxicos, existe uma grande necessidade do desenvolvimento de produtos que não sejam danosos para o ambiente, e as cianobactérias são uma fonte particularmente relevante para a descoberta de novos compostos naturais devido ao potencial bioativo dos seus metabolitos secundários. Neste projeto de doutoramento, um rastreio abrangente foi efetuado com os crudes de 61 estirpes de cianobactérias da coleção de culturas do LEGE (LEGE CC) para identificar o seu potencial anti-incrustante. Várias estirpes demonstraram uma significativa bioatividade contra bactérias marinhas e diatomáceas, e

viii inibiram igualmente o fenómeno de percepção de quórum (quorum-sensing). Trabalho posterior levou à obtenção de uma fração bioativa purificada por HPLC-PDA, cuja estrutura será elucidada por métodos de espectrometria de massa de alta resolução. Adicionalmente, os já conhecidos compostos de cianobactérias, portoamidas, isolados de uma cianobactéria pertencente à LEGE CC, também foram testados contras os mesmo painéis de organismos.

Uma série de ensaios distintos revelou uma abrangente atividade destes compostos, assim como uma significativa atividade disruptiva de biofilmes marinhos, não demonstrando toxicidade para organismos alvo e não alvo. Isto sugere que as portoamidas têm um potencial significativo para serem usados como compostos anti incrustação/anti-biofilmes não danosos para o ambiente. Também foi otimizado um novo protocolo anti-biofilme que pode ser de interesse para a descoberta de novos compostos bioativos com atividade disruptiva contra biofilmes marinhos, o que é necessário para identificar compostos anti-incrustantes eficientes e de largo espectro.

Palavras chave: anti-biofilme; biofilmes marinhos; camadas incrustantes; compostos bioactivos naturais; compostos de cianobactérias; percepção de quórum; sequenciação de nova geração.

Table of Contents

Ackowledgments…………………………………………………………………...... ii

Personal acknowledgements...... iv

Abstract ……………………………………………………………...... v

Resumo ……………………………………………………………...... vii

Table of contents …………………………………………………………………...... ix

List of Tables ……………………………………………………………...... xiv

List of Figures ...………………………………………………………………...... xvi

Abbreviations list………………………………… ...... xxii

1. Chapter 1: General Introduction…….……………………...... 1

1. 1 Marine biofouling: an introduction...... ………………………...... 2

1.2 Marine biofouling: major problems and consequences...... 6

1.3 Antifouling strategies ...... 9

1.4 Antifouling agents of natural origin: current products and future needs………………...... 10

1.5 Cyanobacteria as a source of antifouling agents…...... …………...... 12

1.6 Structure and main objectives...... ………………………...... 14

2. Chapter 2: “Marine biofilms: diversity of communities and of chemical cues” ...... 18

2.1. Introduction...... 20

2.2 Diversity and distribution of marine biofilm communities...... 22

2.3 Marine biofilm formation...... 25

2.4 Marine biofilms and the influence of environmental factors...... 28

2.5 Marine biofilms and chemical cues...... 30

2.6 Marine biofilms and their relation to marine fouling...... 40

2.7 Quorum-sensing inhibition as a strategy to prevent marine biofilm growth.…………...... 43

2.8 Concluding remarks…………………………………...... 46

2.9. References………………………………………………………...... 47

3. Chapter 3: “Distinct profile of early marine biofilms in two different seasons in a Portuguese Atlantic Port” …………………………………………………………………………………...... 65

3.1. Introduction ……………………………………………………………………...... 67

3.2. Material and Methods………………………………………………………...... 70

3.2.1 Study site and sampling……………………………………...... 70

3.2.2 DNA Sequencing and Sequence analysis ……………………………...... 71

3.2.3 PICRUSt functional predictions ...... 72

3.2.4 Classification of chloroplast sequences...... 73

3.3Results...... 74

3.3.1Environmental parameters...... 74

3.3.2 Alpha and Beta-diversity...... 75

3.3.3 Taxonomic temporal succession………………………………………………...... 77

3.3.4 Functional analysis……………………………………...... 84

3.4 Discussion …………………………………………………………………………...... 86

3.4.1 Alpha-Diversity and Beta-diversity………………………………………...... 87

3.4.2 Taxonomic temporal succession ……………………………………………...... 88

3.4.3 Functional analysis……………………………………………………...... 96

3.5. Conclusions………………………………………………………………………...... 98

3.6 References……………………………………………………………………………...... 100

4. Chapter 4: “A multi-bioassay integrated approach to assess the antifouling potential of the cyanobacterial metabolites portoamides” ...... 110

4.1. Introduction………………………………………………………………………...... 112

4.2. Results and Discussion...……………………………………………………………...... 114

4.2.1. Mytilus galloprovincialis Larvae Antisettlement Activity and Recovery...... 114

4.2.2. Antibacterial and Antimicroalgal Activities...... 116

4.2.3. Antibiofilm Activity...... 119

4.2.4. Quorum-Sensing Inhibition (QSI)Activity...... 121

4.2.5.Ecoxocity assesment...... 122

4.2.6.Mode of action assessment...... 123

4.3. Materials and Methods………………………………………………………...... 127

4.3.1. Portoamides Isolation and Purification...... 127

4.3.2. Antifouling Bioassays...... 127

4.3.2.1. Mussel Larvae Antisettlement Bioassays...... 127

4.3.2.2. Mussel Larvae Settlement Recovery Bioassays...... 128

4.3.2.3. Antibacterial Bioassays...... 128

4.3.2.4. Antimicroalgal Bioassays...... 129

4.3.2.5. Biofilm Inhibition Assay...... 129

4. 3.2.6. Quorum-Sensing Inhibition Assay...... 130

4.3.2.6.1 Disk-Diffusion Assay...... 130

4.3.2.6.2 Quorum-Sensing Inhibition (QSI) Quantitative Assay (Dose–Response Violacein Inhibition Assay) ...... 131

4.3.3. Ecotoxicity Bioassay...... 132

4.3.4. Antifouling Mode of Action Towards Mytilus Larvae...... 132

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4.3.4.1. Sample Preparation……………………………………...... 133

4.3.4.2. LC-MS/MS Analysis…………………………………………...... 133

4.3.4.3 Protein Identification and Quantification………………………...... 133

4.3.5. Data Analysis...... 134

4.4. Conclusions……………………………………………………………………...... 134

4.5. References ………………………………………………………………………...... 136

5. Chapter 5: Search for cyanobacterial compounds with antimicrofouling and quorum sensing inhibition activity………………………………………………………………………………….144

5. 1. Introduction ...... 146

5.2 Materials and methods...... 147

5.2.1. Cyanobacteria growth and primary screening...... …………………….…...... 147

5.2.2. Extraction, purification and identification of antifouling compounds...... 150

5.2.3 Anti-microfouling bioassays...... 152

5.2.3.1 Bioassay with fouling marine bacteria………………………...... 152

5.2.3.2. Bioassay with fouling diatoms ………………………………...... 152

5.2.4.1 Quorum sensing (QS) inhibition test …………………………...... 153

5.2.4.2. Quorum-sensing inhibition quantitative assay (Dose-response violacein inhibition assay) ...... 154

5.2.5 Antibacterial screening susceptibility assay ……………………………...... 155

5.2.6 Assay with macroalgae ...... 155

5.2.6.1. Induction of algae spore release ...... ………………………...... 156

5.2.6.2. Bioassay with macroalgae spores ………………………...... 157

5.3. Results and discussion ...... 157

5.3.1. Screening results for the bioassay with microfouling organisms and quorum-sensing inhibition …………………………………………………………………………...... 158

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5.3.2. Results for the assay with marine macroalgae...... 160

5.3.3. Bioactivity-guided isolation of antifouling compounds...... 161

5.4. Main conclusions …………………………………………………………………...... 169

5.5 References....…………………………………………………………………………...... 170

6. Main Conclusions ……………………………………………………………………………....175

7. References ……………………………………………………………………………………...178

8- Publications, conference proceedings and attended courses …………………………………...188

9- Appendix ……………………………………………………………………………………….190

9.1 From Chapter 3 “Distinct profile of early marine biofilms in two different seasons in a Portuguese Atlantic Port” ………………………………………………………………………....191

9.2 From Chapter 4 “A multi-bioassay integrated approach to assess the antifouling potential of the cyanobacterial metabolites portoamides” ...... 191

9.3 Chapter 5 “Search for cyanobacterial compounds with antimicrofouling and quorum sensing inhibition activity” ………………………………………………………………………...... 195

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List of Tables

Chapter 2

Table 1. Non-QS chemical cues from marine biofilms and their relation to affected marine organisms. Page 33

Table 2. Quorum-sensing molecules from marine biofilms and their relation to affected marine organisms. Page 36

Chapter 3

Table 1 - Environmental parameters of the seawater at the Leixões Port at each of the sampled days. Page 74

Chapter 4

Table 1. Bacterial growth inhibition activity of portoamides. Page 117

Table 2. Differentially expressed proteins (p-value < 0.05) in Mytilus galloprovincialis larvae exposed to portoamides (16 M). Besides the mean log10-transformed abundance values, the names of the identified proteins and the main metabolic processes involved are also listed in the table. Protein accession numbers are from NCBI.

Page 124

Chapter 5

Table 1 - LEGE culture collection strains for which biomass extraction has been carried out. Page 148

Table 2: Cyanobacterial strains selected for mass culturing and subsequent extraction and fractioning. Page 162

Table 3: Cyanobacterial strains grown at large-scale and VLC extracted. Page 164

APPENDIX

Supplementary Table 1: Summary of LC-MS/MS protein Identification results. The QTOF data from the Impact HD mass spectrometer equipped with a CaptiveSpray source (Bruker Daltonik, Bremen, Germany) were searched using the Peaks Studio 8.5 search algorithm (Bioinformatics Solutions, Waterlo, ON, Canada). Page 191

Supplementary table 2. Screening results for the crude cyanobacterial extracts against bacteria and diatom growth and against quorum-sensing inhibition. Page 195

Supplementary table 3: Activity for fractions and subfractions of LEGE 00060 (E16156) Page 198

Supplementary table 4: Activity for fractions of LEGE 00247 (E 17159) Page 199

Supplementary table 5: Activity for fractions of LEGE 00246 (E17162) Page 200

Supplementary table 6: Activity for fractions of LEGE 11428 (Subfractioning of E17163) Page 201

Supplementary table 7: Activity for fractions of LEGE 03273 (E17166) Page 201

Supplementary table 8: Activity for fractions of LEGE 11428 (Subfractioning of E17163) Page 202

Supplementary table 9: Activity for fractions of LEGE 11428 (Subfractioning of E17163) Page 203

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List of figures

Chapter 1

Figure 1: Evolution of marine fouling stages (adapted from Maki and Mitchell, 2002). Page 2

Figure 2: Marine biofilm development in the marine environment (adapted from Fuqua et al., 2001). Page 3

Figure 3: Main macrofouling organisms colonizing artificial surfaces (Adapted from Bressy and Lejars, 2014). Page 7

Chapter 2

Figure 1: Global map with dominant taxa characterized in marine biofilm studies, their respective sampling sites, sampled surfaces and sampled periods. Page 24

Figure 2: Marine biofilm formation stages and respective influence of environmental parameters and chemical cues. Page 26

Figure 3: Non-quorum sensing chemical cues identified in marine biofilms. Chemical formulas names and sub-numbers: (1) tetrabromopyrrole; (2) 6,9-heptadecadiene; (3) 12- octadecenoic acid; (4) histamine. See Table 1 for the interaction of each of these non-QS chemical cues with macrofouling species and their affected mechanisms. Page 34

Figure 4: Quorum-sensing chemical cues identified in marine biofilms. Chemical formulas names and sub-numbers: (5) N-hexanoyl-L-homoserine lactone; (6) N-(3-oxodecanoyl) -L- homoserine lactone; (7) N-butanoyl-L-homoserine lactone; (8) N-dodecanoyl-L-homoserine lactone; (9) N-(3-oxobutanoyl)-L-homoserine lactone; (10) N-(3-oxododecanoyl)-L- homoserine lactone;(11) N-octanoyl-L-homoserine lactone; (12) N-(3-oxodecanoyl)-L- homoserine lactone. See Table 2 for the interaction of each of these QS chemical cues with macrofouling species and their affected mechanisms. Page 35

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Figure 5: Influence of quorum sensing (presence of quorum sensing molecules) and quorum quenching processes (presence of quorum quenching enzmyes) resulting in the formation or non-formation of marine biofilms. Page 39

Chapter 3

Figure 1A and 1B: Alpha-diversity values of samples distributed by the two seasons season (Figure 1A) and treatment per season (Figure 1B). Page 76

Figure 2A and Figure 2B: Principal Coordinate Analysis (PCoA) of unweighted UniFrac distances of prokaryote 16S OTUs of the samples plotted by treatment (Figure 2A) and by season (Figure 2B). Page 77

Figures 3A and 3B: Taxonomic diversity and relative abundance at the phylum level of prokaryotes from the samples collected at spring (Figure 3A) and at winter (Figure 3B). Sequences were taxonomically assigned to the Greengenes database (DeSantis et al., 2006) using QIIME (Caporaso et al., 2010). For clarity of presentation, unassigned sequences and phyla comprising <1% of the total number of sequences within a sample were simply classified as “Other”. Page 81

Figure 4A and 4B: Taxonomic diversity and relative abundance at the class level of prokaryotes from the samples collected at spring (Figure 4A) and at winter (Figure 4B). Sequences were taxonomically assigned to the Greengenes database (DeSantis et al., 2006) using QIIME (Caporaso et al., 2010). For clarity of presentation, unassigned sequences and classes comprising <1% of the total number of sequences within a sample were simply classified as “Other”. Page 82

Figure 5A: Taxonomic diversity and relative abundance at the order level of prokaryotes from the samples collected at spring (Figure 5A). Sequences were taxonomically assigned to the Greengenes database (DeSantis et al., 2006) using QIIME (Caporaso et al., 2010). For clarity of presentation, unassigned sequences and orders comprising <1% of the total number of sequences within a sample were simply classified as “Other”. Page 83

xviii clarity of presentation, unassigned sequences and orders comprising <1% of the total number of sequences within a sample were simply classified as “Other”. Page 83

Figure 6: Taxonomic diversity and relative abundance of the Greengenes “chloroplast” sequences at the last day of sampling (day 30). Sequences were classified by the NCBI and PhytoREF databases at the lowest possible taxonomic ranks and chosen based on the best match. For clarity of presentation, unassigned sequences comprising <1% of the total number of sequences within a sample were simply classified as “Other”. Page 83

Figure 7: Relative abundance of 16S rRNA based predictive the main metagenomic functions of in the spring samples, as assigned by KEGG category database (L2) through PICRUSt analysis. Page 85

Figure 8: Relative abundance of 16S rRNA based predictive the main metagenomic functions of in the winter samples, as assigned by KEGG category database (L2) through PICRUSt analysis. Page 85

Figure 9: Relative abundance of 16S rRNA based predictive metagenomic functions of biofilm associated functions in the spring samples as assigned by KEGG category database (L3) through PICRUSt analysis. Page 86

Figure 10: Relative abundance of16S rRNA based predictive metagenomic functions of biofilm associated functions in the winter samples, as assigned by KEGG category database (L3) through PICRUSt analysis. Page 86

Chapter 4

Figure 1. Dose–response anti-settlement activity of portoamides towards plantigrade larvae of the mussel Mytilus galloprovincialis. B: DMSO control (0.01%); C: 5 µM CuSO4 as the positive control. Page 114

Figure 2. Settlement response of plantigrade larvae of the mussel Mytilus galloprovincialis after 15 h of exposure to portoamides followed by 15 h of recovery in filtered seawater. B: DMSO control (0.01%); C: 5 µM CuSO4 as the positive control. Page 116

Figure 3. Antibacterial activity of portoamides towards five biofilm-forming marine bacteria Cobetia marina, Pseudoalteromonas atlantica and Halomonas aquamarina. B: 0.1% DMSO; C: 1:100 dilution of penicilin-streptomycin-neomycin stabilized solution (Sigma P4083). Page 117

Figure 4. Antimicroalgal activity of portoamides at a concentration of 6.5 µM towards four biofilm-forming marine diatoms Cylindrotheca sp., Halamphora sp., Nitzschia sp. and

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Navicula sp. (B: 0.1% DMSO); 3.55 µM cycloheximide was used as the positive control (C+). Page 118

Figure 5. Antibiofilm dose–response activity of portoamides towards the marine bacteria Halomonas aquamarina and Pseudoalteromonas atlantica. B: 0.1% DMSO; C: 4:100 dilution of penicilin-streptomycin-neomycin stabilized solution (Sigma P4083). Page 119

Figure 6. Ratio of the quantification of growth inhibition of Chromobacterium violaceum ATCC 12472 (read at 720 nm) when compared to quantification of the production of violacein by the same bacterium (577 nm). B: 0.1% DMSO; C: 1:100 dilution of penicilin-streptomycin-neomycin stabilized solution (Sigma P4083). Page 122

Figure 7. Mortality rate of Artemia salina nauplii after 48 h of exposure to portoamides. B: 1% DMSO in filtered seawater. C: K2Cr2O7 at a concentration of 13.6 µM. Page 123

Chapter 5

Figure 1. LB agar plate representative of a quorum-sensing inhibition bioassay. The assay uses Chromobacterium violaceum ATCC 12742 and depicts both quorum-sensing inhibition with non-pigmented zones (4), and antimicrobial activities with translucent zones (1 and 3). Translucent zones show inhibition of bacterial growth, while opaque zones represent inhibition of violacein production with viable bacterial growth. (1) denotes positive control, (2) negative control, (3) crude extract from LEGE strain 91342 and (4) crude extract from LEGE strain 06014, respectively. Page 154

Figure 2. a) Ulva lactuca thallus after the release of spores, in the fourth day of incubation in the wells with filtered seawater b) Ulva lactuca spores in the wells with filtered seawater. Page 156

Figure 3: a) Ulva lactuca spores after immersion in 6 days in artificial seawater with visible germination tubes; b) Ulva spores after immersion in copper sulfate without the presence of germination tubes. Page 157

Figure 4: Percentage of spore germination of Ulva lactuca spores exposed respectively to artificial seawater (negative control), copper sulfate (positive control) and fraction E17163I. Page 161

Figure 5a and b: Antibacterial activity of fractions 16156 E3 and 16156 E4 against the growth of Cobetia marina and Pseudoalteromonas atlantica. B: 0.1% DMSO; C: 1:100

xx dilution of penicilin-streptomycin-neomycin stabilized solution (Sigma P4083). T-test results: * p-value <0.05, ** p-value<0.01; *** p-value <0.001. Page 165

Figure 6a and 6b: Antibacterial activity of fraction 17162 A against the growth of Vibrio harveyi (Figure 6a) and Halomonas aquamarina (Figure 6b). B: 0.1% DMSO; C: 1:100 dilution of penicilin-streptomycin-neomycin stabilized solution (Sigma P4083). T-test results: * p-value <0.05, ** p-value<0.01; *** p-value <0.001. Page 166

Figure 7a and 7b: Antibacterial activity of fraction 17166 D against the growth of Pseudoalteromonas atlantica (Figure 7a) and Roseobacter litoralis (Figure 7b). B: 0.1% DMSO; C: 1:100 dilution of penicilin-streptomycin-neomycin stabilized solution (Sigma P4083). T-test results: * p-value <0.05, ** p-value<0.01; *** p-value <0.001. Page 166

Figure 8a and 8b: Anti-diatom activity of fraction 17163 I against the growth of Cylindtrotheca sp. (Figure 8a) and Navicula sp. (Figure 8b). B: 0.1% DMSO. cycloheximide (3.55 µM) was used as the positive control (C+). T-test results: * p-value <0.05, ** p- value<0.01; *** p-value <0.001. Page 167

Figure 9. HPLC-PDA Chromatogram and PDA contour of E17163_I7_D. Page 169

APPENDIX

Supplementary figure 1: Rarefaction plot of observed OTUs for all the samples. Page 191

Supplementary figure 2: Absorption spectra (A) obtained by the analytic method with the absorbance as function of time. The first peak represents portoamide A, while the second peak is portoamide B. The PDA spectrum (B), for each absorbance spectrum, with absorbance in the wavelength of 276.0 nm. Page 192

Supplementary figure 3: 400MHZ 1H NMR spectrum of fraction E 16156_E3. Page 204

Supplementary figure 4: 400MHZ 1H NMR spectrum of fraction E 16156_E4. Page 204

Supplementary figure 5. 400MHZ 1H NMR spectrum of fraction E 17159_A. Page 205

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Supplementary figure 6. 400MHZ 1H NMR spectrum of fraction E 17162_A. Page 205

Supplementary figure 7. 400MHZ 1H NMR spectrum of fraction E 17166_D. Page 206

Supplementary figure 8: 400MHZ 1H NMR spectrum of fraction E 17163_I. Page 206

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Abbreviations list

AI-2 - Autoinducer-2

AF- Antifouling

AHLs- N-acylhomoserine lactones

ANOVA-Analysis of variance

ANOSIM- Analysis of similarities

APDL- Administração dos Portos do Douro, Leixões e Viana do Castelo, SA

ATCC- American Type Culture Collection

ATP-Adenosine Triphosphate

BBE- Blue Biotechnology and Evolution Team

BLAST- Basic Local Alignment Search Tool

BPR- Biocidal Products Regulation

CECT- Spanish Type Culture Collection

CEMUP- Centro de Materiais da Universidade do Porto

CIIMAR- Centro Interdisciplinar de Investigação Marinha e Ambiental

CNP- Cyanobacterial Natural Products Team

DMSO- Dimethyl sulfoxide

DNA-Deoxyribonucleic acid

EC50- Half maximal effective concentration

EPS- Extracellular polymers

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EU- European Union

FASP- Filter aided sample preparation

FCT- Fundação para a Ciência e Tecnologia

FDR- False discovery rate

DGGE- Denaturing gradient gel electrophoresis

FISH-Fluorescence in situ hybridization

HPLC- High Performance Liquid Chromatography

HPLC- DAD- High Performance Liquid Chromatography with Diode Array Detector

KEGG- Kyoto Encyclopedia of Genes and Genomes

KOs- PICRUSt-predicted KEGG orthologs

LC50 - Median lethal dose

LSD- Least Significant Difference

LCL- Lower confidence limit

LC-MS/MS- Liquid chromatography-mass spectrometry

LEGE- Laboratório de Ecotoxicologia, Genómica e Evolução

LEGE CC- Blue Biotechnology and Ecotoxicology Culture Collection

MBEC-Minimum biofilm eradication concentration

MIC- Minimal inhibitory concentration

MS/MS- Tandem mass spectrometry

NCBI- National Center for Biotechnology Information

NGS-Next-generation sequencing

NMWL- Nominal molecular weight limit

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NMR-Nuclear magnetic resonance

NRPS- Non-ribosomal peptide synthetase

OD- Optical density

OTU- Operational taxonomic unit

PCoA- Principal coordinate analysis

PICRUSt- Phylogenetic Investigation of Communities by Reconstruction of Unobserved States

PKS- Polyketide synthase

PVC- Polyvinyl chloride

PSU- Practical salinity unity

QIIME- Quantitative Insights Into Microbial Ecology

QS- Quorum-sensing

QSI- Quorum-Sensing inhibition

QTOF- Quadrupole time of flight

QQ- Quorum-quenching rRNA- Ribonucleic acid

TBT- Tributyltin

TBTO- Tributyltin oxide

TLC- Thin-layer chromatography

T-RFLP- Terminal restriction-fragment length polymorphism

UCL- Upper confidence limit

UNCTAD- United Nations Conference on Trade and Development

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UP- University of Porto

USD – US dollars

UV- Ultraviolet

VLC-Vacuum liquid chromatography

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“Nature knows no pause in progress and development and attaches her curse on all inaction.”

Johann Wolfgang von Goethe

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Chapter 1: General Introduction

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Chapter 1: General Introduction

1.1 Marine biofouling: an introduction

Marine biofouling is known to cause serious problems for marine industries and navies around the world and it is considered to be one of the most important issues facing marine technology (Dobretsov et al., 2013; Dang and Lovell, 2016; Carvalho, 2018). In the marine environment, all sorts of natural and artificial substrata are affected by fouling as they are colonized by marine micro and macroorganisms. Colonization of surfaces in the marine environment has been shown normally to occur by a four-step process: i) adsorption of dissolved organic molecules to a newly submerged surface; ii) colonization of the surface by marine bacteria; iii) colonization by microscopic eukaryotes; iv) protozoan colonization and settlement of invertebrate larvae and/or algal spores (Wahl, 1989; Maki and Mitchell, 2002;

Salta et al., 2013). (See Figure 1)

Figure 1: Evolution of marine fouling stages (adapted from Maki and Mitchell, 2002).

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Microbial biofilms as a rule precede surface colonization by invertebrates and macroalgae, and their establishment is considered a prerequisite for their colonization. In specific, marine invertebrate larvae and algae spores are believed to use biofilms as cues of substratum suitability for their settlement. (Burgess et al. 2003; Patel et al. 2003; Railkin, 2004; Qian et al., 2007). Therefore, biofilms dynamics mediates both microbial surface colonization and are believed to have different effects on modulating the colonization of macrofouling organisms (Mieszkin et al., 2013; Antunes et al., 2018). Marine biofilms are microbial aggregations, forming themselves at interfaces (Flemming and Wingender, 2010), and are mostly consisted of bacteria and diatoms. Physical properties of biofilms as well their biotic composition and chemical compound production constitute processes that shape biofouling communities (Qian et al., 2007, Salta et al., 2013). Overall, it is acknowledged that biofilms constitute highly dynamic systems, whose structure changes over time, and develops with the interaction with environmental parameters (Battin et al., 2007) (See Figure 2). In recent years, several studies have characterized the formation and the composition of marine biofilms, however, most past studies have used traditional community analysis methods that are unable to provide an in-depth picture of the marine fouling diversity in the natural setting

(Brian-Jaisson et al., 2014; Camps et al., 2014; Salta et al., 2013). Likewise, the literature is still sparse concerning the characterization of marine biofilm communities from different geographical areas, climates and/or ecological settings and it was not thoroughly determined if environmental parameters or seasonality are a deciding factor influencing the taxonomical diversity of marine biofilm communities (Dang and Lovell, 2016; Antunes et al., 2018).

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Figure 2: Marine biofilm development in the marine environment (adapted from Fuqua et al., 2001).

However, due to the great diversity of marine fouling organisms, it is not possible to accurately mimic complex natural fouling processes with in vitro experiments and the characterization of these communities remain mostly underexplored (Salta et al., 2013; Dang and Lovell, 2016). Current studies characterizing marine biofilms will need to keep on relying on continued advances in biomolecular techniques, and on the expanding number of sequenced marine prokaryote and eukaryote genomes (Leray and Knowlton, 2015; Zhang et al., 2016). Notably, in the past few years, the development of next- generation sequencing

(NGS) technologies and advances in computational biology opened up new research prospects in the description of microbial communities (Besemer et al., 2012; Lawes et al.,

2018).

An important process for marine biofilm formation is the quorum sensing (QS) phenomenon. QS is a form of bacterial population density-dependent cell–cell communication and gene regulation, which has been shown to contribute to the formation and development of biofilms in different settings. In particular, the change from planktonic to biofilm state is thought to be dependent on quorum sensing phenomena (Davies et al.,

1998; Parsek and Greenberg, 2005) (See Figure 2). In the biofilm stage, bacteria have improved access to nutrients, and a greater resistance towards hostile environments when compared to the planktonic stage (Hall-Stoodley et al., 2004). Since QS controls bacterial biofilm differentiation and maturation, inhibition of quorum sensing has become a strategy to control marine biofilm growth on surfaces immersed in seawater (Manefield et al., 2002;

Dobretsov et al., 2009). Specifically, bacterial QS can be suppressed by the inhibition of QS signal generation (Givskov et al., 1996), by degradation of QS signals (Dong et al., 2001), or by competition with/or suppression of QS receptors (Rasmussen and Givskov 2006). Many marine macroinvertebrates larvae or macroalgae spores are known to settle preferentially on bacterial bioﬁlms with distinct compositions and are believed to be dependent of chemical

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cues for their settlement and/or metamorphosis (Antunes et al., 2018). Therefore, QS inhibition may have an indirect effect on macrofouling settlement on surface by modulating the properties of biofilm (Dobretsov et al., 2007).

In the context of macrofouling, marine algae play a significant role in the marine fouling of a wide range of immersed artificial substrata and are one their most problematic organisms (Hellio, 2002; Mieszkin et al., 2013). Among the most problematic macrofouling algae, the filamentous brown algae of Ectocarpaceae family (phylum Heterokonta) is known to regularly colonize ships’ hulls and different artificial surfaces with coated with antifouling paints (Mineur et al. 2007; Lebret et al., 2009) and are also known to be one of the dominant algae on fish cages of aquaculture (De Nys and Guenther 2009). Another particularly problematic fouling alga is the intertidal green macroalga Ulva, previously called

Enteromorpha (Hayden et al., 2003). Tolerance to a wide range of salinities and water qualities contribute to the ecological success of this cosmopolitan genus and to its significance as the most widespread and troublesome ship fouling macroalgae throughout the world (Callow et al., 1997; Rao et al., 2007; Briand et al., 2009). Ulva is also known to have a great reproductive potential with vast numbers of motile zoospores (or zygotes from the fusion of gametes) being released from each thallus (Callow et al., 1997). Vectors implicated in macroalgal introductions include hull fouling, ballast water, shellfish transfers, and the aquarium trade (Kolar and Lodge 2001; Boudouresque and Verlaque, 2002; Lockwood et al.,

2007). Uncontrolled adhesion of fouling macroalgae such as Ulva sp. can cause serious problems by settling on ships’ hulls, aquaculture systems, and other marine infrastructure

(Hattori and Shizuri, 1996; Rao et al., 2007). Strategies for the development of nontoxic, environmentally benign controls of nuisance algae, such as those involved in marine fouling, require a greater understanding of the cellular mechanisms by which fouling organisms adhere to surfaces (Briand et al., 2009). Algae spore settlement results in a permanent algae attachment, and therefore most studies involve a mechanism to avoid the settlement of algae spores (Hellio, 2002; Rao et al., 2007). Therefore, unravelling the quantitative and qualitative aspects of algae spore settlement, as well to study the life history, overall biology and the

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ecology of the fouling algae (Briand et al., 2009) is important for anti-fouling purposes. A positive correlation was found for instance between zoospore settlement and bacterial biofilm density (Joint et al., 2000), and Ulva zoospores can also utilize bacterial quorum-sensing

(QS) signal molecules, specifically the N-acylhomoserine lactones (AHLs) as cues for the site selection of spores attachment (Joint et al., 2002; Tait et al., 2005; 2009).

1.2 Marine biofouling and marine biofilms: major problems and consequences

Biofouling remains a problem with serious economic and environmental impacts which makes the appearance of environmentally “clean” antifouling compounds essential

(Schultz et al., 2011). In the marine environment, all sorts of natural and artificial substrata are affected by fouling, both by marine micro and macro-organisms. The underwater hull of a ship is exposed not only to the corrosive seawater environment, but also to the constant accumulation of biofouling, and biofouling species includes any attaching organisms such as bacteria, diatoms, tubeworms, mussels, barnacles and algae (Callow and Callow, 2011;

Mieszkin et al., 2013). (Figure 3). Among the most important consequences, marine fouling results on increase of fuel consumption of ships and represents a major vehicle for the spread of invasive marine organisms (Schultz, 2007; Costas et al., 2013). Macrofouling organisms such as macroinvertebrates and macroalgae, in particular, are responsible for a significant percentage of marine fouling biomass and for the diminished hydrodynamic performance of a ship (Yebra et al., 2004). However, marine biofilm attachment to artificial surfaces is also a significant issue for a wide range of submerged engineered structures and is known to have a significant role in the corrosion of surfaces immersed in seawater (Dang et al., 2011;

Carvalho et al., 2018). Attachment of marine fouling species causes the deterioration of coatings favoring corrosion, especially in the case of settlement of invertebrates such as barnacles (Holm et al., 2004). These organisms use bioadhesives to provide a strong

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attachment to the hull, and the growth of their basal edges can penetrate and undermine the protective coatings, leading to premature corrosion (Hellio et al., 2000).

Figure 3: Main macrofouling organisms colonizing artificial surfaces (adapted Bressy and

Lejars, 2014).

Marine biofilms by themselves are known to negatively impact the hydrodynamics of ships leading to an increase in their fuel consumption (Townsin, 2003). Specifically, biofilm growth can alter the velocity structure in the boundary layer of surfaces, increase the wall shear stress, and cause an increase of the frictional drag of ships depending on the fouling type and coverage (Schultz et al., 2011). There is additional repercussion from increased energy consumption necessary to overcome increased frictional drag and heat and mass transfer limitations (Schultz et al., 2007). Accumulations of micro- and macro-organisms generate surface roughness and irregularities which increase the frictional resistance of a boat moving through water and consequently increase fuel consumption and emission of greenhouse gases (Hunsucker et al., 2018). The costs associated with fouling attached to ship

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hulls are mainly due to increased fuel consumption, resulting from higher frictional drag, although the costs for hull cleaning, coating and painting may also be considerable (Schultz,

2007). Over 90% of world trade is carried by the international shipping industry, and the

United Nations Conference on Trade and Development (UNCTAD) estimates that the operating cost of merchant ships alone contributes about US$380 billion in freight rates within the global economy, equivalent to about 5% of total world trade (Howell and

Behrends, 2006). The International Maritime Organization estimated that without new antifouling technologies, air emissions due to increased bunker fuel consumption by the world’s shipping fleet could increase by between 38% and 72% by 2020 (Towsin, 2003).

Antifouling paints and coatings save an estimated 60 billion USD annually and reduced of

384 million and 3.6 million tones, respectively, for carbon dioxide and sulfur dioxide per annum (Schultz et al., 2011). Apart from issues with fouling organisms themselves, surface- associated metabolic activities also exert harmful effects, including biocorrosion (Dang et al., 2011) and may even influence the buoyancy of plastic in seawater (Cao et al., 2011).

Settlement of fouling organisms on hull vessels is also a major issue concerning the introduction of invasive, nonindigenous (“alien”) species into non-native environments, and fouled vessels are the most common vectors of marine (Lejars et al., 2012). The introduced native species can compete with the native species and result on impacts to the local ecosystem, which in turn can demand new eradication and control measures (Hellio and

Yebra, 2009). The increasing prevalence of recreational yachting, coupled with the constantly rising number of alien species in marinas, is likely to enhance opportunities for the dispersal of these organisms (Gollasch, 2002). It should also be pointed out that coastal shipping has the potential to transport organisms and facilitate the spread of fouling species.

Overall, a successful solution to marine biofouling problems requires mechanisms that act effectively and simultaneously on different components and stages of fouling (Patel et al.,

2003; Chiu et al., 2008; Huggett et al., 2009).

1.3 Antifouling strategies

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Ancient Carthaginians and Phoenicians used pitch, and possibly copper sheathing, on the bottoms of ships to prevent biofouling, and later coated hulls with sulphur and arsenical compounds, while the Greeks and Romans both introduced lead sheathing (Durr and

Thomason, 2009). Copper cladding was then “re-invented” in the seventeenth and eighteenth centuries, but it became redundant after the introduction of steel vessels at the end of the nineteenth century, and also because of problems caused by galvanic corrosion (Yebra et al.,

2004). TBT (tributyltin) was invented around150 years ago, it was only used as an antifouling (AF) biocide in the early 1970s in combination with various copper compounds

(Sonak et al., 2009). As early as the late 1980s, TBT was referred to as ‘the most toxic compound ever brought into the marine environment, and several studies indicated that TBT- based compound had adverse effects on aquatic life and more specifically on nontargeted fouling organisms such as oysters, due to its high persistence and toxicity (Jelic-Mrcelic et al., 2006). Organotins like tributyltin (TBT) and tributyltin oxide (TBTO) were banned due to their toxicity, and alternative biocide-based antifouling paints such as Irgarol 1051 have been shown to accumulate in coastal waters and constitute threats to the marine environment

(Dafforn et al., 2011). The International Maritime Organization took into account the adverse effects of TBT on the marine environment, and an order was issued banning the use of this type of biocide in the manufacturing of AF paints from first January 2003, and the presence of these paints on ship surfaces from was banned since first January 2008 (Kotrikla,

2009).The restriction on the use of TBT led to a renewed use of copper-based paints and/or the use of new paints incorporating high levels of copper, however, copper (and other metals) also pose problems for the environment (Thomas and Brooks, 2013). Currently, other toxic metals and organic compounds including, chlorothalonil, dichlofluanid and diuron are used in marine AF coatings (Voulvoulis et al. 2000), however, once again these biocides have also been found to be toxic to the environment (Cresswell et al., 2006). Along with the increasing use of copper in AF paints, several alternatives to TBT have been developed, such as the use of synthetic and natural biocides (Yamada, 2007). The global costs of development of new

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biocides or new AF coatings incorporating biocides have increased, as they are now subject to restrictive legislations (e.g., the European Biocidal Products Directive, which requires several risk studies before registration and marketing authorization) (Salta et al., 2010). The

Biocidal Products Regulation (BPR) of the European Union (528/2012), requires the elucidation of AF mechanisms as a prerequisite for the registration and commercialization of any new antifoulant (Chen et al., 2017). It is thus necessary to understand the molecular targets of newly discovered AF compounds and this will elucidate the molecules or pathways involved in the settlement of biofouling organisms.

1.4 Antifouling agents of natural origin: current products and futures needs

With the ban of organotin-based marine AF products, the industry has a strong motivation and an incentive to discover new effective and environmentally friendly AF compounds from natural sources (Chen et al., 2017). Although over 700 of different compounds have been reported as having AF efficacy, only a few of these can be considered promising candidates (Qian et al., 2015). Part of the limitations of the newly discovered AF compounds include a too narrow inhibitory effect, a low LC50/EC50 ratio and activity based only in laboratory bioassays without field trials, as well the still limited information about the toxicity or ecotoxicity of the compounds (Wang et al., 2017). As an alternative there has been a search for alternative solutions to control marine biofouling, including the use marine- derived natural products or secondary metabolites (Rittschof et al., 2003; Fusetani 2004;

Qian et al., 2010). There are a number of advantages using marine microorganisms as sources of antifoulants. First of all, AF metabolites isolated from marine microorganisms are often chemically diverse and unique, which makes them promising sources for the exploitation of novel candidate antifoulants (Fusetani, 2004). Marine microorganisms can provide a number of benefits for anti-fouling industries like the possibility of having an

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unlimited supply of compounds through fermenter cultivation or scale-up bioreactors, and the possibility of genetic and chemical modification of the source organisms and compounds, respectively (Dobretsov et al., 2006). From an ecological point of view, these compounds are produced in situ and are believed to trigger meaningful responses in marine organisms (Wang et al., 2017). The production of these compounds by the microorganisms may have evolved in the interaction between the different organisms, and it is believed that the presence of neighboring species, as well environmental factors influence their production (Dahms et al.,

2006).

In recent years, marine organisms such as marine bacteria, fungi, cyanobacteria have been explored as a source of these products (Dobretsov et al., 2006; Chen and Qian, 2017;

Dahms and Dobretsov, 2017), and since the early 1980s, a great number of marine natural products have been assayed against organisms implied in the biofouling process (Omae,

2003; Fusetani, 2004). Many AF products have been extracted from various marine organisms and characterized (see the following reviews: Fusetani 2004, 2011; Qian et al.,

2010; Wang et al., 2017). Many marine-derived antifoulants have since been identified from diverse marine organisms, including marine bacteria, algae, soft corals, and sponges (Burgess et al., 2003; Dobretsov et al., 2006). Antifouling marine natural products have been classified according to their biosynthetic pathways into five main compounds: 1) terpenoids

(isoprenoids), 2) steroids and saponins, 3) fatty acid-related, 4) bromotyrosine derivatives and 5) heterocyclic compounds. Lactones, and furanones are some examples of compounds that have shown multiple antifouling activities (Maréchal and Hellio, 2009).

Marine microorganisms are a good source of anti-fouling compounds, and the limitation of the low quantity production of their compounds could be solved by large-scale culturing microorganisms and using genomic tools to identify the genes responsible for biosynthesis of the active metabolites (Penesyan et al., 2010; Wang et al., 2017). In addition, different microbial strains can produce different bioactive metabolites under different temperatures, pH and nutrient conditions, which further enriches the chemical scope of AF compounds (Dahms et al., 2006), and the production of natural products

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derived from microbes has to be through scale-up in bioreactors or by chemical synthesis

(Bhadury and Wright, 2004). Past studies however, focused on testing marine compounds against a limited number of taxonomic groupings, such as macrofoulers (e.g. only algae, barnacles and mussels) and were not tested or were not active against microorganisms in biofilms. (Scardino and de Nys 2011). Often many biocidal and fouling- release coatings are generally free from macrofouling, but they are still subject to colonization by biofilms (Cassé and Swain 2006; Molino et al., 2009; Dobretsov and Thomason 2011; Zargiel et al., 2011;

Briand et al., 2012). Formation of marine biofilms has been increasingly recognized as an important factor in influencing macrofouling (Salta et al., 2013; Antunes et al., 2018).

Nevertheless, existing knowledge of the interaction between biofilms and settling propagules and the molecular mechanism of anti-biofilm compounds is quite limited, and yet so far, and only several marine-derived antibiofilm compounds have been isolated and identified (Antunes et al., 2019). Another limitation of the introduction of antifouling products of marine origin is the relatively long time necessary to develop a new anti-fouling product. The industry prefers a fast and high-return product and there is still insufficient pressure for environmentally friendly antifouling products to force the coating industry to invest in research (Brady and Singer, 2000).

1.5 Cyanobacteria as a source of natural antifouling agents

Cyanobacteria, previously designated as “blue-green algae”, are a group of unicellular and multicellular photosynthetic prokaryotes that occur worldwide in freshwater, brackish and coastal marine ecosystems (Sivonen and Jones, 1999). Both marine planktonic and benthic cyanobacteria species are known, and the latter have been very successful in forming persistent biofilms in aquatic environments (Whitton and Potts, 2000).

Cyanobacteria can exhibit different morphologies and have the ability to perform a range of

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metabolic processes such as oxygenic and anoxygenic photosynthesis, fermentation and nitrogen fixation (Stahl, 1995) and show a remarkable physiological and ecological diversity

(Barberousse et al. 2006; Jelic- Mrcelic et al., 2006). These microorganisms require low light intensities and exhibit optimal growth rates at relatively high temperatures in comparison with eukaryotic microalgae (Paerl et al. 2011). They also have a higher ability to uptake nutrients, phosphorous and nitrogen than many other photosynthetic organisms (Paerl and

Paul, 2012), which means that they can compete with other phytoplankton under conditions of nutrient limitation (Briand et al., 2003). The presence of gas vacuoles provides buoyancy regulation in stratified waters, and this capacity dictates the success of cyanobacteria in the water column (Sunda et al., 2006). Cyanobacteria are among the oldest extant organisms on

Earth, dating in the fossil record to nearly 3.5 billion years ago (Mur et al., 1999). Over this long period, cyanobacteria have evolved to produce an impressive array of biologically active compounds which may have role as allelochemicals, or may have a role as defense against predators and grazers; however the chemical ecology of cyanobacteria and the functional role of the vast majority of these cyanobacterial compounds, remains largely unknown (Nunnery et al, 2010; Leão et al., 2012).

Due to this potential, cyanobacteria and their metabolites have been explored in the last years due to their major potential for the development of those compounds as algaecides, herbicides, insecticides, and also in other expanding areas such as neurosciences or as anti- proliferative or cancer toxic activities where they are being explored as rich sources for drug discovery (Tan et al., 2010). At the same time, there is an increasing knowledge of the biochemical and genetic functions of cyanobacterial production of secondary metabolites

(Pancrace et al., 2017). In the last years, marine cyanobacteria in particular have been considered as important biological sources of structurally novel compounds, particularly compounds containing nitrogen, such as peptides and lipopetides (Gerwick et al., 2001; Leão et al., 2012). Many of these cyanobacterial metabolites are the products of non-ribosomal peptide synthetase (NRPS) or polyketide synthase (PKS) pathways, and in the last years genome mining of biosynthetic gene clusters has become an essential methodology for

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studying NRPS/PKS biosynthetic gene clusters in cyanobacteria (Ehrenreich et al., 2005). At the same time there is an increase of technical knowledge regarding the purification and separation of active cyanobacterial biomolecules which has led to better approaches regarding the screening and selection of cyanobacterial compounds (Shimizu, 2003; Blunt et al., 2006). Due to the prolific production of secondary metabolites by cyanobacteria and due to their large chemical diversity, cyanobacteria have been regarded as an important source of antifouling compounds which inhibit biofouling through active mechanisms (Bhadury and

Wright, 2004; Yebra et al., 2004; Zhong, 2006; Niedermeyer, 2015). The majority of these compounds have been chemically identified as lipopeptides, polyketides, amides, alkaloids, indoles and fatty acids (reviewed by Dahms et al., 2006, Qian et al., 2009; Wang et al., 2017).

Cyanobacterial metabolites have potential to be used due to some of their characteristics including being potentially more effective and less toxic alternatives to combat marine biofouling and replace the biocide-based coatings currently in use (Almeida and

Vasconcelos, 2015; Wang et al., 2017). Among other advantages of using cyanobacteria as a source of natural products is the fact that cyanobacteria can easily be cultured and produce compounds much more rapidly and in large amounts when compared to invertebrates and algae, and also are able to be manipulated to achieve optimal production of bioactive substances, sustaining yields on an industrial scale (Bhadury and Wright, 2004; Dahms et al.,

2006). In spite of this well-recognized potential, cyanobacteria are still underexplored as a source of potential AF compounds (Chen et al., 2017).

1.6 Structure and main objectives

In the first chapter of this thesis includes a general introduction to the topic of marine fouling, its main impacts and hazards, and the main strategies used both historically and currently for controlling the growth of marine fouling organisms. In that chapter it is critically

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discussed how particularly metabolites from marine organisms can be used as an efficient strategy for antifouling purposes. Furthermore, it is emphasized the importance of characterizing marine biofilms in different settings, as well employing efficient assays against marine biofilms, as they constitute the initial, decisive step in the formation of marine fouling communities.

Past research has characterized marine biofilm communities growing on different artificial substrata, or on different geographical locations (Salta et al., 2013; Pollet et al.,

2018), and in the second chapter of this thesis those past field studies are reviewed and the importance of different chemical cues in the formation and development of marine biofilms is discussed. The chemical cues produced by marine microorganisms which are involved in the settlement of macrofouling organisms were also reviewed as they are important aspect for the succession of macrofouling communities. One of the conclusions put forward in this chapter is that few studies so far have comprehensively analyzed the evolution of early biofilms or have compared their communities in different seasons or in different geographic locations.

In the third chapter of this thesis we discuss the work which involves the sampling of marine biofilm samples growing on steel surfaces over a period of 30 days on two distinct seasons in a Northern Portuguese Port. During that study NGS (next-generation sequencing) technology was employed to characterize the biofilm samples taxonomically and the functional profile of those communities was also analyzed. temporal succession of these marine biofilms will likely have importance both for prevention of biocorrosion of steel surfaces and for future efficient antifouling strategies.

In the fourth chapter of this thesis, the screening assays that formed the basis for the discovery of anti-fouling compounds among LEGE CC cyanobacterial strains is discussed.

Our laboratory maintains a culture collection (LEGE Culture Collection) of 386 strains of cyanobacteria, including diverse marine, brackish water and freshwater strains, which have been collected predominantly in the Portuguese aquatic environments, mostly from the continent but also to a minor extent from the Madeira and Azores archipelagos (Ramos et al.,

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2018). Several of these strains have already shown interesting bioactive profiles using other marine organisms (bacteria, microalgae and metazoans) as targets, as well having anti- cancer, anti-viral, anti-microbial properties (Leão et al., 2013; Freitas et al., 2015, Ribeiro et al., 2017). Taking in consideration the antifouling potential of cyanobacterial compounds

(Dahms et al., 2006; Dobretsov et al., 2013) and that this diverse collection of cyanobacterial strains harbors a largely unknown chemical diversity, assays were conducted aiming the control of the growth of microfouling species, the first stage of marine biofouling. In order to find new antifouling cyanobacterial compounds, cyanobacterial extracts were also tested against a panel of several strains of the main microfouling groups (biofilm forming bacteria and diatoms), and assays were conducted both against their planktonic growth and the disruption of their biofilms. Bioassays were also performed to determine the quorum sensing inhibition potential of those compounds (which can disrupt biofilm formation). After this initial screening, the cyanobacterial strains with higher antifouling potential were grown in large scale in order for the conduct the isolation and purification of the bioactive anti-fouling compounds.

Taking as a background the bioactive potential of the cyanobacterial compounds portoamides (Leão et al, 2010; Ribeiro et al., 2018), in the chapter 5 of this thesis we discuss the work involved in determining the antifouling potential of those metabolites. In particular, we report the anti-microfouling assays conducted and the subsequent study of the portoamides mechanisms of action which analysed the potential of those cyanobacterial compounds as new environmentally friendly anti-fouling compounds.

The main aims of this PhD thesis are therefore both to develop natural antifouling/anti-biofilm natural strategies based on cyanobacterial compounds, and also to characterize the temporal variation of marine biofilm communities in situ in order to optimize efficient future strategies for their control.

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More specifically, the main scientific tasks of the present work were:

1. To review and critically discuss past studies involving marine biofilms sampled in different conditions as well the chemical cues associated with their formation, or which are involved in the interaction with marine macroorganisms.

2. Sample marine biofilm communities growing on steel surfaces on the Leixões Port for a period of 30 days on two distinct seasons and to characterize them taxonomically and functionally.

3. Compare those biofilm bacterial communities with the biofilm communities growing on steel surfaces covered with anti-corrosion paints and also to compare them with the local planktonic communities.

4. Conduct an extensive screening in order to identify cyanobacterial strains from the

LEGE culture collection that produce bioactive compounds against recognized and main microfouling species (bacteria and diatoms), against macroalgae species, and which are also capable of inhibiting the quorum sensing phenomenon. Test other bioactive cyanobacterial compounds isolated in our lab for their antifouling potential in the same assays.

5. Isolate and identify the cyanobacterial bioactive compounds responsible for the antimicrofouling activity and quorum sensing inhibition activity of the previous task.

Chapter 2

Review paper: “Marine biofilms: diversity of communities and of chemical cues” (published in Environmental Microbiology Reports)

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Marine biofilms: diversity of communities and of chemical cues

Jorge Antunes1,2, Pedro Leão1, Vitor Vasconcelos1,2,

1CIIMAR/CIMAR - Interdisciplinary Centre of Marine and Environmental

Research, University of Porto. Av. General Norton de Matos, s/n, 4450-208 Matosinhos,

Portugal.

2Department of Biology, Faculty of Sciences, University of Porto, Rua do Campo Alegre, P

4069-007 Porto, Portugal.

Received: 5 March 2018; Accepted: 14 September 2018

Summary

Surfaces immersed in seawater are rapidly colonized by various microorganisms, resulting in the formation of heterogenic marine biofilms. These communities are known to influence the settlement of algae spores and invertebrate larvae, triggering a succession of fouling events, with significant environmental and economic impacts. This review covers recent research regarding the differences in composition of biofilms isolated from different artificial surface types and the influence of environmental factors on their formation. One particular phenomenon - bacterial quorum sensing (QS) - allows bacteria to coordinate swarming, biofilm formation among other phenomena. Some other marine biofilm chemical cues are believed to modulate the settlement and the succession of macrofouling organisms, and they are also reviewed here. Lastly, since the formation of a marine biofilm is considered to be an initial, QS-dependent step in the development of marine fouling events, QS inhibition is discussed on its potential as a tool for anti-biofouling control in marine settings.

Keywords: Marine biofilms; marine fouling; distribution; environmental factors; chemical cues; biofilm formation

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2.1 Introduction

After a structure is submerged in the sea, marine microorganisms rapidly colonize it, forming complex biofilms (Davey and O’Toole, 2000; Huggett et al., 2009) with an important role in ecological cycles (Qian et al., 2007). Marine biofilms are organized in mixed communities of microorganisms dominated by bacteria and diatoms, surrounded by a matrix of extracellular polymeric substances (Allison, 2002), whose constitution depends on the dynamics and the interaction with environmental factors (Battin et al., 2007). In recent years, several studies have described the formation and the composition of marine biofilms

(Chung et al., 2010; Lawes et al., 2016; Briand et al., 2017). Most such studies, however, have used traditional community analysis methods that provide limited information regarding the actual diversity in the natural setting. Characterization of bacterial communities from marine biofilms using metagenomics analysis allows for a better assessment of the diversity present in the natural environment (Delong, 2005). The ability to characterize biofilm community composition in environmental samples has increased substantially through the use of high-throughput next generation gene sequencing (NGS) techniques allowing for ecologically-relevant field-based studies (Besemer et al., 2012). Still, the literature is sparse on what concerns the characterization of marine biofilms communities from different geographical areas, climates, ecological settings or surfaces.

Marine biofilms constitute preferential substrates for the colonization and settlement of larvae and spores of higher sessile organisms, and therefore influence later biofouling stages (Joint et al., 2000; Patel et al., 2003; Hadﬁeld, 2011). The microbial communities that initially attach to surfaces are important triggers of biofouling processes (Salta et al., 2013).

The formation of fouling communities involves a succession of events that has long been acknowledged since Claude Zobell´s pioneer work (Zobell, 1935), however the precise mechanisms and interactions involved in this initial surface adhesion are still unknown (Wahl

FCUP 21 Chapter 2 et al., 2012). Biofilms themselves have a significant role in the corrosion of surfaces immersed in seawater (Dang et al., 2011). Consequently, efficient anti-biofouling strategies need to consider both formation and characteristics of the biofilms. Biofouling remains a problem with serious economic and environmental impacts (Schultz et al., 2011), which makes essential the appearance of environmental “clean” antifouling compounds. Marine organisms are a potential rich source of compounds for antifouling, antibiofilm and antipathogenic purposes (Dahms et al., 2006; Fusetani, 2011; Almeida et al., 2015), which despite the increased interest in the last years, remain underexplored. In general, the formation and succession of marine bioﬁlms communities is a complex multistep process, and it is broadly acknowledged that a better comprehension of biofilm biology is necessary to mitigate the problem of biofouling in marine surfaces.

Cell-to-cell communication process between bacteria, generally known as quorum sensing (QS), controls several important features of bioﬁlm development among other phenomena (Waters and Bassler, 2005; Dobretsov et al., 2009; Antunes et al., 2010). In particular, the change from planktonic to biofilm state is thought to be dependent on quorum sensing phenomena (Rice et al., 1999). In the biofilm stage, bacteria have improved access to nutrients, and have a greater resistance towards hostile environments when compared to the planktonic stage (Hall-Stoodley et al., 2004). Furthermore, as QS controls bacterial biofilm differentiation and maturation (Xavier and Bassler, 2003), the disruption of QS may prevent microfouling. Many marine macroinvertebrates species are known to settle preferentially on bacterial bioﬁlms with distinct compositions and are believed to be dependent of chemical cues for their settlement or metamorphosis (Dobretsov et al., 2007).

QS inhibitors, particularly metabolites obtained from marine organisms can therefore potentially be used for the control of marine biofilm communities, and we decided in this review to focus on this antifouling strategy. Some recent reviews on marine biofilms are available (Qian et al., 2007; Hadfield, 2011; Mieszkin et al., 2013; Salta et al., 2013), focusing mostly on specific microbial groups and substrata. With this review we aim to evaluate the current research about marine biofilm diversity, the influence of environmental

FCUP 22 Chapter 2 factors on their formation and their chemical cues, as well as the potential implications for understanding and manipulating biofouling processes.

2.2 Diversity and distribution of marine biofilms communities

Marine biofilm communities are composed of heterogeneous marine species and their metabolic by-products (Salta et al., 2013). Typically, bacteria are the initial colonizers of surfaces, and are followed by diatoms. Other algae and invertebrates ensue (Cooksey and

Wigglesworth-Cooksey, 1995). Cyanobacteria, benthic dinoﬂagellates, fungi and protozoa are other species which can attach to surfaces by mucilage secretions (Molino and Wetherbee,

2008) and are often part of marine biofilms. The formation and maintenance of structured multicellular microbial communities is dependent on the production of EPS (extracellular polymeric substances) by the fouling microorganisms (Flemming and Wingender, 2010). To understand biofilm development, it is paramount to characterize their associated microbial diversity methods like rRNA gene sequencing, fluorescence in situ hybridization (FISH), denaturing gradient gel electrophoresis (DGGE) and terminal restriction-fragment length polymorphism (T-RFLP) have been used throughout the past decades. In the past few years, the development of Next-Generation Sequencing (NGS) technologies and advances in computational biology that deal with measuring the abundance of different microorganisms present, as well as their combined metabolic capabilities opened up new research avenues in microbial community analysis (Besemer et al., 2012). These NGS methods in particular can effectively be used to describe microbiomes in biofilms (Lawes et al., 2016).

Overall, the current body of work regarding bacterial communities of marine biofilms indicates that Proteobacteria, in particular the Alphaproteobacteria class dominate in these environments (Dang and Lovell, 2000; Huggett et al., 2006; Jones et al., 2007; Chung et al.,

2010). Roseobacter clade members (Alphaproteobacteria) seem to be the dominant group in

FCUP 23 Chapter 2 temperate coastal waters in the Atlantic and the Pacific Ocean (Dang et al., 2008), but also in the Mediterranean Sea (Elifantz et al., 2013; Briand et al., 2017), and Hong Kong (Chung et al., 2010; Lee et al., 2014). (See Figure 1). Other common taxa in most sampled marine biofilms are Sphingomonadaceae (Alphaproteobacteria), Alteromonadaceae

(Gammaproteobacteria) and Bacteroidetes (Dang and Lovell, 2000; Dang and Lovell, 2008;

Lee et al., 2014). Dominant taxa may differ with the geographical origin of the marine biofilms, as well with the type of colonized surface and with seasonality (See Figure 1 for a description of the characterized taxa in marine biofilm studies). However, there is still not enough data to determine any pattern of the distribution as a response to those parameters, and there is a huge diversity of taxa described for all those sampled biofilms which also depends in the sampling and sequencing approach and of the primers being used. Compared to bacterial biofilms in other settings, marine biofilms are more exposed to an influx of species that can influence the succession patterns and the stability of the communities (Røder et al., 2016). Concerning lifestyle, Pepe-Ranney and Hall, 2015 found a difference between biofilm and planktonic bacterial communities, the former showing a higher diversity of

Alphaproteobacteria. Planktonic species of Chlorophyta and Haptophyta are other widely- observed components of biofilm communities (Costas et al., 2013), as are Planctomycetes which adhere to different surfaces in aquatic environments such as marine sediments (Nocker et al., 2004), diatom cells, seaweeds, macrophytes, as well artificial surfaces (Sanli et al.,

2015).

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Figure 1: Global map with dominant taxa characterized in marine biofilm studies, their respective sampling sites, sampled surfaces and sampled periods.

Although the primary bioﬁlm is generally dominated by bacterial species, especially after immersion for a few days, a major, early accumulation of biomass is attributed to diatoms (Railkin, 2004). These organisms are known to adhere to different types of artificial surfaces (Cassé and Swain, 2006; Zargiel et al., 2011; Briand et al., 2012; Sweat and Johnson,

2013). Diatom settlement can be characteristic of an “infection type model”, in which the surface becomes colonized from a single diatom cell, growing exponentially until occupying a large area of the surface (Molino and Wetherbee, 2008). Heterotrophic bacterial species in bioﬁlms require a source of organic carbon for growth, and since the level of free organic material is relatively low in natural waters, the initial bacterial bioﬁlm growth is probably carbon-limited (Landoulsi et al., 2011). Diatoms are autotrophic and require only carbon dioxide and nutrients for growth which are usually not limiting in the marine environment.

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Due to such complementary life strategies, a mutualistic relationship between diatoms and bacteria is often established in biofilms (Landoulsi et al., 2011). Several authors demonstrated that biofilm diatoms growing on surfaces presented dissimilar profiles when compared to the planktonic diatom communities in that area (Patil and Anil, 2005; Cassé and

Swain 2006; Jones et al., 2007; Mitbavkar and Anil 2007; Lee et al., 2008; Shikuma and

Hadﬁeld, 2010). In particular, Patil et al., 2005a demonstrated that while centric diatoms dominate in the water column, pennate diatoms dominated in biofilms and belonged mostly to the genera Navicula, Amphora, Nitzschia, Pleurosigma and Thalassionema. The diversity of these photoautotrophs in biofouling situations is probably underreported, although they are acknowledged as typical fouling species on artificial surfaces (Molino and Wetherbee,

2008). For reviews about the involvement of diatoms in fouling and marine biofilms, see

Cooksey and Wigglesworth-Cooksey, 1995; Falciatore and Bowler, 2002; Molino and

Wetherbee 2008; Landoulsi et al., 2011.

2.3 Marine biofilm formation

Once a surface is submerged in seawater, it is quickly covered with a layer of adsorbed molecules, mostly proteins and glycoproteins, which form a conditioning film before the attachment of any microbial cells (Cooksey and Wigglesworth-Cooksey, 1995;

Flemming and Wigander 2010). After the formation of this conditioning film, bacteria adhere to the surfaces and their microcolonies grow, leading to the development of the biofilm microbial community (Flemming et al., 2016). The adhesion of bacterial cells modifies the surface physicochemical properties, and influences subsequent colonizers such as microalgae, cyanobacteria and diatoms which adhere to the surface after a few days (Dang and Lovell, 2016). Bacterial cells also produce extracellular polymeric substances (EPS), which results in the formation of a highly hydrated biofilm matrix (Flemming and

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Wingender, 2010). Other components like proteins, nucleic acids, lipids and biopolymers determine the biofilm physicochemical properties. (Flemming et al., 2016). Different bacterial taxa may respond differently to environmental factors, allowing for the efficient colonization of preferred surfaces. In addition, microorganisms can also answer to surface environmental signals and initiate surface adhesion by altering gene expression (Steinberg et al., 2002). The last step on the biofilm formation is the protozoan colonization, and the settlement of invertebrate larvae and/or algal spores (Wahl, 1989; Maki, 2002), which happens after some days and culminates in the formation of a mature marine biofilm (See

Figure 2 for a representation of the marine biofilm formation process).

Figure 2: Marine biofilm formation stages and respective influence of environmental parameters and chemical cues.

Evolutionarily, biofilms can provide shelter, nutritional advantages and create a protective structure, in which bacteria are protected from deleterious pathogens and other stressful conditions (Shnit-Orland and Kushmaro, 2009). Furthermore, it is known that biofilms allow the passage of nutrients, extracellular enzymes and metabolites necessary for microorganism growth (Sutherland, 2001). As predation constitutes a major mortality factor for planktonic bacteria (Sherr, 2002), biofilms can function as a protective niche against

FCUP 27 Chapter 2 predation (Matz et al., 2008), allowing microorganisms in biofilms to reach higher cellular densities. Moreover, Matz and co-workers (2008) observed a significantly higher presence of chemical defenses associated with the biofilm lifestyle. For example, diatoms are known to benefit from the biofilm´s properties, as marine biofilms constitute a source of protection from toxic compounds, UV radiation, refuge from protozoan grazing (Matz and Kjelleberg,

2005) and microbial pathogens (Shikuma and Hadfield, 2010). Additionally, diatoms associated with biofilms can similarly benefit directly from bacterial species present in biofilms, as bacteria can compensate from environmental nutrient deficiencies (Boyer and

Wisniewski-Dyé, 2009). Interactions within multi-species biofilms render microorganisms with an ability to grow under a particular environmental stress (Burmølle et al., 2006). As a whole, surfaces seem to represent a more favorable biotope for microorganism growth when compared to the water column (Railkin, 2004). The tendency of bacteria to colonize surfaces is advantageous from an ecological perspective because it preferentially targets specialized microorganisms to speciﬁc locations, encouraging symbiotic relationships (Dunne, 2002).

Biofilm formation can thus be regarded as an adaptive response to long-term coexistence, and its formation is believed to be related to the phylogenetic history of the species that constitute the biofilm (Madsen et al., 2016). The formation of the biofilm matrix is a dynamic process that depends on nutrient availability, the synthesis and secretion of extracellular material, social competition and grazing by other organisms (Flemming et al.,

2016). This multicellular process is triggered by bacterial motility and quorum sensing (QS), among other phenomena (Landini et al., 2010), and is tightly regulated (Nagar and Schwarz,

2015), constituting a cooperative enterprise in which constitutive strains and species work towards a common goal (Klein et al., 2014). In aquatic ecosystems, surface-associated microorganisms vastly outnumber organisms in suspension, and the formation of a biofilm allows a lifestyle that is entirely different from the planktonic state (Flemming and

Wingender, 2010). To enable changes in lifestyle, different genes are expressed in the planktonic and the sessile states, with biofilm-associated proteins not being present in the planktonic state (Hall-Stoodley et al., 2004). It is not known whether the biofilm matrix

FCUP 28 Chapter 2 confers an ecological advantage to all cells that constitute the biofilm, however a strong evolutionary benefit for polymer-producing cells in biofilms has been demonstrated

(Flemming and Wingender, 2010).

2.4 Marine biofilms and the influence of environmental factors

Understanding the factors that inﬂuence bacterial colonization in surfaces is fundamental for the prospective control of fouling on manmade structures (Qian et al., 2007).

Because marine biofilms are found in widely different environmental settings, growing both on natural and artificial surfaces, it is not feasible to generalize about this influence on their structure and on their physiological activities (Allison, 2002). Besides the substratum and the prevailing physical conditions on biofilms, the succession of marine bioﬁlm communities is an evolving process inﬂuenced by a wide variety of environmental conditions (Dobretsov et al., 2009; Qian et al., 2007). Physical parameters of surfaces like surface wettability (Dang and Lovell, 2000), physicochemical properties of substrata, such as surface hydrophobicity, critical surface tension (Jones et al., 2007), and surface molecular topography (Lee et al.,

2008) are known to influence biofilm growth and their characteristics. Equally, other environmental parameters such as temperature (Lau et al., 2005), radiation (Roeselers et al.,

2007, 2008; Behrendt et al., 2012), pH (Baragi and Anil, 2016; Brown et al., 2018) dissolved oxygen (Lagos et al., 2016), CO2 (Nayar et al., 2005; Brown et al., 2018), nutrient availability

(Chiu et al., 2008; Briand et al., 2012) can influence the rate of bacterial colonization as well the composition and ecophysiology of biofilm species (Dang and Lovell, 2000; Hall-Stoodley et al., 2004). (See figure 2 for a representation of the influence of these parameters in biofilm formation). Biomass and polymer accumulation in biofilms are dominated by physical adsorption processes, which in turn are influenced by environmental processes like nutrient

FCUP 29 Chapter 2 recycling (Moss et al., 2006). Since environmental parameters vary seasonally or spatially, and may alter the planktonic communities or bacterial attachment, these can in turn have an impact in the establishment of a biofilm microbial community (Donlan, 2002).

Environmental nutrient variation may trigger the formation of marine biofilms during community assembly, as well as modulate the composition of the communities (McDougald et al., 2011). A recent study demonstrated that the decrease of mineral nutrients available to biofilm photoautotrophs, resulted in a decrease in the diversity of microorganisms responsible for biofilm formation, affecting both the stability and function of marine biofilms

(Lawes et al., 2016).

The specific settings and surfaces where biofilms are formed are differently modulated by environmental factors. Likewise, the lifestyle of the colonizing microorganisms can influence biofilm diversity and the responses to environmental factors by the microbial community (Pepe-Ranney and Hall, 2015). Biofilms formed on ship´s hulls in particular, are normally more complex and dynamic than the ones growing in other types of artificial surfaces and depend on factors like the dynamic conditions of the surfaces

(Zargiel et al., 2014). Microfouling communities growing on these surfaces are subject to selective physical pressures from the environment, and are more protected from factors like insulation, oxidative stress, predation and shearing when compared to biofilm communities growing on other surfaces (Matz et al., 2008). These environmental factors alter the physiochemical and biological dynamics of the water column, and also influence the community structure of the biofilm (Patil and Anil, 2015 a). Higher bacterial densities were found with the maximum age of the biofilm (Wieczorek et al., 1995); it has been proposed that, as the biofilm matures, its composition tends to become progressively more similar to the local microbial community (Pepe-Ranney and Hall, 2015). However, biofilm and planktonic communities remain distinct even from early stages of biofilm formation (Ward et al., 2003).

FCUP 30 Chapter 2

2.5 Marine biofilms and chemical cues

Biofilm formation is cooperatively regulated by intercellular communication, like

QS, which regulates gene expression when cell numbers reach a critical threshold, resulting on phenotypic variations (McDougald et al., 2011). These phenotypic adaptations are believed to be crucial for biofilm development in changing environmental conditions (Chen et al., 2013). Because cells in biofilms are sessile, biofilm formation and motility are considered opposing mechanisms (Hölscher et al., 2015), and the relationship between swarming and biofilm formation is likely to be dependent on the regulation of QS (Waters and Bassler, 2005; Alagely et al., 2011; Guttenplan and Kearns, 2013). In some conditions,

QS may be necessary for EPS synthesis and biofilm formation, while in others it can induce motility and biofilm dispersal (Hoang et al., 2008) (See figure 2). On the other hand, the production of QS inhibitors or antagonists by marine algae and invertebrates associated with biofilms is an important ecological trait that has likely co-evolved with QS (Lau et al., 2005).

Quorum sensing molecules have been detected in several Gram-negative bacteria of marine origin (Mohamed et al., 2008), but are seldom found in marine Gram-positive organisms. For example, Cuadrado Silva et al., 2013 isolated several marine strains with quorum sensing systems mediated by different acyl homoserine lactones (AHLs, a class of common QS autoinducers): N-butanoyl homoserine lactone, N-hexanoyl homoserine lactone, or both from fouled surfaces. Furthermore, a comprehensive study demonstrated the presence of several autoinducer synthases (responsible for the biosynthesis of different QS autoinducers) in three different oceans, encompassing a large phylogenetic diversity and underpinning its global prevalence (Doberva et al., 2015). The production of AHLs has been described in several marine Gram-negative bacteria (Parsek et al., 1999; Daniels et al., 2004), and the widespread occurrence of QS molecules in marine Alphaproteobacteria has been shown for both free- living strains and those associated with eukaryotic algae (Wagner-Döbler et al., 2005).

FCUP 31 Chapter 2

Nevertheless, AHL production has been so far reported chiefly among members of the

Vibrionaceae family of Gram-negative marine bacteria (Purohit et al., 2013). Huang et al.

2007 showed differences in patterns of AHL biosynthesis either due to temporal regulation of AHL production within biofilms or due to different bacterial species dominating at different stages of biofilm development.

Although most of the known QS signals are species specific, QS signals involved in inter-species signaling have also been discovered. In specific, molecules of the autoinducer-

2 (AI-2) family are believed to regulate inter-species and intra-species communication

(Schauder et al., 2001; Bassler and Losick, 2006). AI-2 mediated signaling was first identified in the early 1990’s in Vibrio harveyi (Bassler et al., 1994), and unlike AHL’s, AI-

2 molecules are present in both gram-positive and gram-negative bacteria (Bodor et al.,

2008). The luxS gene is in involved in AI-2 biosynthesis and it was first discovered in Vibrio harveyi and Escherichia coli (Surette et al., 1999). Afterwards, it was found also in over 70 bacterial species and it was demonstrated that inactivation of luxS gene results in the elimination of AI-2 production (Whiteley et al., 2017).AI-2 system is correlated with pathogenicity in a variety of organisms, and is known to regulate a host of bacterial processes including virulence and biofilm formation (Bassler and Losick 2006). Therefore, the identification of molecules capable of interfering with AI-2 signaling has attracted interest for anti-virulence and anti-biofilm purposes. The specific role of AI-2 signaling in marine microbial communities remains to be demonstrated. It has been hypothesized that it may be involved in the regulation of interspecies interactions in complex microbial communities

(Hmelo, 2017). However, not all bacteria which produce AI-2 molecules encode a known cognate sensor or respond to AI-2 in a way consistent with a signaling system (Vendeville et al., 2005). Among marine bacteria, AI-2 is only known to function as a signal in Vibrio species (Milton, 2006), and AI-2 producing Vibrio species were found to be predominant and have an important role in microbial communities present in coral reefs or sponges (Tait et al.,

2010, Zimmer et al., 2014). There is not much data regarding the presence of AI-2 signals in the ocean beyond coral communities, despite the fact that luxS gene sequences seem to be

FCUP 32 Chapter 2 prevalent in the Global Ocean Sampling database (Doberva et al., 2015). Therefore, it is difficult so far to fully determine the importance of AI-2 in quorum sensing of marine bacteria, and its role marine biofilm formation.

The prevalence of QS-mediating compounds in marine environments is probably underestimated and has been reported only for a few taxonomic groups (McLean et al., 1997;

Mohamed et al., 2008; Leão et al., 2012). Non-QS compounds like tetrabromopyrrole (Figure

3-1) (Sneed et al., 2014), 6,9-heptadecadiene (Figure 3-2) (Hung et al., 2009), 12- octadecenoic acid(Figure 3-3) (Hung et al., 2009), amino acids (Jian and Qian, 2005), exopolysaccharides from marine biofilms (Khandeparker et al., 2002, 2006), or histamine

(Figure 3-4) (Swanson et al., 2006),are also known to induce the metamorphosis, or the settlement of larvae of different marine organisms, particularly species associated with macrofouling (See Table 1 for a description of non-QS chemical cues from biofilms and their role with other marine organisms and see figure 3 for their chemical formula).Complex community assembly on marine environments is likely led by QS signaling, with enrichment of specific members with QS signaling, and with the exclusion of member with anti-QS signaling traits (Tait et al., 2005; Tan et al., 2015b). Additionally, marine eukaryotes have been reported to compete with other bacterial signals, inhibiting the expression of bacterial signaling phenotypes (Rice et al., 1999). It is likely that many aquatic multicellular organisms evolved different mechanisms to interfere with bacterial QS (reviewed by Dobretsov et al.,

2009; Goecke et al., 2010). In particular, bacterial bioﬁlms have been shown to enhance or inhibit settlement of larvae from many invertebrate phyla, and therefore have been studied regarding the settlements cues for a broad range of marine invertebrate taxa, both in laboratory, and in field studies (Wieczorek et al., 1995; Olivier et al., 2000; Qian et al., 2013;

Bao et al., 2007; Zardus et al., 2008).

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Table 1. Non-QS chemical cues from marine biofilms and their relation to affected marine organisms

Affected Target species Chemical cue Producing Reference mechanism species Caribbean corals: Porites astreoides, Biofilms of Steinberg et Induction of Orbicella marine al., 2002; settlement and franksi, Tetrabromopyrrole bacterial Tebben and metamorphosis Acropora (Figure 3-1) Pseudoalterom Tapiola, palmate and onas sp. PS5 2011; Sneed Acropora et al., 2014 millepora

6,9-heptadecadiene (Figure 3-2), 12- octadecenoic acid (Figure 3-3; Hung et al., dissolved amino 2009; acids and Multispecies Beckmann et Hydroides extracellular marine al., 1999; Jin Induction of larval elegans; polymers; biofilms; and Qian, settlement Holopneustes Histamine Macrococcus 2005; Lam et purpurascens; (Figure 3-4); sp. AMGM1 al., 2005; Perna Unidentified and Bacillus Swanson et canaliculus; glycolipids from sp. AMGB1 al., 2006; Amphibalanus from Bacillus sp. from biofilm Ganesan et amphitrite and non-polar exudates al., 2012; protein; LCA- Jouuchi et al., binding sugar 2007 chain(s) compound

Pseudomonas aeruginosa Szewzyk et Balanus biofilms al., 1991; Induction of larval amphitrite and Biofilm isolated from Khandeparker metamorphosis Ciona exopolysaccharides the shell of et al., 2002, intestinalis Balanus 2006; Amphitrite, B. Patil and pumilus and C. Anil, 2005b freundii and marine diatom biofilm

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Macroinvertebrate larval attachment is thought to act as a response to chemical cues that are present in marine biofilms as well as in the surrounding water (Bao et al., 2007;

Ganesan et al., 2010). Despite the importance of these chemical cues from microbial biofilms, few have been identified, mostly AHL’s associated with QS phenomena with different roles in the settlement of macroinvertebrates or macroalgae (Steinberg et al., 2002; Jin and Qian,

2005; Hung et al., 2009). (See Table 2 for a summary of the reported QS chemical cues from marine biofilms and their action with other marine organisms and see figure 4 for their respective chemical formula).

Figure 3: Non-quorum sensing chemical cues identified in marine biofilms. Chemical formulas names and sub-numbers: (1) tetrabromopyrrole; (2) 6,9-heptadecadiene; (3) 12-octadecenoic acid; (4) histamine. See Table 1 for the interaction of each of these non-QS chemical cues with macrofouling species and their affected mechanisms.

Marine bacterial biofilms are also known to enhance the settlement of spores of the green alga Ulva sp., with settlement being very dependent on the nature of the biofilms

(Mieszkin et al., 2012, 2013). In this particular type of signaling, Ulva zoospores are attracted by marine bacterial biofilms through the release of N-Acyl-L-homoserine lactones (AHLs)

(Joint et al., 2007), which facilitate the settlement of zoospores, namely N-hexanoyl-L- homoserine lactone (Figure 4-5), and N-(3-oxodecanoyl) -L-homoserine lactone (Figure 4-

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6) (Joint et al., 2000, 2007; Williams, 2007). Liberation of carpospores from the algae

Gracilaria dura was also found to be associated to the prodution of quorum sensing signals by bacteria, specifically N-butanoyl-L-homoserine lactone (Figure 4-7) (Singh et al.,

2015).According to Tait et al., 2005, these compounds are required for the attraction of Ulva zoospores to biofilms, which are the preferred sites for their attachment. Ulva can sense different AHL molecules, which are thought to be used for the selection of surface sites for zoospore attachment through chemokinesis (Tait et al., 2005; Tait et al., 2009). These molecules are N-dodecanoyl-L-homoserine lactone (Figure 4-8), N-(3-oxobutanoyl)-L- homoserine lactone (Figure 4-9) and N-(3-oxododecanoyl)-L-homoserine lactone (Figure 4-

10) (Tait et al., 2005; Tait et al., 2009).

Figure 4: Quorum-sensing chemical cues identified in marine biofilms. Chemical formulas names and sub-numbers: (5) N-hexanoyl-L-homoserine lactone; (6) N-(3-oxodecanoyl) -L-homoserine lactone; (7) N-butanoyl-L-homoserine lactone; (8) N-dodecanoyl-L-homoserine lactone; (9) N-(3- oxobutanoyl)-L-homoserine lactone; (10) N-(3-oxododecanoyl)-L-homoserine lactone;(11) N- octanoyl-L-homoserine lactone; (12) N-(3-oxodecanoyl)-L-homoserine lactone. See Table 2 for the interaction of each of these QS chemical cues with macrofouling species and their affected mechanisms.

Interestingly, QS molecules from Ulva-associated endophytic bacteria interfere with the settlement, growth and maturation of the macroalga (Park et al., 2011; Twigg et al., 2014).

(See Table 2). This sort of signal transduction between eukaryotes and bacteria has been

FCUP 36 Chapter 2 denominated as “interkingdom signaling” (Williams, 2007; Lowery et al., 2008). (See Table

2 for examples of these interactions). The inﬂuence of bioﬁlm age on larval and mussel settlement is generally reported to be quantitative, increasing with the age of the biofilm

(Harder et al., 2002; Bao et al., 2007; Toupoint et al., 2012; Wang et al., 2012; Yang et al.,

2014; Li et al., 2010; Li et al., 2016; Freckelton et al., 2017); however, in some cases, induction of larval settlement is favored by younger, less dense biofilms (Dobretsov et al.,

2007; Yang et al., 2016). The role of marine biofilm bacteria on the attachment of the barnacle Balanus amphitrite has been well documented on bioassay studies, and its settlement varies with biofilm substrata (Wieczorek et al., 1995; Kamino, 2013), as well the production of QS molecules like N-(3-oxodecanoyl) -L-homoserine lactone (Figure 4-6), N-

(3-oxodecanoyl) N-dodecanoyl-L-homoserine lactone (Figure 4-8) and N-octanoyl-

Lhomoserine lactone (Figure 4-11). (Lee et al., 2014) Increasing bacterial load of different genera in biofilms seems to be responsible for the primary colonization of submerged surfaces by the tubeworm Hydroides elegans (Olivier et al., 2000; Harder et al., 2002; Huang and Hadfield, 2003; Shikuma and Hadfield, 2005, Lau et al., 2002). Contact between larvae and biofilm appears to be necessary for this interaction to occur (Hadfield et al., 2014), and there is also the mediation of AHLs’s such N-hexanoyl-L-homoserine lactone (Figure 4-5),

N-dodecanoyl-L-homoserine lactone (Figure 4-8) and N-(3-oxodecanoyl)-L-homoserine lactone (Figure 4-12) (Huang et al., 2007).

Table 2. Quorum-sensing molecules from marine biofilms and their relation to affected marine organisms

Affected Producing Target species Chemical cue Reference mechanism species

N-hexanoyl-L- Vibrio Wheeler et Stimulation of homoserine lactone anguillarum al., 2006; zoospore attachment Ulva sp. biofilms

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(Figure 4-5), N-(3- Joint et al., oxodecanoyl) -L- 2007 homoserine lactone (Figure 4-6).

N-butanoyl-L- Gram-negative homoserine lactone epi- and Induction of Ulva sp. and (Figure 4-7) endophytic Singh et al., carpospore Gracilaria sp. , N-hexanoyl-L- seaweed- 2015 liberation homoserine lactone; associated (Figure 4-5) bacteria

N-dodecanoyl-L- homoserine lactone (Figure 4-8); N-(3-oxobutanoyl) - L-homoserine Sulﬁtobacter Increase of zoospore Ulva sp. lactone (Figure 4-9); spp. and Tait et al., germination N-(3-oxodecanoyl) Shewanella sp. 2009; Twigg L-homoserine from a et al., 2014 lactone (Figure 4-6); polymicrobial N-(3- biofilm oxododecanoyl) -L- homoserine lactone (Figure 4-10).

C8-HSL, N- octanoyl-L- homoserine lactone Single species Increase of cyprid (Figure 4-11); bioﬁlms of settlement Balanus N-dodecanoyl-L- Vibrio Tait et al., improvises homoserine lactone anguillarum, 2013 (Figure 4-8) and Aeromonas 3-oxo- C10-HSL, N- hydrophila and (3-oxodecanoyl) L- Sulﬁtobacter homoserine lactone sp. strain BR1 (Figure 4-6)

N-hexanoyl-L- homoserine lactone (Figure 4-5) N- Induction of larval Hydroides dodecanoyl-L- Subtidal Huang et al., settlement elegans homoserine lactone biofilm 2007 (Figure 4-8); and 3- oxo-C8-HLS, N-(3-

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oxodecanoyl) -L- homoserine lactone (Figure 4-12).

Marine bacteria, as well eukaryotes, can produce enzymes like AHL acylases and

AHL lactonases that inactivate AHL signals in a process known as “quorum quenching” (QQ)

(Dong et al., 2001). Signal concentration in the environment can be regulated via specific enzymatic degradation activity, or quorum quenching by members of the community

(Daniels et al., 2004). One possible mechanism for QS behavior regulation, is through the control of signal concentrations (Antunes et al., 2010), and this can be regulated via speciﬁc enzymatic signal degradation activity or by quorum quenching activity (Dong et al., 2007).

Complex community assembly on marine environments is probably led by QS signaling, with enrichment of specific members with QS signaling and with the exclusion of member with anti-QS signaling traits (Tait et al., 2005). (See Figure 5 for an example of QS and QQ processes in marine biofilms). For example, in some marine Gram-negative biofilm bacteria, furanones are strong inhibitors of quorum sensing regulated behavior, and also affect the expression of virulence in genes, bacteria motility and formation of biofilms (Kjelleberg et al., 1997; Dobretsov et al., 2007; Dobretsov et al., 2009). The presence of these compounds results on a shift in the composition of microbial communities from being dominated by

Gram-negative bacteria to be dominated by Gram-positive species (Kjelleberg and Molino,

2002). Production of QS inhibitors can have a role in securing a foothold in a surface environment by inhibiting the attachment of marine bacteria, and consequently of larvae, algae and barnacles (Goecke et al., 2010; Wahl et al., 2010). Described quorum sensing inhibitors include furanones and their analogs (Zang et al., 2009; Liu et al., 2010), bismuth porphyrin complexes (Galkin et al., 2015) or glycosylated flavonoids (Brango-Vanegas et al., 2015). A study demonstrated that Kojic acid inhibited quorum sensing activity and formation of microbial communities on glass slides, decreasing both the densities of bacteria and diatoms (Dobretsov et al., 2011).

FCUP 39 Chapter 2

Recently the production of QS inhibitory metabolites has also been reported in different marine bacteria, mostly Alpha and Gammaproteobacteria (Mohamed et al., 2008;

Kanagasabhapathy et al., 2009; Teasdale et al., 2009; Dobretsov et al., 2011). QQ activity was already evaluated in cultivable bacteria obtained from ocean and estuarine seawaters and was found to be more common in specific habitats in the marine environment, like isolates from subtidal biofilms and sponges (Romero et al., 2011). Quorum sensing and quorum quenching activities are associated with phylogenetically diverse marine microbial groups, and there are recognized AHL producers such as Acidovorax, Pantoea, Rhizobium,

Rhodobacter, Sphingomonas and Streptomonas and quenchers such as Ochrobacterium,

Variovorax, Bacillus and Rhodoccus (Tan et al., 2015). A recent metagenomic study revealed that both QS and QQ gene sequences were common among free-living bacteria, and their prevalence shifted according to the depth of the sampled bacteria (Muras et al., 2018).

Although QQ activity has been demonstrated in diverse microbial species, the ecological role of QQ on QS signaling remains largely unknown but may be related to the selection of stable

FCUP 40 Chapter 2 and functional microbial assemblages (Tan et al., 2015b). In these communities, quorum quenching is hypothesized to perform a role in the regulation of quorum sensing signaling and community assembly (Tan et al., 2015). In the last years, there has been an increasing number of reports of marine bacteria presenting AHL degrading mechanisms like the above- mentioned AHL lactonases or acylases under laboratory conditions (Decho et al., 2009,

Romero et al., 2012). The significant number of sequences coding for putative acylases and lactonases in metagenome collections (Romero et al., 2012) suggests prevalent, uncharacterized quorum quenching activity in marine bacteria. For example, a marine metagenomic screening (Yaniv et al., 2017) found that anti-biofilm and anti-quorum sensing activities are common among uncultured marine planktonic bacteria, proving that functional screenings can provide a source of quorum sensing inhibitor compounds.

2.6 Marine biofilms and their relation to marine fouling

Biofouling is known to cause serious problems for marine and naval industries as well as navies around the world and is considered to be one of the most important issues facing marine technology today (Abarzua et al., 1999; Yebra et al., 2004; Salta et al., 2013). In the marine environment, all sorts of natural and artificial substrata are affected by both micro- and macrofouling. Although organisms such as mussels and barnacles are responsible for a significant part of macrofouling, and for the diminished hydrodynamic performance of a ship, as mentioned earlier microorganisms have an important role in surface colonization.

(Steinberg et al., 2002; Qian et al., 2007, Salta et al., 2013). Therefore, understanding the development of microfouling communities is an important aspect in the selection and management of anti-fouling (AF) coatings for marine operations. Marine biofilms are known to alter the velocity structure in the boundary layer of surfaces, increase the wall shear stress, and cause an increase of the frictional drag of ships depending on the fouling type and

FCUP 41 Chapter 2 coverage (Schultz et al., 2011).In order to determine the effect of biofilms in frictional drag, an early study was conducted with rotating disk devices which simulated fouling conditions

(Loeb et al., 1984). The results showed that disks covered with biofilms had up to a 5% higher resistance at a velocity of 40 knots when compared the disks with fouling-released coatings.

Another study by Haslbeck and Bohlander (1992) was conducted in a Knox class frigate to more correctly determine the effect of biofilms on the frictional drag of ships. In that study, the removal of biofilms from a ship stationed for 22 months in Pearl Harbor resulted in an

18% decrease of shaft horsepower to propel that ship. Additionally, a later study found that the skin friction coefficient of flat plates colonized by microfouling communities had an average increase of between 33 to 186 % relatively to surface covered by antifouling paints

(Schultz et al., 1999). More recently, Holm et al., (2004) employed rotating disk machines similar to Loeb et al., 1984 to determine frictional drag on ships and compared the drag penalties between surfaces with accumulated biofilms and surfaces with different anti-fouling release coatings. Hydrodynamic drag penalties resulting from biofilm growth were between

9% to 29% higher when compared to anti-fouling release coating surfaces.

Overall, marine biofilms are recognized to be a significant issue for a wide range of submerged engineered structures. Biofouling remains today a problem with serious economic and environmental impacts (Yebra et al., 2004), which makes the appearance of environmental “clean” antifouling compounds essential. Biofouling is responsible for friction on boats, inducing excessive fuel consumption (Townsin, 2003) increased maintenance costs, and generates considerable monetary losses annually (Schultz, 2007). There is additional repercussion from increased energy consumption necessary to overcome increased frictional drag and heat and mass transfer limitations (Schultz et al., 2011). The costs associated with fouling attached to ship hulls are mainly due to increased fuel consumption, resulting from higher frictional drag, although the costs for hull cleaning, coating and painting may also be considerable. Antifouling paints and coatings save an estimated 60 billion USD annually

(Schultz et al., 2011). Apart from issues with fouling organisms themselves, surface- associated metabolic activities also exert harmful effects, including biocorrosion (Dang et al.,

FCUP 42 Chapter 2

2011) and may even influence the buoyancy of plastic in seawater (Cao et al., 2011). The environmental impact of biofouling control strategies is also a cause of concern. Certain eﬃcient but environmentally harmful biocides, commonly used for controlling biofouling have been banned, notably tributyltin (TBT), as well as some herbicides used as “booster biocides” (Yebra et al., 2004). Likewise, alternative biocide-based antifouling paints such as

Irgarol 1051 have also shown to accumulate in coastal waters and constitute threats to the marine environment (Dafforn et al., 2011). Therefore, during the last years, much attention has been given to the use of environmentally benign antifoulants, particularly compounds from marine organisms (Fusetani, 2011). Among them, surfaces incorporating natural substances isolated from living organisms such as antimicrobial peptides, bacteriolytic enzymes or secondary metabolites have been proposed (Hellio et al. 2001; Qian et al., 2007).

Recent reviews into marine fouling control using textured and biomimetic surfaces have identified the numerous challenges of developing antifouling (AF) technologies (Salta et al.,

2010; Scardino and de Nys, 2011). Importantly, it was highlighted that these technologies usually target specific taxonomic groups. For instance, altering the physico-chemical properties of the surfaces (like surface roughness and fluid hydrodynamics) may inhibit macrofoulers but may not be active against biofilms which is necessary for an effective antifouling strategy (Magin et al., 2010). On the other hand, the use of “cleaner” antifouling compounds derived from marine organisms in particular, have the limitation of having to be produced in significant amounts which carries technical and economic challenges, and information regarding “antifoulants” compounds derived from the marine organisms is still limited (Fusetani, 2004, 2011). See also Magin et al., 2010 and Dafforn et al., 2011 for a review of different antifouling strategies used in marine settings. In recent years marine organisms such as marine bacteria, fungi, cyanobacteria have been explored as a source of these products (Dobretsov et al., 2006). There has been a search for quorum sensing inhibiting metabolites (Dobretsov et al., 2011) which can inhibit quorum sensing and biofilm formation, and as a consequence have an impact in the succession of marine fouling communities (Dobretsov et al., 2009; Salta et al., 2013). The environmental fate and effects

FCUP 43 Chapter 2 of these compounds should also be considered during biofouling management and control.

Natural products can be successfully used for antifouling applications by incorporating in a suitable paint. However, preparation of an antifouling coat using microbial products is a major challenge as these compounds will breakdown rapidly in the environment. So far, there have been few attempts to incorporate natural products into paint and successful mid- term to long-term field trials are still scarce (Abarzua et al., 1999).

2.7 Quorum-sensing inhibition as a strategy to prevent marine biofilm growth

Different strategies have been studied to control marine biofilms, namely the use of specific enzymes like proteases and oxidases, the use of microcapsules with biocides and the application of quorum sensing inhibitors, particularly those derived from marine organisms

(Dobretsov et al., 2006, 2013). Since QS controls bacterial biofilm differentiation and maturation, inhibition of quorum sensing has the requirements to become a privileged strategy to control marine biofilm growth and corrosion of surfaces immersed in seawater.

Bacterial QS can be suppressed by the inhibition of QS signal generation (Givskov et al.,

1997), by degradation of QS signals (Dong et al., 2001), and by competition with/or suppression of QS receptors (Rasmussen and Givskov 2006). Marine organisms not only respond to bacterial QS signals, but also interfere and block them, presumably to control the growth of alien bacteria, which would lead to biofilm formation (Skindersoe et al., 2008). In addition to QS inhibition molecules, enzymatic degradation is a different strategy to inhibit bacterial signaling molecules. In specific, AHL-lactonases and decarboxylases hydrolyze lactone rings, while AHL-acylases and deaminases cleave the acyl side chain (Amara et al.,

2011). It may also be possible to degrade QS molecules by increasing pH and using enzymes from QS degrading bacteria to control bacterial QS (Defoirdt et al. 2004).

FCUP 44 Chapter 2

Although more studies are needed to fully realize the potential of targeting QS for antifouling purposes, possible applications of QS inhibitors have attracted a lot of research, and main applications of QS inhibitors include therapeutic use of QS components to scale- up production of natural or engineered bacterial products as well biofouling control (Hentzer and Givskov, 2003). The attachment of microorganisms on the surface of sold matrixes is the main hindrance during the operation of membrane bioreactors and AHL autoinducers have been commonly found during biofouling of this surfaces. Interference with quorum sensing has the benefit of greater anti-fouling resistance, low toxicity and less stimulation of bacterial resistance (Xiong and Liu, 2010). Previous studies already showed the potential of using QS as a method of biofilm formation to reduce the biofouling of reverse osmosis membranes by controlling the presence of AHL’s. The use of QQ compounds as a biofouling-control agent in membrane bioreactors was reported by Yeon et al., 2009 in which acylase I-immobilized nanofiltration membrane was used. AHLs C6-HSL and C8-HSL were detected and correlated with the degree of fouling, which could be reduced by disrupting the quorum sensing activity by the acylases (Yeon et al., 2009). Commercially available vanillin has shown promise has a natural quorum quenching compound, being able to suppress up to 97% of biofilm formation in a reverse osmosis membrane in a reactor (Kappachery et al., 2010). Another study also used vanillin to interfere with AHLs which resulted in a significant decrease of biofilm formation by Aeromonas hydrophila (Ponnusamy et al., 2010, 2013). A recent study found that non-halogenated furanones blocked the production of AHL and inhibited the formation of biofilms in a reverse osmosis membrane (Kim et al., 2011).Oh et al., 2013 employed a quorum quenching bacteria strain (Rhodococcus sp. BH4),which has degrading activity against C8-HSL, in membrane reactors, substantially delaying the development of biofilm in a membrane reactor. Although quorum sensing compounds have been described as having potential and have already been applied in bioreactor fouling situations, no similar progress has yet been made for marine biofouling (Dobretsov et al., 2011). The main constraint is the lack of stability of enzymes within anti-fouling paints as well as their possible limited activity in seawater (Bzdrenga et al., 2017). QQ enzymes from extremophile

FCUP 45 Chapter 2 organisms can constitute promising candidates as they usually are highly robust and may be active in non-conventional environments (Choudhary et al., 2010). A promising work tested the effect of known QS blockers 5-hydroxy-3[(1R)-1- hydroxypropyl]-4-methylfuran-2(5H)- one (FUR1), (5R)-3,4-dihydroxy-5-[(1S)- 1,2-dihydroxyethyl]furan-2(5H)-one (FUR2) and triclosan (TRI) on the formation of bacterial biofilms. The tested QS blockers caused changes in bacterial density and bacterial community structure of the biofilms as well affecting larval attachment on these modified biofilms (Dobretsov et al., 2007). Another work involved the use of Kojic acid, which had been reported as a non-toxic QS inhibitor (Dobretsov et al.,

2011). Kojic acid incorporated into painting in glass slides and inhibited microfouling growth of bacteria and of the diatom Amphora coffeaeformis over a period of one month (Dobretsov et al., 2011).

Quorum sensing inhibition strategies are important because they can reduce QS- regulated biofilm formation involved in microfouling but may also impact the attraction and fixation of macrofouling species. QQ enzymes can constitute an environmentally friendly solution as they may be active while incorporated irreversibly into paints or coatings, and their action would be localized to the ship hull. Furthermore, in case of releasing, enzymes could be degraded in the environment and would not bio-accumulate (Olsen et al., 2007;

Bzdrenga et al., 2017). Alternatively, immobilized quorum-quenching enzymes or marine microbes that secrete QS inhibitors may be incorporated into anti-fouling treatments (Callow and Callow, 2011). However, the major problems of this technology are its low durability, and high cost (Siddiqui et al., 2015). It is also necessary to determine their effect in environment and the ecosystem, as quorum quenching activity may potentially inhibit all QS regulated pathways (Bzdrenga et al., 2017). Quorum sensing inhibition compounds can be used to prevent biofouling formation or to destroy biofilms. However, mature biofilms remain difficult to degrade and therefore acting early in biofilm formation is generally better.

Different studies have also demonstrated the influence of bacterial biofilm on the settlement of spores from algae or others (Tait et al., 2005) and considered the incorporation of QS inhibition compounds into paints or coatings as non-toxic antifouling alternatives (Dobretsov

FCUP 46 Chapter 2 et al., 2007, 2011). We envision that QS-inhibition approaches will most likely not fully prevent the settlement of macrofouling but may be a powerful strategy if used in combination with other approaches.

2.8 Concluding remarks

It is acknowledged that marine biofilms constitute highly dynamic communities, whose structure changes over time and with the interaction with different environmental parameters. The technical ability to characterize the composition of biofilm communities in environmental samples has increased substantially through recent high-throughput next generation gene sequencing, enabling ecologically-relevant field-based studies. However, most efforts to characterize microbial biofilms in the marine environment focused on substrata such as mats, sediments, rocks, organisms, and not much attention has been given to biofilm growth on artificial surfaces. Filling in this gap will surely strongly impact the development of effective anti-biofouling strategies. Therefore, we argued that marine biofilms from different climates and environmental origins should be characterized to convey a real portrait of their overall diversity. This approach should be complemented with functional data (e.g. transcriptomics) and integrated with chemical ecology, as the different systems become better characterized. Despite some recent intensive research, few biofilm chemical cues which can stimulate larval or spore settlement or their morphogenesis have been identified. Previous studies involving biofilm-macrofouler interactions substantiate the existence of a significant relationship between biofilms and macroinvertebrates mediated by chemical signaling, that is still however ill-characterized. Along these lines, knowledge on quorum QS activity and QS inhibition in marine microorganisms will be crucial for a more in-depth understanding of the development of marine biofilms communities. With such knowledge, controlling, disrupting or manipulating the development of marine biofouling

FCUP 47 Chapter 2 communities may become accessible through the action of small-molecules or enzymes that exist in nature. Lastly, we consider that as revealed by substantial past research, marine organisms have a great potential as a source of these products, and their prospective use still remains underexplored.

Acknowledgements

This work was supported by FCT - Foundation for Science and Technology - under project UID/Multi/04423/2013 and by the Structured Program of R&D&I INNOVMAR -

Innovation and Sustainability in the Management and Exploitation of Marine Resources

(reference NORTE-01-0145-FEDER-000035, Research Line NOVELMAR), which was funded by the Northern Regional Operational Programme (NORTE2020) through the

European Regional Development Fund (ERDF) and by the project CVMAR+I

(0302_CVMAR_I_1_P) funded by the program Interreg V Portugal-Spain (POCTEP) through the ERDF. J. Antunes acknowledges a PhD scholarship SFRH/BD/99003/2013 from

FCT (Fundação para a Ciência e Tecnologia)

2.9 References

Abarzua S., Kacan, S., Fuchs, P. (1999) Status and potential of natural product antifoulants. In: Fingerman M, Nagabhushanam R, Thompson F, editors. Recent advances in marine biotechnology. Vol. 3. Biofilms, bioadhesion, corrosion, and biofouling. Enfield, NH: Science Publishers Inc. pp 37 – 65

Alagely A., Cory J. K., Ritchie,K.B., Teplitski, M (2011). Signaling-Mediated Cross-Talk Modulates Swarming and Biofilm Formation in a Coral Pathogen Serratia Marcescens. The ISME Journal 5:1609–20.

FCUP 48 Chapter 2

Allison, D.G. (2002) The bioﬁlm matrix. Biofouling 19:139–150.

Almeida, J. R., Vasconcelos, V (2015) Natural antifouling compounds: Effectiveness in preventing invertebrate settlement and adhesion. Biotechnology Advances 33: 343–357.

Amara, N., Krom, B.P., Kaufmann, G.F., Meijler, M.M. (2011) Macromolecular inhibition of quorum sensing: enzymes, antibodies, and beyond. Chem Rev 111: 195–208.

Antunes, L., Caetano, M., Rosana, B. R., Ferreira, B.R. Buckner, M.C.C., Finlay, B.B. (2010) Quorum Sensing in Bacterial Virulence. Microbiology 156: 2271–82. doi:10.1099/mic.0.038794-0.

Bassler, B.L., Wright, M., Silverman, M.R. (1994) Multiple signaling systems controlling expression of luminescence in Vibrio harveyi: sequence and function of genes encoding a second sensory pathway. Mol Microbiol 13:273–86.

Bassler, B.L. Losick, R. (2006) Bacterially speaking. Cell 125:237–246.

Bao, W.Y., Satuito, C.G., Yang, J.L., Kitamura, H. (2007) Larval settlement and metamorphosis of the mussel Mytilus galloprovincialis in response to bioﬁlms. Mar Biol. 150:565-574.

Baragi, L.V., Chandrashekar A. (2016) Synergistic effect of elevated temperature, pCO2 and nutrients on marine biofilm. Mar Pollut Bull 105(1): 102-9.

Battin, T.J., Sloan, W.T., Kjelleberg, S., Daims H., Head, I.M., Curtis, T.P. (2007) Microbial landscapes: new paths to bioﬁlm research. Nat. Rev. Microbiol 5: 76–81.

Beckmann, M., Harder, T., Qian, P.Y. (1999) Induction of larval settlement and metamorphosis in the serpulid polychaete Hydroides elegans by dissolved amino acids: mode of action in laboratory bioassays. Mar Ecol Prog Ser 190: 167–178.

Bellou, N., Papathanassiou, E., Dobretsov, S., Lykoussis, V., Colijn. F. (2012) The effect of substratum type, orientation and depth on the development of bacterial deep-sea biofilm communities grown on artificial substrata deployed in the Eastern Mediterranean. Biofouling 28: 199-213.

Behrendt, L, Schrameyer, V, Ovortrup, K, Lundin, L, Søren, J, Sørensen, J, Larkum, AWD, Kü hl, M. (2012) Biofilm Growth and Near-Infrared Radiation-Driven Photosynthesis of the Chlorophyll d-Containing Cyanobacterium Acaryochloris marina. Appl. Environ Microbiol 78: 3896-3904.

FCUP 49 Chapter 2

Besemer, K., Peter, H., Logue, J.B., Langenheder, S., Lindstorm, E.S., Tranvik, L.J. (2012) Unraveling assembly of stream biofilm communities. ISME J 6: 1459–6.

Bodor, A., Elxnat, B., Thiel, V., Schulz, S., Wagner-Döbler, I., (2008) Potential for luxS related signalling in marine bacteria and production of autoinducer-2 in the genus Shewanella. BMC Microbiol 8: 13–22.

Boyer , M., Wisniewski-Dye F. (2009) Cell-cell signalling in bacteria: not simply a matter of quorum. FEMS Microbiol Ecol 70:1–19.

Brango-Vanegas, J., Costa, G., Ortmann, C., Schenkel, E., Reginatto, F., Ramos, F., Arévalo- Ferro C, Castellanos, L. (2014) Glycosylflavonoids from Cecropia pachystachya Trécul are quorum sensing inhibitors. Phytomedicine 21:670-675.

Briand, J., Djeridi I., Jamet, D., Coupé, S., Bressy, C., Molmeret, M., Le Berre, B., Rimet F., Bouchez A., Blache, Y. (2012) Pioneer Marine Biofilms on Artificial Surfaces Including Antifouling Coatings Immersed in Two Contrasting French Mediterranean Coast Sites. Biofouling 28: 53–63.

Briand, J.F., Barani, A, Garnier, C, Réhel, K, Urvois, F, LePoupon, C, Bouchez, A, Debroas, D, Bressy, C (2017) Spatio-Temporal Variations of Marine Biofilm Communities Colonizing Artificial Substrata Including Antifouling Coatings in Contrasted French Coastal Environments. Microb Ecol 74:585-598.

Brian-Jaisson, F., Ortalo-Magné A., Guentas-Dombrowsky L., Armougom, F., Blache Y., Molmeret, M. (2014). Identification of Bacterial Strains Isolated from the Mediterranean Sea Exhibiting Different Abilities of Biofilm Formation. Microb Ecol 68: 94–110.

Brown, N.E.M., Milazzo M., Rastrick, S.P.S., Hall-Spencer J.M., Therriault, T.W., Harley, C.G.D. (2017). Natural acidificationchanges the timing and rate of succession, alters community structure, and increases homogeneity in marine biofouling communities. Glob Chang Biol 24:112-127.

Burmølle, M., Webb, J.S., Rao, D., Hansen, L.H., Sørensen, S.J. (2006) Enhanced biofilm formation and increased resistance to antimicrobial agents and bacterial invasion are caused by synergistic interactions in multispecies biofilms. Appl Environ Microbiol 72: 3916–3923.

Bzdrenga J, Daudé D, Rémy B, Jacquet P, Plener L, Elias M, et al. (2017) Biotechnological applications of quorum quenching enzymes. Chem Biol Interact. 267:104-115.

Camps M., Barani A., Gregori G., Bouchez A., le Berre B., Bressy C,. Blache Y., Briand J.F. (2014). Antifouling Coatings Influence Both Abundance and Community Structure of

FCUP 50 Chapter 2

Colonizing Biofilms: A Case Study in the Northwestern Mediterranean Sea. Appl Environ Microbiol 80: 4821–31.

Callow, J.A., Callow, M.E. (2011) Trends in the development of environmentally friendly fouling-resistant marine coatings. Nat Commun 2: 244.

Cao, X., Pettitt, M.E., Wode, F., Sancet, M.P.A., Fu, J., Ji, J., Callow, M.E., Callow, J.A., Rosenhahn, A., Grunze, M. (2010) Interaction of zoospores of the green alga Ulva with bioinspired micro- and nanostructured surfaces prepared by polyelectrolyte layer-by-layer self-assembly. Adv Funct Mater 20:1984–1993.

Cassé F., Swain G.W. (2006) The Development of Microfouling on Four Commercial Antifouling Coatings under Static and Dynamic Immersion. International Biodeterioration and Biodegradation 57: 179–85.

Celikkol-Aydin, S., Gaylarde,. C., Lee, T., Melchers, R.E., Witt, D.L., Beech, I.B. (2016) 16S rRNA gene profiling of planktonic and biofilm microbial populations in the Gulf of Guinea using Illumina NGS. Mar Environ Res.122:105-112.

Chen, C., Lung, J, Maki, S, Rittschof, D Teo, SLM. (2013) Early Marine Bacterial Biofilm on a Copper-Based Antifouling Paint. International Biodeterioration and Biodegradation 83: 71–76..

Chiu, J.M. Y, Zhang, R., Wan, H., Thiyagarajan, V., Qian, P.Y. (2008) Nutrient Effects on Intertidal Community: From Bacteria to Invertebrates. Marine Ecology Progress Series 358: 41–50.

Choo, J.H., Rukayadi, Y., Hwang, J.K. (2006) Inhibition of bacterial quorum sensing by vanilla extract. Lett Appl Microbiol 42:637–41

Choudhary, S., Schmidt-Dannert, C. (2010) Applications of quorum sensing in biotechnology. Appl Microbiol Biotechnol 86:1267–1279.

Chung, H.C., Lee, O.O., Huang, Y.L., Mok, S.Y., Kolter R., Qian, P.Y. (2010) Bacterial Community Succession and Chemical Profiles of Subtidal Biofilms in Relation to Larval Settlement of the Polychaete Hydroides Elegans. The ISME Journal 4: 817–28.

Cooksey, K.E., Wigglesworth-Cooksey, B. (1995). Adhesion of Bacteria and Diatoms to Surfaces in the Sea: A Review. Aquatic Microbial Ecology 9: 87–96.

Cooksey, K.E., Wigglesworth-Cooksey, B. (2001) Diatoms in bioﬁlms. In: Flemming H-C, editor. Encyclopedia of environmental microbiology. New York: Wiley. p. 1052– 1063.

FCUP 51 Chapter 2

Costas, E., Gonzalez, R., López-Rodas, V., Emma Huertas, I (2013) Mutation of Microalgae from Antifouling Sensitivity to Antifouling Resistance Allows Phytoplankton Dispersal through Ships’ Biofouling. Biological Invasions 15: 1739–50.

Cuadrado-Silva, C.T., Castellanos, L., Arévalo-Ferro, C.,Osorno, O.E. (2013) Detection of Quorum Sensing Systems of Bacteria Isolated from Fouled Marine Organisms. Biochemical Systematics and Ecology 46: 101–7.

Dafforn, K.A., Lewis, J.A., Johnston, E.L., (2011) Antifouling strategies: history and regulation, ecological impacts and mitigation. Mar Pollut. Bull 62: 453–465.

Dahms, H.U., Xu, Y., Pfeiﬀer, C. (2006) Antifouling potential of cyanobacteria. Biofouling 22:317–327.

Dang, H., Lovell, C.H. (2000) Bacterial Primary Colonization and Early Succession on Surfaces in Marine Waters as Determined by Amplied rRNA Gene Restriction Analysis and Sequence Analysis of 16S rRNA Genes. Appl Environ Microbiol 66: 467–75.

Dang, H., Lovell, C.H. (2002) Numerical Dominance and Phenotype Diversity of Marine Rhodobacter Species during Early Colonization of Submerged Surfaces in Coastal Marine Waters as Determinated by 16S Ribosomal DNA Sequence Analysis and Fluorescence in Situ Hybridization. Appl Environ Microbiol 68: 496–504.

Dang, H., Li, T., Chen, M., Huang, G. (2008) Cross-Ocean Distribution of Rhodobacterales Bacteria as Primary Surface Colonizers in Temperate Coastal Marine Waters. Appl Environ Microbiol 74:52–60.

Dang, H., Chen, R., Wang, L., Shao, S., Dai, L., Ye, Y. (2011) Molecular characterization of putative biocorroding microbiota with a novel niche detection of Epsilon- and Zetaproteobacteria in Pacific Ocean coastal seawaters. Environ Microbiol 13:3059 – 74.

Dang, H., and Lovell, C. R. (2016). Microbial surface colonization and biofilm development in marine environments. Microbiol Mo Bio Rev 80: 91–138.

Daniels, R., Vanderleyden, J., Michiels , J. (2004) Quorum sensing and swarming migration in bacteria. FEMS Microbiol Rev 28: 261–289.

Davey, M.E., O'Toole, G.A (2000) Microbial biofilms: from ecology to molecular genetics. Microbiol. Mol Biol Rev 64: 847-867.

Decho, A.W., Visscher, P.T., Ferry, J., Kawaguchi, T., He, L., Przekop, K.M., Norman, R.S., Reid, R.P. (2009) Autoinducers extracted from microbial mats reveal a surprising diversity

FCUP 52 Chapter 2 of N-acylhomoserine lactones (AHLs) and abundance changes that may relate to diel pH. Environ Microb 11:409–420.

Defoirdt, T., Sorgeloos, P., Bossier, P. (2011) Alternatives to antibiotics for the control of bacterial disease in aquaculture. Curr Opin Microbiol 14:251–258.

DeLong, (2005) Microbial Community Genomics in the Ocean. Nat Rev Microbiol 3: 459– 69.

De Tender, C., Devriese, L., Haegman, A., Maes, S., Vangeyte, J., Cattrijsee, A., Dawyndt, P., Rutttink, T. (2017) Temporal Dynamics of Bacterial and Fungal Colonization on Plastic Debris in the North Sea. Environ Sci Technol. 51:7350-7360.

Doberva, M. S., Sanchez-Ferandin, E., Toulza, P., Lebaron, and Lami R. (2015). Diversity of Quorum Sensing Autoinducer Synthases in the Global Ocean Sampling Metagenomic Database. Aquatic Microbial Ecology 74: 107–19.

Dobretsov, S.V., Qian, P.Y., Wahl, M.(2005) Effect of solar ultraviolet radiation on the formation of shallow, early successional biofouling communities in Hong Kong. Mar Ecol Prog Ser 290:55–65

Dobretsov, S., Dahms, H.U., Qian, P.Y.(2006) Inhibition of Biofouling by Marine Microorganisms and Their Metabolites. Biofouling 22: 43–54.

Dobretsov, S., Dahms, H.U., Yili, H.,Wahl, M., Qian, P.Y. (2007) The Effect of Quorum- Sensing Blockers on the Formation of Marine Microbial Communities and Larval Attachment. FEMS Microbiol Ecol 60: 177–88.

Dobretsov, S., Xiong, H., Xu, Y., Levin, L., Qian, P.Y. (2007) Novel Antifoulants: Inhibition of Larval Attachment by Proteases. Mar Biotechnol 9: 388–97.

Dobretsov, S.,Teplitski, M., Paul, C. (2009) Mini-Review: Quorum Sensing in the Marine Environment and Its Relationship to Biofouling. Biofouling 25: 413–27.

Dobretsov, S., Teplitski, M., Alagely, A., Gunasekera, S.P., Paul,V.J. (2010) Malyngolide from the Cyanobacterium Lyngbya Majuscula Interferes with Quorum Sensing Circuitry. Environ Microbiol Rep. 2: 739–44.

Dobretsov, S., Teplitski, M., Bayer, M., Gunasekra, S., Proksch, P., Paul, V.J. (2011) Inhibition of marine biofouling by bacterial quorum sensing inhibitors. Biofouling: 27:893– 905.

FCUP 53 Chapter 2

Dong, Y.H., Wang, L.H., Xu, J.L. et al. (2001) Quenching quorum-sensing-dependent bacterial infection by an N-acyl homoserine lactonase. Nature 411: 813-817.

Dong, Y.H., Wang, L.H., Zhang, L.H. (2007) Quorum-quenching microbial infections: mechanisms and implications. Philos T Roy Soc B 362: 1201–1211.

Donlan, R.M. (2002) Bioﬁlms: Microbial Life on Surfaces Emerg Infect Dis 8:881-890.

Dunne, W.M.(2002) Bacterial adhesion: seen any good biofilms lately? Clin Microbiol Rev.15:155-66.

Elifantz, H, Horn, G, Ayon, M., Cohen, Y., Minz, D. (2013) Rhodobacteraceae are the key members of the microbial community of the initial biofilm formed in Eastern Mediterranean coastal seawater. FEMS Microbiol Ecol 85:348–357.

Falciatore, A., Bowler, C. (2002) Revealing the Molecular Secrets of Marine Diatoms. Annual Review of Plant Biology 53: 109–30.

Flemming, H.C., Wingender, J. (2010) The Biofilm Matrix. Nature Reviews. Microbiology 8: 623–33.

Flemming, H.C., Wingender, J., Szewzyk, U., Steinberg, P., Rice, S.A., Kjelleberg, S., (2016) Biofilms: an emergent form of bacterial life. Nat. Rev. Microbiol. 14: 563-575.

Freckelton, M.L., Nedved, B.T., Hadfield, M.G. (2017) Induction of Invertebrate Larval Settlement; Different Bacteria, Different Mechanisms? Sci Rep. 14:42557.

Fusetani, N. (2004) Biofouling and antifouling. Natural Product Reports 21: 94-104.

Fusetani, N. (2011) Antifouling marine natural products. Nat Prod Rep 28: 400–41

Galkin, M., Ivanitsia, V., Ishkov, Y., Galkin, B., and Filipova, T. (2015) Characteristics of the Pseudomonas aeruginosa PA01 intercellular signaling pathway (quorum sensing) functioning in presence of porphyrins bismuth complexes. Pol J Microbiol. 64: 101–106.

Ganesan, A.M., Alfaro, A.C., Brooks, J.D., Higgins, CM. (2010) The role of bacterial biofilms and exudates on the settlement of mussel (Pernacanaliculus) larvae. Aquaculture 306: 388–392.

Ganesan, A.M., Alfaro, A.C., Brooks, J.D., Higgins, C.M. (2012) Characterization of bioﬁlm exudates and their effects on settlement of mussel (Perna canaliculus) larvae. Journal of Experimental Marine Biology and Ecology 434–435: 34–46.

FCUP 54 Chapter 2

Givskov M, Eberl L, Molin S. (1997) Control of exoenzyme production, motility and cell differentiation in Serratia liquefaciens. FEMS Microbiol Lett 148:115–122.

Goecke, F., Labes, A., Wiese, J., Imhoff, J.F. (2010) Review Chemical Interactions between Marine Macroalgae and Bacteria.Marine Ecology Progress Series 409: 267–300.

Guo, X., Zuo-shin, N., Da-pei, L., Feng, J., Chen, Y., Tou, F., Liu, M., Yi, Y. (2017) Bacterial community structure in the intertidal biofilm along the Yangtze Estuary, China. Mar Pollut Bull124:314-320.

Guttenplan, S.B., Kearns D.B. (2013) Regulation of Flagellar Motility during Biofilm Formation. FEMS Microbiology Revs 37: 849–71.

Hadﬁeld, MG. (2011) Bioﬁlms and marine invertebrate larvae: what bacteria produce that larvae use to choose settlement sites. Ann. Rev. Mar. Sci 3: 453–470.

Hadfield, M.G., Nedved, B.T., Wilbur, S., Koehl, M.A.R. (2014) Biofilm cue for larval settlement in Hydroides elegans (Polychaeta): is contact necessary? Mar Biol. 161:2577– 2587.

Hall-Stoodley, L., Costerton, J., Stoodley, W., Paul, V (2004) Bacterial bioﬁlms: from the natural environment to infectious diseases. Nat. Rev. Microbiol. 2: 95–108.

Harder, T., Lam C, M., Qian, P.Y. (2002) Induction of Larval Settlement in the Polychaete Hydroides Elegans by Marine Biofilms: An Investigation of Monospecific Diatom Films as Settlement Cues. Marine Ecology Progress Series 229: 105–12.

Haslbeck, E, G., Bohlander, G. (1992) Microbial biofilm effects on drag-lab and field. In Proceedings 1992 SNAME Ship Production Symposium. New Orleans, LA.

Hellio, C., Thomas-Guyon, H., Culioli, G., Piovetti, L., Bourgougnon, N., Le Ga, Y. (2001) Marine antifoulants from Bifurcaria bifurcata (Phaeophyceae, Cystoseiraceae) and other brown macroalgae. Biofouling 17:189–201.

Hentzer, M., Givskov, M. (2003) Pharmacological inhibition of quorum sensing for the treatment of chronic bacterial infections. J Clin Invest 112: 1300–1307.

Hmelo, L.R. Quorum sensing in marine microbial environments (2017) Annu Rev Mar Sci 9: 257–281.

Hoang, H.H., Gurich, N., Gonzalez, J.E. (2008) Regulation of motility by the ExpR/Sin quorum-sensing system in Sinorhizobium meliloti. J Bacteriol 190: 861–871.

FCUP 55 Chapter 2

Holm, E. R., Schultz, M. P., Haslbeck, E. G., Talbott, W. J., Field, A. J. (2004) Evaluation of hydrodynamic drag on experimental fouling-release surfaces, using rotating disks. Biofouling 20: 219–226.

Hölscher, T, Bartels, B., Lin, Y.C., Gallegos-Monterrosa, R., Price-Whelan, A., Kolter, R., Dietric, L.E.P., ,Kovács, A.P. (2015) Motility, Chemotaxis and Aerotaxis Contribute to Competitiveness during Bacterial Pellicle Biofilm Development. J Mol Biol 427:3695-3708

Huang, S., Hadﬁeld, M. (2003) Composition and density of bacterial bioﬁlms determine larval settlement of the polychaete Hydroides elegans. Mar Ecol Prog Ser 260: 161–172.

Huggett, M.J., Williamson, J.E., de Nys,R., Kjelleberg, S., Steinberg, P. D. (2006) Larval settlement of the common Australian sea urchin Heliocidaris erythrogramma in response to bacteria from the surface of coralline algae. Oecologia 149:604 – 19.

Huggett, M.J., Nedved, B.T., Hadfield, M.G. (2009) Effects of initial surface wettability on biofilm formation and subsequent settlement of Hydroides elegans. Biofouling 25:387–399.

Hung, OS, Lee, OO, Thiyagarajan, V., He, HP, Xu, Y., Chung, H.C. (2009) Characterization of cues from natural multi-species bioﬁlms that induce larval attachment of the polychaete Hydroides elegans. Aquat Biol 4: 253– 262.

Inbakandan, D., P Sriyutha Murthy, R., Venkatesan, Khan, S.J. (2010) 16S rDNA Sequence Analysis of Culturable Marine Biofilm Forming Bacteria from a Ship’s Hull. Biofouling 26: 893–99.

Jin, T., Qian, P.Y. (2005) Amino acid exposure modulates the bioactivity of bioﬁlms for larval settlement of Hydroides elegans by altering bacterial community components. Mar Ecol Prog Ser 297: 69–179.

Joint, I., Callo, M.E, Callow, J., Clarke, R.,K.. (2000) Zoospores To a Bacterial Biofilm Assemblage. Biofouling 16: 151–58.

Joint, I., Tait, K., Wheeler, G. (2007) Cross-Kingdom Signalling: Exploitation of Bacterial Quorum Sensing Molecules by the Green Seaweed Ulva. Philos Trans R Soc Lond B Biol Sci 362: 1223–33.

Jones, P.R., Cottrell, M.R., Kirchman, D.L, Dexter, S.C. (2007) Bacterial Community Structure of Biofilms on Artificial Surfaces in an Estuary. Microb Ecol 53: 153–62.

Jouuchi T., Satuito, C.G., Kitamura H. (2007) Sugar compound products of the periphytic diatom Navicula ramosissima induce larval settlement in the barnacle, Amphibalanus amphitrite. Mar Biol 152:1065-1072.

FCUP 56 Chapter 2

Kamino, K. (2013) Mini-Review: Barnacle Adhesives and Adhesion. Biofouling 29:735-49.

Kanagasabhapathy, M. G. Yamazaki, A., Ishida,, H, Nagata, S. (2009) Presence of Quorum- Sensing Inhibitor-like Compounds from Bacteria Isolated from the Brown Alga Colpomenia Sinuosa. Lett Appl Microbiol 49: 573–79.

Kappachery, S., Paul, D., Yoon, J, Kweon, J.H. (2010) Vanillin, a potential agent to prevent biofouling of reverse osmosis membrane. Biofouling 26:667–72.

Keller, M,Zengler,K. (2004) Tapping into Microbial Diversity. Nat Rev Microbiol 2: 141– 50.

Khandeparker, L., Anil, A.C., Raghukumar, S. (2002) Factors regulating the production of different inducers in Pseudomonas aeruginosa with reference to larval metamorphosis in Balanus amphitrite. Aquat Microb Ecol 28:37-54.

Khandeparker, L, A, Raghukumar, S. (2006) Relevance of biofilm bacteria in modulating the larval metamorphosis of Balanus amphitrite. FEMS Microbiol Ecol 58: 425–438.

Kjelleberg, S., Steinberg, P., Givskov, M., Gram, L., Manefield, M., De Nys, R.,(1997) Do Marine Natural Products Interfere with Prokaryotic AHL Regulatory Systems? Aquat Microb Ecol 13: 85–93.

Kjelleberg, S., Molin, S. (2002) Is there a role for quorum sensing signals in bacterial biﬁlms? Curr. Opin. Microbiol 5: 254–258.

Kim, S.J., Lee, S.Y., Hong, S.K., Oh, Y.S., Seoul, M.J., Kweon, J.H., Kim, T.H. (2009) Biofouling of reverse osmosis membranes: Microbial quorum sensing and fouling propensity. Desalination 247: 303–315.

Kim, J.H., Choi, D.C., Yeon, K.M., Kim, S.R.; Lee, C.H. (2011) Enzyme-immobilized nanofiltration membrane to mitigate biofouling based on quorum quenching. Environ Sci Technol 45: 1601–1607.

Klein, G., Pierre, L., Bellon-Fontain,G., Zhao, M.N., Breret, J.M., Maugard, M, Graber, T (2014) Marine diatom Navicula jeffreyi: from biochemical composition and physicochemical surface properties to understanding the first step of benthic biofilm formation. Journal of Adhesion Science and Technology 28: 739-1753

Lagos, M.E., White, C.R., Marshall, D.J.( 2016) Biofilm history and oxygen availability interact to affect habitat selection in a marine invertebrate Biofouling 32: 645-655.

FCUP 57 Chapter 2

Lam, C., Harder, T., Qian, P.Y. (2005) Induction of larval settlement in the polychaete Hydroides elegans by extracellular polymers of benthic diatoms. Mar Ecol Prog Ser 286: 145–154.

Landini, P., Antoniani, D., Burgess, J., Nijland, R. (2010) Molecular mechanisms of compounds affecting bacterial biofilm formation and dispersal. Appl Microbiol Biotechnol 86: 813–823.

Landoulsi, J., Cooksey, K.E., Dupres, V. (2011) Review – Interactions between Diatoms and Stainless Steel: Focus on Biofouling and Biocorrosion. Biofouling 27: 1105–24..

Lau, S.C., Mak, K.K., Chen, F., Qian, P.Y. (2002) Bioactivity of bacterial strains isolated from marine biofilms in Hong Kong waters for the induction of larval settlement in the marine polychaete Hydroides elegans. Mar Ecol Prog Ser 226:301–310.

Lau, S.C.K., Vengatesen, T.,, Cheung SCK, Qian, P.Y. (2005) Roles of Bacterial Community Composition in Biofilms as a Mediator for Larval Settlement of Three Marine Invertebrates. Aqua MicrobiolEcology 38: 41–51.

Lawes, J.C, Neilan B.A Brown M.V, Clark G.F, Johnston E.L. (2016) Elevated nutrients change bacterial community composition and connectivity: high throughput sequencing of young marine biofilms. Biofouling 32: 57-69.

Leão, P.N., Engene, N, Antunes, A., Gerwick, W.H., Vasconcelos, V. (2012) The chemical ecology of cyanobacteria. Nat Prod Rep 29: 372–391.

Leary, D., Li, R.W., Hamdan, L.J., Hervey, W.J., Lebedev, Wang Z., Deschamps, J.R., Kusterbeck, A.W., Vora, G.J (2014) Integrated Metagenomic and Metaproteomic Analyses of Marine Biofilm Communities. Biofouling 30:1211-1223.

Lee, J.W., Ji, H.W., Yang, H.K., Lee, K.H., Lee (2008) Bacterial Communities in the Initial Stage of Marine Biofilm Formation on Artificial Surfaces. Journal of Microbiology.46: 174– 82.

Lee, O.O., Chung, H.C., Yang J., Wang, Y., Dash, S., Wang, .H, Qian, P.Y. (2014) Molecular techniques revealed highly diverse microbial communities in natural marine bioﬁlms on polystyrene dishes for invertebrate larval settlement. Microb Ecol 68:81 – 93.

Lee, Y.M., Cho, K.H., Hwang, K., Kim, E.H., Kim, M., Hong, S.G., Lee, H.K. (2016) Succession of bacterial community structure during the early stage of biofilm development in the Antarctic marine environment. Korean Journal of Microbiology: 52:1: 49.

FCUP 58 Chapter 2

Loeb G I, Laster D, Gracik T (1984) The influence of microbial fouling films on hydrodynamic drag of rotating discs. In: Costlow J D, Tipper R C (eds) Marine Biodeterioration: an Interdisciplinary Study. Naval Institute Press, Annapolis, MD: 88 – 9.

Lowery, C., Dickerson, T.J., Janda. (2008) Interspecies and Interkingdom Communication Mediated by Bacterial Quorum Sensing. Chem Soc Rev 37: 1337–46.

Madsen, J.S., Røder, H.L., Russel, K., Sørensen, H., Burmølle, M., Sørensen, S.J. (2016) Coexistence facilitates interspecific biofilm formation in complex microbial communities. Environ Microbiol. 18:2565-74

Magin, C.M., Finlay, J.A., Clay, G., Callow, M.E., Callow, J.A., Brennan, A.B. (2011) Antifouling performance of cross-linked hydrogels: refinement of an attachment model. Biomacromolecules 12:915–922.

Maki JS. 2002. Biofouling in the marine environment. In: Bitton G, editor. Encyclopedia of environmental microbiology. New York (NY): Wiley; p. 610–619. Martínez-García M, Koblížek M, López-Lege.

Matz, C., Kjelleberg, S. (2005) Off the hook – how bacteria survive protozoan grazing. Trends Microbiol 13:302 – 307.

Matz, C.,Webb, J.S., Schupp, P.J., Phang, P.Y., Penesyan, A., Egan, S., Steinberg, P., Kjelleberg S. (2008) Marine Biofilm Bacteria Evade Eukaryotic Predation by Targeted Chemical Defense. PLoS ONE 3: 1–7.

McDougald, D., Rice, S.A., Barraud, N., Steinberg, P.D., Kjelleberg S. (2011) Should we stay or should we go: mechanisms and ecological consequences for bio ﬁ lm dispersal. Nat Rev Microbiol 10:39 – 50.

McLean, .R.J.C., Whiteley, M., Stickler, D.J., Fuqua, W.C. (1997) Evidence of autoinducer activity in naturally occurring bioﬁlms. FEMS Microbiol Lett 154: 259–263.

Mieszkin, S., Pierre, M., Callow, M.E., Callow, J. (2012) Effect of bacterial biofilms formed on fouling-release coatings from natural seawater and Cobetia marina, on the adhesion of two marine algae. Biofouling 28: 953–68.

Mieszkin , S., Callow , M.E., Callow , J.A.( 2013) Interactions between microbial bioﬁlms and marine fouling algae: a mini review. Biofouling 29:1097 – 1113.

Milton, DL. (2006) Quorum sensing in vibrios: complexity for diversification. Int J Med Microbiol 296:61–71.

FCUP 59 Chapter 2

Mitbavkar, S., Anil, A.C. (2007) Species Interactions within a Fouling Diatom Community: Roles of Nutrients, Initial Inoculum and Competitive Strategies Biofouling 23: 99–112.

Mohamed, N.M., Cicirelli, E.M., Kan, J., Chen, F., Fuqua, C.,Hill., R.T. (2008) Diversity and Quorum-Sensing Signal Production of Proteobacteria Associated with Marine Sponges. Environ Microbiol 10: 75–86.

Molino, P. J., Wetherbee, R. (2008). The biology of biofouling diatoms and their role in the development of microbial slimes. Biofouling 24:365–379.

Moss, J., Nocker, A., Lepo, J.E., Snyder, R.A. (2006) Stability and Change in Estuarine Biofilm Bacterial Community Diversity. Appl Environ Microbiol 72: 5679–88.

Muras, A., Lopez-Perez, M., Mayer, C., Parga, A., Amaro-Blanco, J., Otero, A. (2018) High prevalence of quorum-sensing and quorum-quenching activity among cultivable bacteria and metagenomic sequences in the Mediterranean Sea. Genes 9: 100.

Nagar, E., Schwarz, R. (2015) To be or not to be planktonic? Self-inhibition of biofilm development. Environ Microbiol 17:1477–1486.

Nayar, S., Goh, C.L., Chou, L.M. (2005) Settlement of marine periphytic algae in a tropical estuary. Estuarine Coast Shelf Sci 64: 241–248.

Nocker, A. Lepo, J.E., Snyder, R.A. (2004) Inﬂuence of an oyster reef on development of the microbial heterotrophic community of an estuarine bioﬁlm. Appl Enviro Microbiol. 70: 6834–6845.

Olivier, F., Tremblay, R., Bourget, E., Ritschoff, D. (2000) Barnacle settlement: field experiments on the influence of larval supply, tidal level, biofilm quality and age on Balanus amphitrite cyprids. Mar Ecol Prog Ser 199:185–204.

Olsen SM, Pedersen LT, Laursen MH, Kiil S, Dam-Johansen K (2007) Enzyme-based antifouling coatings: a review. Biofouling 23:369–383

Park, S.R., Kang, Y.H., Choi, C.G. (2011) Biofilm: A crucial factor affecting the settlement of seaweed on intertidal rocky surfaces. Estuarine, Coastal and Shelf Science 91: 163-167.

Parsek, M.R.,Val, D.L., Hanzelka, B.L., Cronan,J.E., Greenberg.E.P .(1999) Acyl Homoserine-Lactone Quorum-Sensing Signal Generation. PNAS 96: 4360–65.

Patel, P., Callow, M.E.Joint, I.J., Callow, J.A. (2003) Specificity in the Settlement–modifying Response of Bacterial Biofilms towards Zoospores of the Marine Alga Enteromorpha. Environ Microbiol 5: 338–49.

FCUP 60 Chapter 2

Patil, J.S., Anil, A.C.,. (2005a) Biofilm Diatom Community Structure: Influence of Temporal and Substratum Variability. Biofouling 21: 189–206..

Patil, J. S., Anil. A.C. (2005b) Influence of Diatom Exopolymers and Biofilms on Metamorphosis in the Barnacle Balanus Amphitrite. Marine Ecology Progress Series 301: 231–45.

Pepe-Ranney C., Hall E. (2015) The effect of carbon subsidies on marine planktonic niche partitioning and recruitment during biofilm assembly. Front Microbiol 6:703.

Ponnusamy K., Paul D., Kweon J.H. (2009) Inhibition of quorum sensing mechanism and Aeromonas hydrophila biofilm formation by vanillin. Environ Eng Sci 26:1359–63.

Ponnusamy, K., Kappachery, S., Thekeettle, M., et al. (2013) Anti-biofouling property of vanillin on Aeromonas hydrophila initial biofilm on various membrane surfaces. World J Microb Biot 29: 1695–703.

Purohit, A. A., Johansen, J. A., Hansen, H., Leiros, H. K., Kashulin, A., Karlsen, C., et al. (2013). Presence of acyl-homoserine lactones in 57 members of the Vibrionaceae family. J. Appl. Microbiol 115: 835–847.

Qian, P. Y., Lau, S.C.K, Dahms, H.U., Dobretsov, S., Harder, T. (2007) Marine Biofilms as Mediators of Colonization by Marine Macroorganisms: Implications for Antifouling and Aquaculture. Marine Biotechnology 9: 399–410.

Qian, P.Y., Chen, L., Xu, Y. (2013) Mini-Review: Molecular Mechanisms of Antifouling Compounds. Biofouling 29 : 381–400.

Rampadarath, S., Bandhoa, K., Puchoaa, D., Jeewon, R., Bal., S. (2017) Early bacterial biofilm colonizers in the coastal waters of Mauritius. Electronic Journal of Biotechnology 29: 13-21.

Rasmussen, T.B. and Givskov, M. (2006) Quorum-sensing inhibitors as anti-pathogenic drugs. Int. J. Med. Microbiol. 296: 149–161

Rice, S. ,Givskov, M., Steinberg, P., Kjelleberg, S. (1999) Bacterial Signals and Antagonists: The Interaction between Bacteria and Higher Organisms. J Mol Microbiol Biotechnol.1: 23– 31.

Røder,H.L., Sørensen, SJ, Burmølle, M. (2016) Studying Bacterial Multispecies Biofilms: Where to Start? Trends Microbiol. 24:503-13.

FCUP 61 Chapter 2

Roeselers, G., Van Loosdrecht, M.C., Muyzerm G.(2007) Heterotrophic Pioneers Facilitate Phototrophic Biofilm Development. Microbial Ecology 54: 578–85.

Roeselers, G., Van Loosdrecht, M.C., Muyzer, G.(2008) Phototrophic biofilms and their potential applications". J Appl Phycol 20: 227 – 235

Romero, M., Martin-Cuadrado, A., Roca-Rivada, A., Cabello, A.M., Otero A. (2011) Quorum quenching in cultivable bacteria from dense marine microbial communities. FEMS Microbiol Ecol 75: 205–17.

Romero, M., Martin-Cuadrado, A.B. Otero, A. (2012) Determination of whether quorum quenching is a common activity in marine bacteria by analysis of cultivable bacteria and metagenomic sequences. Appl Environ Microbiol 78:6345–6348

Salta, M.,Wharton, J.A., Stoodley, P., Dennington, S.P, Goodes, L.R., Werwinski, S., Mart, U.,Wood, R.J.K.; Stokes, K.R. (2010) Designing biomimetic antifouling surfaces. Philos Trans A Math Phys Eng Sci 368: 4729–4754.

Salta, M., Wharton, J.A., Blache, Y., Stokes, K.R., Briand JF. (2013) Marine biofilms on artificial surfaces: structure and dynamics. Environ Microbiol 15:2879.

Sanli, K., Bengstsson-Palme, J., Nilsson, R.H., Kristiansson, E., Rosenblad, A., Blanck, H., Eriksson, K.M. (2015) Metagenomic sequencing of marine periphyton: taxonomic and functional insights into biofilm communities. Front Microbiol 30:1192.

Scardino, A.J., de Nys, R. (2011) Mini review: biomimetic models and bioinspired surfaces for fouling control. Biofouling 27:73–86.

Schauder, S., Shokat, K., Surette, M. G., Bassler, B. L. (2001) The LuxS family of bacterial autoinducers: biosynthesis of a novel quorum-sensing signal molecule. Mol Microbiol 41: 463–476.

Schultz, M.P., Swain, G.W. (2000) The influence of biofilms on skin friction drag. Biofouling. 15:129:139.

Schultz, M.P. (2007) Effects of coating roughness and biofouling on ship resistance and powering. Biofouling 23:331–41

Schultz, M.P., Bendick, J.A., Holm, E.R., Hertel, W.M. (2011) Economic impact of biofouling on a naval surface ship. Biofouling 27: 87–98.

Sherr, E B. (2002) Significance of Predation by Protists. Antonie van Leeuwenhoek 81: 293– 308.

FCUP 62 Chapter 2

Shikuma, N.J., Hadfield, M.G. (2005) Temporal variation of an initial marine biofilm community and its effects on larval settlement and metamorphosis of the tubeworm Hydroides elegans. Biofilms 2:231–238

Shikuma, N.J., Hadfield, M.J. (2010) Marine Biofilms on Submerged Surfaces Are a Reservoir for Escherichia Coli and Vibrio Cholerae. Biofouling 26 (1): 39–46.

Shnit-Orland, M, and Kushmaro, A. (2009) Coral mucus-associated bacteria: a possible ﬁrst line of defense. FEMS Microbiol Ecol 67: 371–380.

Singh R.P, Baghel, RS., Reddy, CRK, Jha, B. (2015) Eﬀect of quorum sensing signals produced by seaweed-associated bacteria on carpospore liberation from Gracilaria dura. Front Plant Sci 6:117.

Sneed, J.M., Sharp, K.H., Ritchie, K.B. , Paul, V.J.(2014) The chemical cue tetrabromopyrrole from a biofilm bacterium induces settlement of multiple Caribbean corals. Proc R Soc B: 281. 20133086.

Steinberg, P.D., De Nys R., Kjelleberg S. (2002) Chemical cues for surface colonization. J Chem Ecol 28: 1935–1951.

Surette, M. G., Miller, M. B., Bassler, B. L. (1999) Quorum sensing in Escherichia coli, Salmonella typhimurium, and Vibrio harveyi: a new family of genes responsible for autoinducer production. Proc Natl Acad Sci USA 96: 1639–1644.

Sutherland, I.W (2001) The Biofilm Matrix - An Immobilized but Dynamic Microbial Environment. Trends in Microbiology 9: 222–27.

Swanson, R.L., de Nys R., Huggett, M.J., Green, J.K., Steinberg, P.D. (2006) In situ quantification of a natural settlement cue and recruitment of the Australian sea urchin Holopneustes purpurascens. Mar Ecol Prog Ser 314:1–14

Sweat, , L.H., Johnson, K.B. (2013) The Effects of Fine-Scale Substratum Roughness on Diatom Community Structure in Estuarine Biofilms. Biofouling 29: 879–90.

Szewzyk, U., Holmstroem, C., Wrangstadh, M., Samuelsson, M.O., Maki, J.S., Kjelleberg S.(1991) Relevance of the exopolysaccharide of marine Pseudomonas sp. strain S9 for the attachment of Ciona intestinalis larvae. Mar Ecol Prog Ser 75:259–265

Tait, K., Joint,I., Daykin, M., Debra, L. Milton, P.W., Cámara, M. (2005) Disruption of Quorum Sensing in Seawater Abolishes Attraction of Zoospores of the Green Alga Ulva to Bacterial Biofilms. Environ Microbiol 7: 229–40.

FCUP 63 Chapter 2

Tait K., Holly Williamson, S.A., Williams P., Cámara M., Joint I. (2009) Turnover of Quorum Sensing Signal Molecules Modulates Cross-Kingdom Signalling Environ Microbiol 11: 1792–1802.

Tait, K., Hutchison, Z, Thompson, F.L., Munn, C.B. 2010. Quorum sensing signal production and inhibition by coral-associated vibrios. Environ Microbiol Rep 2:145–50

Tait, K., Havenhand, J. (2013) Investigating a possible role for the bacterial signal molecules N-acylhomoserine lactones in Balanus improvisus cyprid settlement. Mol Ecol 22:2588-602

Tan, D, Svenningsen, L., Middelboe, M. (2015) Quorum Sensing Determines the Choice of Antiphage Defense Strategy in Vibrio Anguillarum. mBio 6: 1–10.

Tan CH,, Koh KS, XiC, Zhang J, Xiao HT, Guo P Yan, Zhou, Wun, Jern, Wun, Rice, Scott A., Kjelleberg, S. (2015) Community quorum sensing signalling and quenching: microbial granular bioﬁlm assembly. Bioﬁlms and Microbiomes 15006.

Teasdale, M.E., Liu J., Wallace, J., Akhlaghi, F., Rowley, DC. (2009) Secondary Metabolites Produced by the Marine Bacterium Halobacillus Salinus That Inhibit Quorum Sensing- Controlled Phenotypes in Gram-Negative Bacteria. Appl Environ Microbiol 75: 567–72.

Tebben, J.D., Tapiolas, M. (2011) Induction of larval metamorphois of the coral Acropora millepora by tetrabromopyrrole isolated from a Pseudoalteromonas bacterium. PLoS ONE 6: e19082.

Toupoint, N., Mohit, V., Linossier, I., Bourgougnon, N., Myrand, B., Olivier, F. (2012) Effect of bioﬁlm age on settlement of Mytilus edulis. Biofouling 28: 985–1001.

Townsin, R.L. (2003) The ship hull fouling penalty. Biofouling 19(Suppl.): 9 – 15.

Twigg, M.S., Tait, K., Williams, P. Atkinson, M., Camara M. (2014) Interference with the Germination and Growth of Ulva Zoospores by Quorum-Sensing Molecules from Ulva- Associated Epiphytic Bacteria. Environ Microbiol. 16: 445–53.

Vendeville, A., Winzer, K., Heurlier, K., Tang, C.M., Hardie, K.R. (2005) Making “sense” of metabolism: Autoinducer-2, LuxS and pathogenic bacteria. Nature Rev Microbiol 3: 383– 396.

Wagner-Döbler I, Thiel C, Eberl L, Allgaier L, Bodor A, Meyer S, Ebner S, Hennig S, Pukall R., Schulz S. (2005) Discovery of Complex Mixtures of Novel Long-Chain Quorum Sensing Signals in Free-Living and Host-Associated Marine Alphaproteobacteria. ChemBioChem 6 : 2195–2206.

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Wahl, M. (1989). Marine epibiosis. I. Fouling and antifouling: some basic aspects. Mar. Ecol. Prog. Ser. 58: 175–189.

Wahl, M., Shahnaz, L., Dobretsov, S., Saha, M., Symanowski, F., David, K., Lachnit, T., Vasel, M., and Weinberger, F. (2010) Ecology of antifouling resistance in the bladder wrack Fucus vesiculosus: patterns of microfouling and antimicrobial protection. Mar. Ecol. Prog. Ser. 411, 33–48.

Wahl, M., Goecke , L.A.,, Dobretsov, S., Weinberger. (2012) The second skin: ecological role of epibiotic bioﬁlms on marine organisms. Front Microbiol 3: 292.

Wang, C., Bao, W.Y., Gu, Z.Q., Li, Y.F., Liang, X., Ling, Y., Cai ,L.Y., Shen, H.D., Yang J.L. (2012) Larval settlement and metamorphosis of the mussel Mytilus coruscus in response to natural bioﬁlms. Biofouling 28:249–256.

Ward, J.P., King, J.R., Koerber, A.J., Croft, JM., Sockett, R.E., Williams P. (2003) Early development and quorum sensing in bacterial bioﬁlms. J Math Biol 47: 23–55.

Water, C.M., Bassler, BM. (2005) Quorum Sensing: Cell-to-Cell Communication in Bacteria.Annu Rev Cell Dev Biol 21: 319–46.

Webster, N.S., Negria, A.P. (2006) Site-specific variation in Antarctic marine biofilms established on artificial surfaces. Environ Microbiol 8:1177–1190.

Wheeler, G.L., Tait, K., Taylor, A., Brownlee, C., Joint I. (2006) Acyl-homoserine lactones modulate the settlement rate of zoospores of the marine alga Ulva intestinalis via a novel chemokinetic mechanism. Plant Cell Environ 29: 608–618.

Wieczorek, S.K., Clare, S., Todd, CD. (1995) Inhibitory and Facilitatory Effects of Microbial Films on Settlement of Balanus Amphitrite Amphitrite Larvae. Mar Eco Prog Ser 119: 221– 28.

Whiteley, M., Diggle, S.P., Greenberg, E.P. (2017) Progress in and promise of bacterial quorum sensing research. Nature 551:313–320.

Williams, Paul. (2007) Quorum Sensing, Communication and Cross-Kingdom Signalling in the Bacterial World. Microbiology 153: 3923–38.

Xavier, KB, Bassler BL. (2005) Interference with Al-2-mediated bacterial cell-cell communication. Nature 437: 750–753.

Xiong, Y., Liu, Y. (2010) Biological control of microbial attachment: a promising alternative for mitigating membrane biofouling. Appl Microbiol Biotechnol 86:825–837.

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Yang, J.L., Zhou, X., Li, Y.F., Guo, X.P., Liang, X., Li, J.L., (2014) Plantigrade settlement of the mussel Mytilus coruscus in response to natural bioﬁlms on different surfaces. J Mar Biol Assoc U. K. 94: 1639-1649.

Yang, Y., Wikieł, A.J., Dall’Agnol, L.T., Eloy, O., Genet, M., Moura, J.J.G., Sand, W, Dupont-Gillain, C.C., Rouxhet, P. (2016) Proteins dominate in the surface layers formed on materials exposed to extracellular polymeric substances from bacterial cultures. Biofouling 32: 95-108.

Yaniv, K., Golberg, K., Kramarsky-Winter, E., Marks, R., Pushkarev, A., Béjà, O. (2017) Functional marine metagenomic screening for anti-quorum sensing and anti-biofilm activity. Biofouling 33: 1–13.

Yebra, D.M., Kiil, S., Dam-Johansen, K. (2004) Antifouling technology – past, present and future steps towards efficient and environmentally friendly antifouling coatings. Prog Org Coat. 50:75–104.

Yeon, K-M, Lee C-H, Kim, J. (2009) Magnetic enzyme carrier for effective biofouling control in the membrane bioreactor based on enzymatic quorum quenching. Environ Sci Technol 43:7403–9.

Zang, T., Lee, B. W., Cannon, L. M., Ritter, K. A., Dai, S., Ren, D., et al. (2009). A naturally occurring brominated furanone covalently modifies and inactivates LuxS. Bioorg Med Chem. Lett 19: 6200–6204.

Zardus, J.D., Nedved, B.T., Huang, Y., Tran, C,, Hadfield, M.G. (2008) Microbial Biofilms Facilitate Adhesion in Biofouling Invertebrates. Biol Bull 214: 91–98.

Zargiel, K., Coogan, J.S., Swain, G.W.. (2011) Diatom Community Structure on Commercially Available Ship Hull Coatings. Biofouling 27: 955–65.

Zargiel, K.A., Swain, G.W.(2014) Static vs dynamic settlement and adhesion of diatoms to ship hull coatings. Biofouling 30:115 – 129.

Zimmer, B.L., May, A.L., Bhedi, C.D., Dearth, S.P., Prevatte, C.W, et al. (2014) Quorum sensing signal production and microbial interactions in a polymicrobial disease of corals and the coral surface mucopolysaccharide layer. PLOS ONE 9:e108541.

Zobell C.E., Allen E.C. (1935) The significance of marine bacteria in the fouling of submerged surfaces. J Bacteriol 29: 239-251.

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Chapter 3

Research paper: “Distinct profiles of early marine biofilms in two different seasons in a Portuguese Atlantic Port” (manuscript to be submitted)

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Distinct profiles of early marine biofilms in two different seasons in a Portuguese Atlantic Port

Jorge Antunes1,2, Joana Azevedo1,2, António Sousa1, Adriana Rego1,2, Pedro Leão1, Vitor Vasconcelos1,2

2Departamento de Biologia, Faculdade de Ciências, Universidade do Porto, Rua do Campo Alegre, P 4069-007 Porto, Portugal

Abstract

The formation of marine biofilms communities is the first stage of marine biofouling, involving a succession of prokaryotic and eukaryotic communities; however not much is known about their temporal or seasonal succession. In this work we used amplicon-based

NGS (Next-generation sequencing) to characterize early marine biofilms growing on steel surfaces in a Northern Portugal port. Sampling spanned 30-day periods in two distinct seasons (late spring and winter). Distinct temporal patterns were observed, namely a significantly higher abundance of Gammaproteobacteria and Mollicutes on the first spring days and a higher abundance of Alphaproteobacteria during the early winter sampling in the same period, while at intermediate biofilm days there was a stronger taxonomic similarity between the two sampled seasons. At the last sampled day (day 30), the spring biofilms significantly shifted towards a dominance of photoautotrophic groups (mostly diatoms), and predictive functional profiling using PICRUSt further suggested that those biofilms were already evolving towards mature biofilms, something not observed in the winter sampling.

Pathogenicity-associated taxa (particularly the Vibrio and Mycoplasma genera) were also detected, some of which have not previously been reported in marine biofilms, and their presence was strongly influenced by temporal variations. Furthermore, we detected the presence of taxonomic groups known to be associated with the corrosion of steel surfaces, whose relative abundance was mostly stable along the sampled days and months.

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Noteworthy, the presence of some of those taxonomic groups was influenced by the application of the tested anti-corrosion paint, while others seem to be resistant to it. Overall, we suggest that the herein reported temporal succession of early marine biofilm communities should be taken in consideration for future antifouling applications, as well for the effective control of pathogenic and steel-corroding associated bacterial taxa present in marine biofilms.

Keywords: marine biofilms; marine fouling; pioneer bacteria; steel surfaces; temporal succession

3.1 Introduction

Marine biofilms are surface-associated aggregates of microbes (including bacteria, fungi and protists) that rapidly colonize all submerged surfaces in marine ecosystems

(Antunes et al., 2018; Dang et al., 2016). Although the primary biofilm is dominated by bacterial species, especially after immersion for a few days, a significant accumulation of biomass is also attributed to diatoms (Railkin, 2004). Other group of microorganisms like microalgae, cyanobacteria, benthic dinoflagellates, fungi and protozoa follow the colonization after a few days, resulting in the formation of mature marine biofilms (Molino and Wetherbee, 2008). Despite several published studies in the last 10 years, the characterization of early marine biofilm communities in different environmental settings remains incomplete (Salta et al., 2013; Lawes et al., 2016; de Carvalho, 2018). The initial stages of marine biofilm formation, in particular, is not very well understood despite its crucial importance in the development of the biofilm and of the subsequent biofouling communities (Dang and Lovell, 2000; Dang et al., 2008). It is believed that biofilm and planktonic prokaryotic communities are distinct even from the first stages of biofilm

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formation, however microbial biofilms in aquatic environments are very heterogeneous and dynamic and are influenced by the surrounding environmental parameters (such as nutrient status, temperature, hydrodynamics and water chemistry) as well by the physicochemical properties of solid surfaces (Ward et al., 2003; Jones et al., 2007). The microbial communities that initially attach to surfaces are the decisive triggers of the marine biofouling processes (Salta et al., 2013) as marine biofilm constitutes preferential substrates for the colonization and settlement of larvae and spores of higher sessile organisms like macroinvertebrates (Dobretsov et al., 2009; Hadfield, 2011). Marine biofouling is known to cause serious problems to marine associated industries as well as to navies around the world and it is considered one of the most important issues facing marine technology today (Yebra et al., 2004; Schultz et al., 2011; Salta et al., 2013). Biofilms themselves have a significant role in the corrosion of steel surfaces immersed in seawater (Dang et al., 2011; Carvalho,

2018), are responsible for diminished hydrodynamic performance in ships (Holm et al., 2004) and can be vectors for the spread of pathogenic bacteria (Shikuma and Hadfield, 2010).

Certain bacterial taxa are routinely described as early colonizers of metal surfaces and are present in low-water corrosion (ALWC) structures (Jin et al., 2015; McBeth et al., 2011), a problem with a significant economic impact (Hou et al., 2017).

The ability to characterize biofilms community composition in environmental samples has increased substantially through the use of high-throughput next generation gene sequencing (NGS) techniques allowing for unprecedented analyses of the ocean microbiome

(Besemer et al., 2012). The spatial and temporal dynamics of bacterioplankton community are known to be driven by multiple environmental factors such as latitudinal gradient

(Fuhrman et al., 2008; Milici et al., 2016), temperature, as well biogeochemical and physical interactions, seasonal and geographical variations (Kim et al., 2016), Similarly, biofilm temporal evolution may have form a consistent time series, each being associated with a different bacterial community, but those patterns have not been thoroughly analyzed

(Stoodley et al., 2002). Previous studies have characterized the marine biofilm communities growing on different artificial substrata or on different geographical locations (Briand et al.,

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2012; Elifantz et al., 2013; Briand et al., 2017; Pollet et al., 2018). Overall, the current body of knowledge regarding bacterial communities of marine biofilms indicates that

Proteobacteria, and in particular the Alphaproteobacteria class dominate in these environments (Dang and Lovell, 2000; Huggett et al., 2006; Jones et al., 2007; Chung et al.,

2010). Roseobacter clade members (Alphaproteobacteria) seem to be the dominant group colonizing artificial surfaces immersed in temperate coastal waters in the Atlantic and the

Pacific Ocean (Dang et al., 2008). However, few studies have comprehensively analyzed the evolution of early biofilms and have compared their communities in different seasons or in geographic locations (Antunes et al., 2018). The impact of seasonality in the evolution of these pioneer biofilm microbial communities may be important to determine the correct application timeframe and the selection of anti-fouling or anti-corrosion paints, and to predict the timing of the settlement of the subsequent macrofouling communities.

Our main objective in this study was therefore to characterize the temporal succession of early marine biofilm communities over a one-month period growing on steel surfaces both during winter and during spring. We also aimed at characterizing the biofilm communities growing in surfaces covered with anti-corrosion paint, which would allow to determine the efficiency of those products and also to characterize the planktonic communities of the same location. We conducted our sampling on the Leixões Port, a major ship hub in Northern

Portugal and employed Next-generation sequencing (NGS) technology and universal prokaryotic primers. In addition to the metagenomic amplicon-based analysis, we also used a functional metagenome prediction analysis approach to infer probable functions performed by the different bacterial biofilm communities. We found that the early pioneer colonizing bacteria, their temporal succession, as well their functional profiles were strikingly different in the sampled seasons. This suggests that marine biofilms communities strongly respond to temporal shifts, and their diversity may depend on the sampled day of the season.

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3.2 Material and Methods

3.2.1. Study site and sampling

The Port of Leixões, North of Portugal (Matosinhos) was selected for the field assays

(41º10´39.32´´N, 8º42´8.78´´W). After some visits to the site, a pier in the northwest part of the Leixões Cruise Terminal marina was selected as the study site. This harbor was selected because submerged artificial surfaces in the location are known to be significantly covered with marine fouling and it is a port with a great traffic of cargo ships and cruisers. We employed 66 stainless steel foil plates (10 × 10 × 0.2 cm) of the 316L type, an austenitic chrome nickel steel with superior corrosion resistance typically used in surfaces exposed to chemical corrodents as well as submerged in marine environments. These plates have a chemical composition of a maximum of: Carbon: 0.08%; Manganese: 2.00%; Silicon: 0.75%;

Phosphorous: 0.045%; Sulfur: 0.03%; Chromium: 16-18%; Molybdenum: 2%-3% and

Nickel: 10%-14%. Three panels per season were professionally covered with Intersheen 579

(International Paint Ltd, Gateshead, UK), by the Port Authority of Douro, Leixões and Viana do Castelo (APDL). Intersheen 579 is a fast drying, single component, modified acrylic finish, non-biocidal anti-corrosion paint employed to avoid corrosion of metal surfaces.

Before being used in situ, the steel plates were pre-cleaned with HCl (0.1M) and autoclaved.

Afterwards, the plates were submerged vertically at a depth of approximately 1 m next to a pier deck in the Leixões cruise terminal and in two different season periods: during the spring of 2016 (24th of May-23rd of June 2016), and during the winter of 2017 (7th of February-6th of March 2017). The stainless-steel plates were attached by a kernmantle rope, supported by a grey PVC tube, vertically placed within a distance of 10 cm from each plate, in order to avoid contact between them, and positioned with a northeast/southwest orientation.

To conduct the sampling, an independent set of three steel plates were retrieved at days 1,2,4,7,10,14,21,25 and 30 of immersion in each of the sampled seasons, in order to obtain triplicate samples for each of the sampled days. The microbial biofilm biomass

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attached to the plates was scraped by using a sterilized razor blade and stored into sterilized microcentrifuge tubes. Approximately one liter of seawater at the same location was collected into an autoclaved polycarbonate bottle on days 1, 15 and 30 of plate immersion, for each season. Harvested biofilm samples and sampled seawater were transported in ice coolers for subsequent DNA extraction. Microorganisms present in the water sample were concentrated by using a filtration with 0.22 µm Sterivex filters (MOBIO, Carlsbad, CA, USA), and isolation of DNA from filtered water was conducted using PowerWater DNA Isolation Kit

(MOBIO, Carlsbad, CA, USA). Biofilms samples were preserved in LifeGuard Soil

Preservation Solution (MOBIO, Carlsbad, CA, USA), and kept at -80ºC until genomic DNA extraction was conducted. DNA from each biofilm sample was extracted and isolated using the using the Powersoil DNA Isolation Kit (MOBIO, Carlsbad, CA, USA) then stored at

−20ºC until sequencing. Environmental parameters of the seawater were measured at the test site with a multiparameter sensor from HQ40D (Hach Lange, Berlin, Germany) - over the experimental period.

3.2.2. DNA Sequencing and Sequence Analysis

The 16S rRNA gene sequencing was performed by LGC Genomics (XX, XX) using a 2× 300 bp paired-end read (Illumina MiSeq V3). To amplify 16S rRNA gene sequences, the prokaryotic universal primers 341 F (5’-CGC CCG CCG CGC CCC GCG CCC GTC

CCG CCG CCC CCG CCC GCC TAC GGG AGG CAG CAG-3’) and 785 R (5’-

CTACCAGGGTATCTAATCC-3’) were used (Klindworth et al., 2013). All processing and data analysis were performed using the Quantitative Insights Into Microbial Ecology software package (QIIME, v1.8.0) (Caporaso et al., 2010). In short, FASTQ files received were previously subjected to base calling and demultiplexing (Illumina bcl2fastq v.1.8.4).

Pairs of forward and reverse primer/adapter-trimmed sequences were overlap-combined

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using BBMerge 34.48. Reads with a final length of less than 100 bp were discarded, and the quality of paired-end Illumina reads was filtered based on quality (Phred) scores per- nucleotide using the quality parameters recommended by Bokulich et al., (2013) with a maximum unacceptable Phred quality score of Q20. The chimeric sequences were filtered out using UCHIME v4.2.40. Reads were assigned to OTUs using UCLUST (Edgar, R. C.,

2010), clustering at 97% sequence identity. The filtered reads were classified using the

Greengenes release gg 13 5 database (DeSantis et al., 2006), as implemented in the QIIME pipeline. Alpha-diversity indices (Observed OTU Number, Chao1, Shannon and Simpson) and beta-diversity were calculated using the Phyloseq package (McMurdie and Holmes,

2013), and the samples rarefaction curves were calculated with the vegan package (Oksanen et al., 2016), both in RStudio (RStudio Team, 2015). Variation of sample composition at each hierarchical level was examined using principal coordinate analysis (PCoA) conducted with both Bray-Curtis and weighted UniFrac distance matrices using QIIME. Analysis of similarity (ANOSIM) based on both Bray-Curtis and UniFrac distances was then performed to assess differences among groups within each hierarchical level using the same program.

One-way ANOVA was used to determine significant differences for each taxon for each of treatments, and Fisher’s unprotected LSD test was carried out to identify significant differences between treatments (GraphPad Prism v.5, GraphPad Software Inc., La Jolla, CA,

USA).

3.2.3 PICRUSt functional predictions

PICRUSt (Phylogenetic Investigation of Communities by Reconstruction of

Unobserved States) software (Langille et al., 2013) was used to predict the functional traits of the microbial communities associated with the different treatments based on 16S rRNA gene amplicon sequences. Firstly, a new OTU table was created with QIIME, (v1.8.0), using a closed-reference picking OTU protocol against the Greengenes database (version

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gg_13_5_otus) (Caporaso et al., 2010). The frequency of each OUT was then normalized by dividing the abundances of each OTU by known or predicted 16S rRNA gene copy number abundances. The normalized-OUT table was used to predict the functional genes, and the accuracy of the metagenome prediction was assessed by the nearest sequenced taxon index, using KEGG (Kyoto Encyclopedia of Genes and Genomes) Orthology (Kanehisa et al.,

2000), as the annotation reference database at different pathway levels (levels 1–3). To determine whether a certain functional family of gene was enriched in a sample or treatment, the ratios of each of the predicted genomic features were calculated accordingly. To further explore the functionality of the biofilm communities, specific predicted genomic features normally associated with biofilm functions were also analyzed and compared in the different samples and treatments.

3.2.4. Classification of chloroplast sequences

A separate OTU table was generated for OTUs classified as “chloroplast” by the

Greengenes classification in the last day of sampling for both seasons (day 30). Because the taxonomic assignments of chloroplasts via Greengenes was generally restricted to the

Phylum or Class level, we used the BLASTn NCBI classification as the primary way to more specifically classify those chloroplast sequences. Additionally, we classified the same chloroplast sequences by BLASTn search of the PhytoRef database (Decelle et al., 2015), a curated database with an access to plastidial 16S rRNA gene sequences of photosynthetic eukaryotes (downloaded in July 2018). Because the exact taxonomic identity often varied slightly between the two databases, we established rules for determining which classification to use following the protocol of Needham and Fuhrman et al., 2016. Accordingly, if a chloroplast sequence had a NCBI match was <95%, and the PhytoRef was >95%, we used the PhytoRef classification.

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3.3 Results

3.3.1. Environmental parameters

During the spring sampling (24th of May-23rd of June of 2016) water temperature

values ranged between a minimum of 16.1ºC and a maximum of 18.5 ºC, with an average

temperature of 17.1 ºC  0.9 ºC. Values of pH ranged between 7.09 to 7.67 with an average

pH of 7.37  0.20. For the winter sampling (7th February -6th March 2017) the temperature values varied between a minimum of 13.3 ºC to a maximum of 13.9 ºC with an average

temperature of 13.6 ºC  0.3 ºC. The pH values during winter varied between 7.64 and 7.94

with an average of 7.88  0.09. Regarding evolution of biofilm biomass in the sampled days, during spring there was an average increase of 87.6 ± 3.1 grams per bare plate, and an

increase of 80.2 ± 1.8 grams per plate covered with corrosion paint. During the winter there

was an average increase of 83.3 ± 2.1 grams per bare plate, and an average increase of 78.1

± 1.1 grams plate with anti-corrosion paint. Salinity values during our sampling varied

between 28.9 PSU and 33.3 PSU with an average of 31.3 PSU, while the dissolved oxygen

values were between 7.41 mg/L and 7.94 mg/L, with an average of 7.71 mg/L.

Table 1 - Environmental parameters of the seawater at the Leixões Port at each of the sampled days

Parameters Day 1 Day 2 Day 4 Day 7 Day 10 Day 14 Day 21 Day 25 Day 30 spring

pH 7.31 7.52 7.51 7.67 7.21 7.56 7.21 7.09 7.21

Temperature 16.1 16.5 16.6 17.5 17.5 18.2 18.6 16.3 16.5 (C) Parameters Day 1 Day 2 Day 4 Day 7 Day 10 Day 14 Day 21 Day 25 Day 30 winter

pH 7.65 7.87 7.91 7.94 7.92 7.91 7.85 7.92 7.91

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Temperature 13.3 13.6 13.6 13.3 14.0 13.5 13.9 13.9 13.5 (C)

3.3.2. Alpha-diversity and beta-diversity

Reads with a final length of less than 100 bases were discarded and the remaining

reads were primer clipped. Forward and reverse reads were combined to yield a complete

dataset comprised of 4 950 919 combined reads (154 784 reads or 3.03 % of the total reads

were uncombinable reads pairs, from 66 samples with an average 75 014 reads per sample

(minimum 762; maximum 453 850 reads). After quality filtering and chimera removal, a total

of 2 961 008 sequences were obtained across all samples (average 44 863; minimum 495;

maximum 269 867). All of the rarefaction curves for the winter samples reached a saturation

level, as well most of the spring samples, suggesting that the number of sequences per

samples was generally adequate to cover the whole diversity of the biofilm and planktonic

samples (Supplementary figure 1).

Regarding the alpha diversity, despite a higher average in estimates of OTU richness

(Chao1, Shannon and Simpson) in winter than in spring, no significant differences were

found between the combined samples of those two seasons regarding the diversity indexes

(Figure 1A). As for the effect of treatment, there was a significant lower diversity biofilm

growing in the plates when compared to the planktonic communities (for the observed OUT’s

and Chao1 indexes) (p-values <0.05). These differences, however, were only observed for

the spring sampling (Figure 1B). Regarding the effect of the day of sampling, there is an

average lower value of diversity in the second day of sampling with an increase of diversity

until day 25, followed by a decrease in the last day of sampling but without any statistically

significant differences between those days. There were however major differences in the

values of diversity for the sampled days within seasons: during spring there was a progressive

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increase of diversity from the first day of sampling until day 25 of sampling, followed by a decrease of diversity in the last day of sampling (day 30) (p-values < 0.01). During winter, despite the average increase of diversity with the daily succession of biofilms, there were no significant differences between the sampled days. Regarding the beta-diversity, the ANOSIM analysis demonstrated that the bacterial composition was affected more by the sampled day of the season (ANOSIM, R=0.34; p<0.01), but less so affected by the overall effect of the season (ANOSIM, R=0.10; p=0.1) and the effect of the day (ANOSIM, R=0.11; p=0.2).

Furthermore, we found a distinct cluster composed of most of the early spring biofilm samples (Figure 2A and 2 B), suggesting that these spring pioneer communities are distinct both from later spring bacterial communities and also from the almost totality of the winter samples.

Figure 1A: Alpha-diversity values of samples distributed by the two seasons

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Figure 1B: Alpha-diversity of samples per different treatment among the two

different seasons

Figure 2A and Figure 2B: Principal Coordinate Analysis (PCoA) of unweighted UniFrac distances of prokaryote 16S rRNA gene OTUs of the samples plotted by treatment (Figure 2A) and by season

(Figure 2B).

3.3.3. Taxonomic temporal succession

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Taking in consideration the total of the samples, the Prokaryotic communities were dominated by the phylum Proteobacteria (52.2  21.3%), followed by Bacteroidetes (15.6 

5.09%), Cyanobacteria (14.9  7.08%), and Actinobacteria (7.75  2.34%). Proteobacteria were strongly represented by Alphaproteobacteria (30.1  0.65 %), and

Gammaproteobacteria (25.0  5.6 %). Chloroplast (11.6  3.69) %, and Actinobacteria (7.17

 2.46%) were the subsequent most dominant classes in all the samples. Regarding the taxonomic rank of order, Rhodobacterales was the most abundant (15.7  4.60%), followed by Stramenopiles (11.1  4.31%), Flavobacteriales (8.73  4.02%), Burkholderiales (7.81 

3.02%) and Actinomycetales (7.79  2.63%). Rhodobacteraceae (22.1  2.25%) is by far the most common family in all of the samples, the next more common families were

Flavobacteriaceae (10.9  4.31%), Halomonodaceae (7.43  2.68%), and Alteromonodaceae

(5.36  3.90%). Considering the genera, the most abundant were Octadecabacter (4.17 

1.13%); Friedmanniella (4.04  2.15%); Clostridium (2.81  2.39%); Propionibacterium

(2.77  0.89%); Oleispira (1.97  2.37%) and Pseudoalteromonas (1.96  1.29%). Regarding combined differences between the tested seasons, no major differences were found for the abundance at the phyla level (Figures 3A and 3B), except that Actinobacteria and

Planctomycetes were significantly more abundant in the entire winter sampling than during spring sampling (p-values < 0.05). On the other hand, Tenericutes were significantly more common in spring (0.51  0.11%) than in winter (0.02  0.01%) (p-value < 0.001). The classes Chloroplast, Clostridia, Cytophagia and Mollicutes were also significantly more abundant in all samples collected during spring than during the winter (Figure 4A and 4B)

(p-values < 0.05). The same was verified for the orders Burkholderiales, Clostridiales and

Methylococcales (p-values < 0.01), and for the families Altermonodaceae and Clostridiaceae

(p-values < 0.01) (Figure 5A and 5B). On the other hand, during winter there was a higher abundance of the orders Actinomycetales and Deinococcales (p-values < 0.01), as well of the families Nocardioidaceae and Xenococcaceae (p-values < 0.01 and < 0.05, respectively). The genera Friedmanniella, Hymenobacter, Sulfurimonas and Proteus were more highly represented in spring than in winter, while in winter Candidatus Portiera, Clostridium,

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Mycoplasma, Oleispira, Pseudoalteromonas were more abundant. (all p-values < 0.05). The genera Friedmanniella, Hymenobacter, Sulfurimonas Proteus were more abundant in spring than in winter, while Candidatus Portiera, Clostridium, Oleispira, Pseudoalteromonas,

Proteus were more observed during winter than during in spring (all p-values <0.05).

Regarding the effect of the different treatments on the taxa, at the phylum level,

Acidobacteria and Actinobacteria are more abundant in seawater than in the sampled biofilms growing in surfaces with anticorrosion paint, while Tenericutes was much more abundant in steel surfaces with no anticorrosion paint than in the collected seawater, or in steel plates covered with anticorrosion paint (p-values < 0.001) (Figure 3A and 3B). Regarding the taxonomic level of class, Bacilli, Cytophagia and Epsilonproteobacteria are much more common in steel plates with no paint (p-values < 0.01), where there was also a significantly higher abundance of the orders Cytophagales, Flavobacteriales, Oceanospirillales,

Stramenopiles (Figure 4A and 4B). The orders Flavobacteriales, Methylococcales and

Stramenopiles, as well of the respective families Flavobacteriaceae and Vibrionaceae are more abundant growing in the steel surfaces than in the planktonic water suggesting a tendency to be generally more abundant in the biofilm stage. Additionally, the families

Clostridiaceae, Flavobacteriaceae, Phormidiaceae, Vibrionaceae are more abundant in steel surfaces with no paint. This is particularly true of the genus Mycoplasma (p-value < 0.001), as well of the genera Loktanella and Sulfurimonas (p-values < 0.01). On the other hand,

Alteromonadales (as well the Alteromonodaceae family), Deinoccocales (and the

Deinococcaceae family) and the family Xenococcaceae are much more common in the sampled planktonic communities than in the sampled biofilms either with or without paint

(p-values <0.01, except for Alteromonadales, p-value < 0.001). The same is verified for the genera, Candidatus Portiera, Chroococcidiopsis, Friedmanniella, Phormidium and

Sphingomonas which are significantly more abundant in the planktonic communities than in the sampled biofilms. Regarding the taxonomic differences per day of the combined seasons, no major differences were found for most taxa, suggesting the biofilm day of growth is generally not a major factor influencing taxonomic differences. At the phylum level the only

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exceptions are Acidobacteria which seems to be more common in early biofilm days

(between days 2 and 14) (Figure 3A and 3B). Other taxa abundance increases progressively while the biofilm matures, such as the classes Sphingomonodales, Rhodobacterales (and

Rhodobacteraceae family), Vibrionales (and Vibrionaceae family), as well the families

Alteromonodaceae, Flavobacteriaceae, Halomonadaceae and Clostridiaceae, however this increase was not always progressive, and some intermediary days may have a slight decrease of their abundance. The dominant Rhodobacteraceae family has a significant lower abundance in the first day of biofilm (0.72  0.68%) (p-value <0.001), starting to dominate in the second of sampling until reaching a maximum in the last day (15.3  7.86%). On the contrary, Alteromonodaceae, and Halomonodaceae see their dominance increase significantly in the last day of sampling (day 30) (p-values <0.001). When the different days of biofilm are compared in the two sampled seasons, more significant differences are found among the taxa. Firmicutes are more abundant in early days in spring than during winter, as well in the last days, in control treatment and in seawater. On the other hand, Proteobacteria and Cyanobacteria are more common since day 14 until the last day of sampling during spring, being much more common in those days in spring than during winter, particularly in the last day of sampling (p-values < 0.001) (Figures 3A and 3B). The dominant classes

Alphaproteobacteria and Flavobacteria are more abundant in the intermediate days of the spring biofilm, while other classes that dominate spring biofilms in the later days include

Chloroplast and Planctomycetia (p-values <0.01) (Figures 4A and 4B). Regarding the sequences classified as Chloroplast by the Greengenes database, the blast analysis of NCBI and phytoREF databases determined that at the last day of spring the phototrophic communities were dominated by the diatom Haslea sp. which no significant difference between the surfaces with anti-corrosion paint and the surfaces without paint. There was also a low abundance of the red algae Ahnfeltia (with no differences between treatments) and the green algae Desmochlorosis which was less abundant in the presence of the anti-corrosion paint (p-value <0.05). On the winter sampling, a dominance of the dinoflagellate

Thoracosphaera was observed, followed by the low abundance of the diatom Haslea sp.,

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with no significant difference among treatments for any of the most abundant taxa (Figure

6).

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Figure 5B: Taxonomic diversity and relative abundance at the order level of prokaryotes from the samples collected at winter (Figure 5B). Sequences were taxonomically assigned to the Greengenes database (DeSantis et al., 2006) using QIIME (Caporaso et al., 2010). For clarity of presentation, unassigned sequences and orders comprising <1% of the total number of sequences within a sample were simply classified as “Other”.

Figure 6: Taxonomic diversity and relative abundance of the Greengenes “chloroplast” sequences at the last day of sampling (day 30). Sequences were classified by the NCBI and

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PhytoREF databases at the lowest possible taxonomic ranks and chosen based on the best match. For clarity of presentation, unassigned sequences comprising <1% of the total number of sequences within a sample were simply classified as “Other”.

3.3.4. Functional analysis

For the functional profile analysis, no major differences were found for the main functional groups (L2) when the totality of samples for each season were compared (Figures

7 and 8). No difference was also found among treatments, except for the metabolism category which was less common in seawater than in sampled biofilms, while cellular processes abundance was higher in planktonic communities than in biofilms (p-value < 0.01).

Additionally, there was an apparent metabolism category enrichment in plates with anti- corrosion paint than in plates without paint (p-value < 0.001). Furthermore, cellular processes were more represented in seawater communities than in biofilms (p-value <0.05). Regarding differences per day for both seasons, there was a major decrease of metabolism for both seasons on day 7, followed by a significant increase in day 10 (p-values < 0.001). When the daily evolution per season was considered, genetic information processing was more prevalent during the first spring days (days 1 and 2), however this was not observed on winter

(Figures 7 and 8). Considering the L3 level analysis there was generally higher values for photosynthesis and photosynthetic proteins in spring than during winter (p-values < 0.05), while photosynthesis was lower in seawater than in the biofilm communities (p-value < 0.01)

(Figures 9 and 10). Bacterial chemotaxis and bacterial motility proteins were also more

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predominant in plates with paint than in planktonic communities. On the contrary, bacterial motility, sporulation, and glycan biosynthesis and metabolism were more associated to the later spring biofilm days (p-values < 0.05). Cytoskeleton proteins, peptidoglycan biosynthesis, and cell division were more represented during the last day of spring sampling, while flagellar assembly and bacterial chemotaxis were important components in the first days of sampling and less so in the last day of sampling (p-values <0.05). Those daily functional shifts were not observed, however for the winter sampling (Figures 9 and 10).

Figure 7: Relative abundance of 16S rRNA gene based predictive the main metagenomic functions of the spring samples, as assigned by KEGG category database (L2) through PICRUSt analysis.

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Figure 8: Relative abundance of 16S rRNA gene based predictive the main metagenomic functions of the winter samples, as assigned by KEGG category database (L2) through PICRUSt analysis.

Figure 9: Relative abundance of 16S rRNA gene based predictive metagenomic functions of biofilm associated functions in the spring samples as assigned by KEGG category database (L3) through PICRUSt analysis.

Figure 10: Relative abundance of16S rRNA gene based predictive metagenomic functions of biofilm associated functions in the winter samples, as assigned by KEGG category database (L3) through PICRUSt analysis.

3.4. Discussion

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3.4.1. Alpha- and Beta-diversity

Biofilms and planktonic communities are known to present dissimilar communities, with the latter often being associated with higher diversity values than the former (Jones et al., 2007; Lee et al., 2008; Shikuma and Hadfield, 2010). In our results we did not observe a major difference of the alpha-diversity indexes among the entirety of biofilm and planktonic samples. In our results, the Chao metric varied between 193.8  24.7 for the first day of spring and 4053.2  1237.2 for the fourth day of winter, the lowest and highest values, respectively.

We found an average Chao value for spring (1387.5 969.7) and for winter (2332.4 

1729.3), with the average of plates with no paint (1939.8 1501.6), and for the plates with anti-corrosion paint (1201.8 1241.0). Characterizing marine microbial community dynamics some authors (Gilbert et al., 2011; Cram et al., 2015) found distinct cyclic alpha- diversity patterns with a maximum in winter and a minimum in summer with seasonal differences tending to be greater than inter-annual changes (Furhman et al., 2015). We also found an average maximum of diversity in the winter samples which is contradictory with some previous biofilm studies which did not show clear diversity seasonal patterns (Patil and

Anil, 2015), but similar to a study which presented highest diversity during winter (Elifantz et al., 2013). During the spring sampling, the application of the tested anti-corrosion paint resulted in a lower diversity compared with biofilms growing without paint (p- value < 0.05).

This is in accordance with previous studies demonstrating that paints employed in surfaces can significantly influence prokaryotic diversity (Muthukrishnan et al., 2014; Briand et al.,

2017; Noble et al., 2016; Dussud et al., 2017). However, this effect was not observed during winter, which suggests that the effect of this paint in the prokaryotic diversity may be also dependent of seasonal variations. Compared to planktonic communities, surface-associated microorganisms normally possess compositional variations and dynamics responding to seasonal, daily or even hourly variations (Grossart, 2010; Ghiglione et al., 2007).

Suggestively, in our spring sampling, there was an average increase of alpha-diversity of all

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observed indexes (Observed OTUS, Chao, Shannon and InvSimpson) from day 1 until day

25 of biofilm growth, with a subsequent decrease of diversity. This may suggest that during spring there was a transition to mid-stage biofilms, which have moderate to low diversity when compared to early biofilms, and this relatively lower diversity may be explained by competitive species dominating the biofilms (Jackson et al., 2001; Noble et al., 2006). During winter, we observed a different pattern, with an increase of diversity until the fourth day of growth, followed by a non-continuous and slight decrease, and another increase the last days of sampling, indicating that there was still no stabilization of diversity. This confirms the notion that biofilm development and diversity are interconnected, and diversity is dependent on the testing time (Jackson et al., 2001), and our results seem to suggest they are also dependent on the season. Regarding the beta-diversity analysis we observed that the bacterial diversity (bacterial community composition) was more influenced by the seasonal day of biofilm growth, with a PCoA analysis clearly separating most early spring biofilm samples from all other samples (Figure 1). There were, however, no major differences between the seawater and biofilm communities, or the biofilm communities growing with paint or without paint, suggesting that there is no major influence of those treatments in the diversity of the communities.

3.4.2. Taxonomic temporal succession

At the phylum level, the most abundant group in our samples was Proteobacteria

(Figures 3A and 3B), confirming the existing body of work that points out that Proteobacteria, particularly the class Alphaproteobacteria dominate marine biofilms in different settings

(Dang and Lovell, 2000; Huggett et al., 2006; Jones et al., 2007; Chung et al., 2010).

Proteobacteria are pioneer surface colonizers, (Slightom and Buchan, 2009; Dang et al.,

2008, 2011) and are abundant across a range of different environmental conditions in biofilms

(Liu et al., 2015; Qi et al., 2016) and in seawater (Suh et al., 2015; Yang et al., 2015;

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Mancuso et al., 2016). Among the Alphaproteobacteria class, we observed a dominance of the order Rhodobacterales and of the Rhodobacteraceae family, similar to previous studies involving marine biofilms growing on steel surfaces (Emerson et al., 2010; Dang and Lovell,

2015). The Alphaproteobacteria class is known to survive well in poor environments (Alonso and Pernthaler, 2006), as is the case of initial phase of biofilm formation, when the amount of nutrients and organic carbon close to the surface is limiting (Flemming, 2002). The success of this taxonomic group is believed to be associated to their ability to react to low level of nutrient enrichment faster than other bacteria and are able to use a wide spectrum of carbon sources (Dang et al., 2008). We did not observe any major variation of either Proteobacteria or Alphaproteobacteria relative abundance as a response to temporal variations, confirming the dominance of this taxonomic group in the temporal evolution of biofilms (Figures 3A,

3B, 4A and 4B). Bacteroidetes were the second most common phylum in our samples, followed by Cyanobacteria. Within the phylum Bacteroidetes, the dominance of the order

Flavobacteriales was observed, which is in line with past studies sampling marine biofilm in temperate waters (Dang et al., 2011; Pollet et al., 2018). In contrast to Alphaproteobacteria,

Bacteroidetes requires higher levels of nutrients and organic matter (Kirchman, 2002;

Elifantz et al., 2013), which could be available as the biofilm matures and the matrix develops, and accordingly, we observed an increase of Bacteroidetes abundance in later biofilm days (starting on day 21). Bacteroidetes are a group well-known for colonizing steel surfaces in the marine environment and to be associated to biological corrosion (Kirchman,

2002; Dang and Lovell 2015). Members of this group are efficient surface colonizers and can take some advantage of primary production by chemolithotrophic Fe- and S-oxidizing bacteria during biofilm formation (Doghri et al., 2015).

In a recent study on microbial community succession over mild steel, the dominance of Proteobacteria and Bacteroidetes were described in different conditions, with the oxidizing iron group, the class Zetaproteobacteria within the Proteobacteria being the most abundant

(Rajala et al., 2015; McBeth and Emerson 2016). In our study however, the

Zetaproteobacteria class had a very low abundance (always lower than 0.1%).

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Burkholderiales (Betaproteobacteria) which are also known to oxidize iron (Rajala et al.,

2015) were common in our biofilm samples (1.05 0.20%) but were much less common in the planktonic communities (0.47 0.21%). Among the Deltaproteobacteria class, the order Desulfobacterales, an order particularly associated to corrosion of artificial surfaces

(Korenblumet al., 2010; Li et al., 2016; Tian et al., 2017) was also more detected in the biofilm growing in steel surfaces (with paint and without paint) but it was not detected in the seawater. This taxa like the Clostridia class, produce acetic, butyric, or formic acids so that their presence may also lead to corrosion of steel surfaces (Broda et al., 2000), and our results seem to suggest that these corroding associated taxa are particularly associated to the biofilm lifestyle. The Acetobacteraceae family (Alphaproteobaceria class, Rhodospirillales order), are known to produce acetic acid which can corrode steel (Paarup et al., 2006), and this family was less common in plates with anti-corrosion paint Intersheen 579, demonstrating a possible a role of this paint in controlling the growth of some steel corrosion associated taxa.

The Alteromonadales order (and the Alteromonadaceae family) were found to be significantly more common in the planktonic samples in our study, despite being also relatively common on the sampled biofilms. The orders Alteromonadales, Oceanospirillales,

Pseudomonadales and Vibrionales were previously described as the main taxa responsible for biocorrosion on carbon steel (Dang and Lovell, 2015; Xu et al., 2017), and in this work we observed that those taxa colonize steel surfaces since the very early days of surface colonization with no major effect of the anti-corrosion paint. Cyanobacteria are known to be dominant in biofilms reported from eutrophicated areas (Muthukrishnan et al., 2014; Sanli et al., 2015). Despite the presence of these taxa being in average higher in spring than in winter, probably as a consequence of higher temperature and higher solar irradiance, the differences were not significant. The Gram-positive bacteria in our samples were mainly represented by two phyla, Actinobacteria and Firmicutes, and the first of those groups was significantly more common in winter (Figures 3A and 3B). There was a relatively lower abundance of

Firmicutes compared to previous reports (Dang and Lovell, 2002), however their abundance was high on the first 10 days of winter biofilms (higher than 10%), while its abundance was

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lower than 1% in the same days of spring. Therefore, the abundance of this taxon seems therefore to experience a strong effect of seasonality. The particular importance of Firmicutes growth on steel surfaces is due to the fact that some members of this phylum generate H2S and organic acids which can result on corrosion (Paarup et al., 2006).

A growing number of human pathogens have been reported from marine environments (Ortega-Morales et al., 2006) and these microorganisms have been described as competitive, opportunistic and rapid colonizers, which can explain their recurring presence in young biofilms (Brinkmeyer, 2016). Vibrio sp. (Gammaproteobacteria) was a significant main component in carbon steel surfaces that had been immersed for one month in the

Cherbourg Harbour, France (Bermont- Bouis et al., 2007). In our samples, the presence of

Vibrio was higher in the winter sampling than in the spring sample (p-value < 0.05) which is surprising considering the optimum growth of this taxa is at higher temperatures but may explained by competition phenomena. Marine Vibrio species are also a major cause of wound infections in humans, which may be life-threatening (Oliver, 2005). Among the Firmicutes, the class Clostridia was the second most abundant taxa growing in our steel surfaces, a presence which may lead to corrosion (Muthukrishnan, 2014; Tian et al., 2017). In our results, the genus Clostridium was also more common in plates with no anti-corrosion covering and were particularly abundant in the early spring days (10.4%4.31% in the first day and 13.5% 2.31%in the second day). The genus Mycoplasma (Phylum Tenericutes, Class Mollicutes) was also particularly abundant in early spring biofilms (28.1% in the first day and 22.7% in the second day) being almost entirely absent in any of the winter sampling days, as well in spring sampling after day 4 (always lower than 0.1%). Mycoplasma are facultative anaerobes which are known to be associated with homeothermic animals and can also be human respiratory pathogens (Tully et al., 1993; Jacques et al., 2010). They are capable to grow attached to biotic or abiotic surfaces and to form prolific biofilms on glass and plastic surfaces in vivo studies conducted in the lab (Simmons and Dybvig, 2009).

Mycoplasma however were not described in previous in situ marine biofilm studies, and their relative high presence in our samples further demonstrates the role of marine biofilm

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communities as vectors of potential health hazardous taxa. Mycoplasma abundance is likely influence by environmental factors, with higher temperature likely playing a key role in influencing the abundance of these taxa in our data, as the genus has a preference for higher temperatures (Tully et al., 1993). In the generality of aquatic systems, temperature plays an important role in the changes of most biofilm propagation and metabolism (Bott, 1996; Rao,

2010). In general, temperature, salinity and pH are part of variables that are the most connected with bacterial diversity are believed to have the greatest influence on the biofilm formation and evolution (Donlan, 2002; Chiu et al., 2006; Briand et al., 2017). In our sampling we detected significant differences between the average temperature of the two sampled seasons, but not major variations of pH or salinity were observed (see Table 1), which may suggest that temperature is the main factor driving bacterial diversity patterns.

Questioning which group should be considered as the major and genuine pioneer bacterial group in marine biofilms, Lee et al., (2008) suggested that this role is performed by

Gammaproteobacteria, a conclusion supported partially by Pollet et al., 2018. In our results, however we observed important major seasonal changes of the early surface colonizers. In the first four days of spring, we also observed a strong presence of Gammaproteobacteria and of also Mollicutes. In winter, however Alphaproteobacteria was much more abundant in the early days, with a much lower presence of Gammaproteobacteria and a complete absence of the Mollicutes class (Figures 4A and 4B). This indicates that main pioneer substrate colonizing groups may vary substantially along the seasons of the year in the same location.

The overall dominance of the orders Rhodobacterales (Alphaproteobacteria) and

Flavobacteriales (Bacteroidetes), however did not seem to be affected by temporal variations

(Figures 5A and 5B) and corroborate the dominance of those taxa among marine biofilms sampled in different locations and sampling periods (Salta et al., 2013; Antunes et al., 2018).

Among the Gammaproteobacteria class, the Oceanospirillales order was found to be much more abundant in biofilms than in seawater, particularly during the first days of sampling, reaching a maximum in the first day of spring (18.8 5.40%), which is in accordance with previous reports suggesting this taxa are particularly abundant in early marine biofilms

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(Briand et al., 2017). The second most common family was Flavobacteriaceae, followed by

Halomonodaceae, Alteromonodaceae and Nocardioidaceae. This is in contrast with some previous studies which revealed a stronger dominance of other families like

Sphingomonadaceae (Chung et al., 2010; Briand et al., 2017) which was not verified in our sampling. In general, the observed seasonal variations suggest that early marine biofilm colonization may strongly dependent of seasonality, which was particularly evident in the pioneer bacterial communities sampled during spring which were particularly distinct from all other biofilm samples (See Figure 1A and 1B, and the ANOSIM analysis results).

However, after day 7 of sampling, biofilm communities in both seasons seem to converge towards a similar composition (Figures 4A, 4B, 5A and 5B). As the cells in the biofilms can closely interact with others and are protected by its exopolymeric matrix, the impact of seasonal variation of seawater parameters could be minimized after some days of growth, unlike what happens in planktonic communities (Salta et al., 2013; Dang and Lovell, 2016).

This is particularly true for mature biofilms, which probably can explain the stabilization of the relative abundance of the prokaryotic diversity in the mature biofilms sampled during the spring. Another important taxonomic shift was observed on the last day of spring sampling

(day 30) when there was a significant taxonomic shift towards the class Chloroplast

(eukaryotic autotrophs) (average of 70.2%), which was not reported during winter (average lower than 2%) (Figure 4A and 4B). These results suggest that seasonal changes in environmental variables may be more important than other interactions like competition or trophic interactions among biofilms (Pollet et al., 2018). When mentioning “seasonal” patterns however, it should be considered that there is an annual repeating pattern and not changes over a period of a few months (Furhman et al., 2015). Therefore, future biofilms studies should be collected over different seasons for several years to confirm if there is a real seasonal pattern as at least partially suggested by our results, and not a monthly or occasional taxonomic change.

Marine biofilm communities are selected from planktonic communities but appear to be dissimilar (Lee et al., 2008; Briand et al., 2012). However, it is unclear if the community

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structure of biofilms on artificial surfaces differs from free-living bacteria because artificial surfaces and free-living bacteria have not often been examined together. In our results, the majority of the planktonic bacteria were affiliated with Alphaproteobacteria, Bacteroidetes,

Cyanobacteria and Actinobacteria taxa, similar to previous results describing planktonic communities on temperate waters (Giovannoni and Stingl, 2005) (Figures 4A, 4B,5A and

5B). It has been proposed that biofilm and planktonic communities remain distinct even from early stages of biofilm formation but as the biofilm matures, its composition tends to become progressively more similar to the local microbial community (Ward et al., 2003; Pepe-

Ranney and Hall, 2015). We observed a similar taxonomic shift with a progressive increase of Proteobacteria abundance (particularly Gammaproteobacteria) with their relative abundance on later days becoming more similar to the seawater communities. The early spring biofilm communities were strikingly distinct from the planktonic communities, and in our planktonic communities we observed different temporal abundance of some taxa such as a higher abundance of Actinobacteria in winter compared to spring, while the class

Flavobacteria was much more abundant in the spring. Furthermore, Planctomycetes abundance was found to be higher during spring, suggesting its abundance is higher in warmer seasons, something which has been verified also in freshwater biofilms (Moss, 2006).

The domain Bacteria contribute the most to the initial surface microbiota, while the domain Archaea is rarely detected, but is known to be detected occasionally on marine biofilms sampled from steel surfaces (Dang et al., 2011; Webster et al., 2006). Archaea abundance in our results (0.2%  0.06%) is considerably lower than in those previous works with no major seasonal or daily variations. Similar to previous studies characterizing marine biofilms (Li et al., 2017; Pollet et al., 2018) there seems to be significant differences at the genus level as a response to the effect of seasonal, daily or treatment variations, with more significant differences at the genus level than at higher taxonomic ranks. Overall,

Gammaproteobacteria from the genus Oleibacter strongly dominated microbial communities during the first days of biofilm formation, something which had been previously been reported (Pollet et al., 2018), and we also observed an early strong presence of Clostridium

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(Firmicutes), Friedmanniella (Actinobacteria), Sphingomonas (Alphaproteobacteria) and

Sulfurimonas (Epsilonbacteria), which were the other main pioneer bacteria genera in our samples (particularly until day 4), and have not been commonly described in previous marine biofilm studies. On the other hand, later biofilms had a higher presence of Propionibacterium

(Actinobacteria), while Loktanella (Alphaproteobacteria), Octadecabacter

(Alphaproteobacteria) and Pseudoalteromonas (Gammaproteobacteria). All these last taxa seem to be the dominant primary surface-colonizing bacterial groups in different locations at relatively later biofilm sampling days (Dang et al., 2008; Dang and Lovell, 2016).

Concerning the biofilm microeukaryotes organisms, diatoms are key members being the main microphytobenthic group in marine epipelagic zone, especially in temperate areas

(Cassé and Swain 2006). Diatoms colonizing surfaces are believed to have a relative low diversity and only species from about 8–10 genera have commonly been documented to be problematic biofoulers of the ‘niche’ habitats provided by modern antifouling and fouling release coatings (Molino et al., 2008), most of them being pennate diatoms (Mitbavkar and

Anil 2008; Zargiel et al., 2011). Some past studies did not show any major differences on the colonization rates on the tested artificial surfaces both on temperate and tropical locations after two weeks of submersion (Molino et al., 2009) and diatom colonization of tested surfaces are believed to normally do not present clear temporal patterns (Elifantz et al.,

2013; Patil and Anil, 2005). In our results, we found a strong dominance of the motile pennate diatom Haslea sp. (over 80%) in the last day spring sampling (Figure 6), a genus which is known to colonize artificial surfaces (Winfield, 2018), while it had a much lower presence during winter (lower the 5%). Haslea sp. presence did not seem to be modulated by the application of the anti-corrosion paint which was not reported as a biocide. During winter, there was a high abundance of the phototrophic, calcareous dinoflagellate Thoracosphaera among all the chloroplast sequences, and a much lower abundance of diatom colonization when compared to spring (Figure 6). Low water temperatures are known to influence negatively the phototrophs community abundance, and diatom abundance is more common in warmer seasons, particularly as a part of spring or summer blooms (Molino et al., 2009),

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while Thoracosphaera is known to thrive at relatively lower water temperatures (Vink et al., 2003). Dinoflagellates species and other marine periphytic algae are known to colonize substrata as glass, with an average abundance of around 1% of the total of the covered surfaces (Railkin, 2004; Salta et al., 2013). The total abundance of that dinoflagellate in our results was around 2.5%, with no major influence from the anti-corrosion paint on its abundance. Thoracosphaera is known to be a common genus among Atlantic waters phytoplankton but has not been reported as a major colonizer of artificial surfaces, particularly steel surfaces. Dinoflagellate cysts have been recorded in ballast-tank sediments of ships arriving at ports around the globe (McCollin et al., 2000) and are a vector for their potential global dispersal (Hallegraeff, 1998), probably explaining the presence of this organisms in our sampling.

In general, there is a significant lack of information regarding the timing of the onset of the phototrophic colonization that precedes the macrofouling colonization of surfaces. On a study conducted near Melbourne, Australia, it was found that the diatom Cocconeis completely covered the areas of paint surface after 32 days of colonization (Molino et al.,

2009). In our results, we observed a major colonization of steel surfaces on spring by diatoms after a 30-day period (over 70% of total abundance of all the sequence reads), while their colonization was much lower during the winter (lower than 2%). This suggests a major difference in the temporal shift of biofilm colonization towards phototrophic communities in this Atlantic temperate water port. Furthermore, the appearance of the green algae

Desmochloris, and of the red macroalgae Ahnfeltia during the last day of spring sampling, also suggests these fouling communities growing on the panels are moving beyond the initial stages of microfouling and are entering the first stages of macrofouling, while the winter communities remained dominated by heterotrophic prokaryotes.

3.4.3. Functional analysis

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For the functional profile analysis, no major differences were found for most of the major groups (L2) suggesting that the functional profile of the biofilm samples displayed no major temporal variations (Figures 7 and 8). This suggests that there is an overall functional redundancy among biofilm samples in the different days which may provide a significant advantage of biofilm communities to changing environmental conditions along time, as the core functional traits are conserved (Allison and Martiny, 2008). There was however, a major difference for metabolic functions (Figures 7 and 8), with higher prevalence for biofilms growing in the steel surfaces when compared to the planktonic seawater communities, suggesting an increase of metabolic activity of microbial groups as soon as they attach to the surfaces in the early biofilm days. A higher metabolic activity in the surfaces covered with corrosion paint can be explained by an adaptive response of the prokaryotic communities to the compounds present in the anti-corrosion paint. This is also relevant as the level of metabolic functionality is believed to be correlated to the corrosion of surfaces (Videla and

Herrera, 2005; Rajala, et al., 2015). Interestingly, there was a predicted major decrease of metabolism in both seasons in the day 7 of biofilm growth, followed by a significant increase in day 10 (p-values <0.001), and this shift coincides with the important taxonomic shift during the same sampled day. In order to gain more insight into the functional repertoire of the samples, we compared the relative abundance of PICRUSRt predicted KOs (predicted

KEGG orthologs) at level-3 of functional groups associated with biofilm formation and development (Figure 9 and 10). Bacterial chemotaxis and bacterial motility proteins were more represented in plates with paint than both in planktonic communities and in plates without paint (p-values <0.05). The increase in abundance of genes encoding bacterial motility will likely influence the development of biofilms and may be associated an adaptation response to the anti-corrosive compounds of the paint with a further activation of genes associated with chemotaxis (Alon et al., 1999), and chemotaxis allows bacteria to swim towards optimum conditions, thus providing bacteria with a competitive advantage

(Wadhams and Armitage, 2004). Additionally, in topologically constrained environments such as surfaces, chemotaxis may be important for assembling a quorum of cells that can

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initiate biofilm development (Park et al., 2003). On the other hand, reduced bacterial motility, bacterial chemotaxis and flagellar assembly observed in the last spring biofilm days may be a result of the existence of an already well-developed biofilm (Lai et al., 2009), which was not verified in the winter sampling. Higher motility may be necessary to overcome the repulsive forces generated between cellular and abiotic surfaces and to allow favorable cell- surface interactions required for attachment (Geesey, 2001), which are required in the very first early biofilm days. Accordingly, motility genes are repressed after the bacterium attaches to the surface (Prigent-Combaret et al., 1999), and our analysis predicts both of those patterns in the last day of spring sampling. Furthermore, there was a significant increase of peptidoglycan biosynthesis functionality in the last spring day compared to all other days.

Peptidoglycan is major biofilm component and an integral part of the EPS (Weibel, 2011), enabling bacteria to form discrete multicellular communities (Varki, 2016). The predicted increase of the peptidoglycan biosynthesis in those samples was accompanied by an increase of cell division and of cytoskeleton evidencing a maturation of the biofilms (Figure 9). The increase in abundance of photosynthesis and photosynthesis proteins in the last spring biofilm day confirms a shift towards the dominance of photoautotrophic communities (Figures 9 and

10). Interestingly, in the plates with anti-corrosion paint there was an increase in abundance of sporulation functions, which may be related with the protection from deleterious environments (Kievit and Iglewski, 2000). Overall, we observed very distinct functional patterns in our samples, when both sampled seasons were compared, suggesting a strong functional and metabolic response of marine biofilm communities to temporal variation

3.5. Conclusions

Our study revealed distinct profiles among the sampled biofilms, suggesting that pioneer biofilm communities may face significant seasonal taxonomic and functional variations in the same location. These differences were particularly observed during the first

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four days of biofilm growth, after which there was a stabilization of diversity and a higher taxonomic similarity between the two seasons. Our results have also showed that in the last sampled day of the spring, biofilms shifted from heterotrophic towards phototrophic dominated communities, something which did not occur in the winter sampling. Furthermore, the conducted functional analysis also suggest that the spring biofilms were already converging and becoming mature biofilms, while the winter biofilms were still likely in the earlier stages of development. Overall, we confirmed the dominance of the Rhodobacterales order, as well of the Bacteroidetes phylum and the Flavobacteriales order among the totality of the samples, further proving the dominance of these taxa among marine biofilms. Among the pioneer colonizing groups, Alphaproteobacteria and Gammaproteobacteria were the dominant group depending on the sampled season. Some previously unreported taxa in marine biofilms were also detected; importantly, we detected a significant abundance of pathogenic associated taxa, and their presence was likely correlated to the higher temperatures detected on spring. These results further suggest the role of marine biofilms as vectors of harmful taxa, particularly in big ports and with access to effluents or to ship cargo waste. Additionally, taxa associated to the corrosion of the steel surfaces were also detected among the pioneer colonizing bacteria. Overall, our results should be taken in consideration for the efficient control of biofilm growth, particularly for the control of corrosion associated bacteria and of pathogenic bacteria. Future studies should be conducted in several and consecutive seasons to fully infer if marine biofilms taxonomic and functional shifts respond mostly to seasonal variations, or to more specific/broader temporal patterns.

Acknowledgments

Jorge Antunes was supported by the FCT grant SFRH/BD/99003/2013. This research was also supported by the projects NOVELMAR – Novel marine products with biotechnological applications (INNOVMAR NORTE-01-0145-FEDER-000035) and UID/Multi/04423/2019,

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through national funds provided by the Portuguese Foundation for Science and Technology

(FCT) and the European Regional Development Fund (ERDF), in the framework of the programme PT2020 and funding was provided by the project NASCEM PTDC/BTA-

BTA/31422/2017 (POCI-01-0145-FEDER-031422), financed by the FCT, COMPETE2020 and PORTUGAL2020. The work performed was done in collaboration with APDL

(Administração dos Portos do Douro, Leixões e Viana do Castelo, SA). We thank our colleagues from the CIIMAR animal facility (Bioterium), Hugo Santos, and Ricardo Branco, who helped in the construction and in the installation of the structure which supported the steel plates in the Leixões Port sampling. We also thank André Machado for assistance in the

BLASTn search and data analysis.

3.6 References

Alon, U., Surette, M.G., Barkai, N., Leibler, S.1999. Robustness in bacterial chemotaxis. Nature 397(6715):168–171.

Allison S.D., Martiny, J.B. 2008. Resistance, resilience, and redundancy in microbial communities. Proc Natl Acad Sci. 105:11512–11519

Alonso, C., Pernthaler, J,2006. Roseobacter and SAR11 dominate microbial glucose uptake in coastal North Sea waters. Environ. Microbiol. 8: 2022–2030

Antunes, J. Leão, P., Vasconcelos, V. 2018. Marine biofilms: Diversity of communities and of chemical cues. Environ. Microbiol, Rep. doi:10.1111/1758-2229.12694.

Bermont-Bouis D., Janvier, M., Grimont, P.A., Dupont, I., Vallaeys, T. 2007. Both sulfate- reducing bacteria and Enterobacteriaceae take part in marine biocorrosion of carbon steel. J. Appl. Microbiol. 102: 161–168 61.

Besemer, K., Peter, H., Logue, J. B., Langenheder, S., Lindström, E. S., Tranvik, L. J., et al. 2012. Unraveling assembly of stream biofilm communities. ISME J. 6: 1459–1468.

FCUP 101 Chapter 3

Bokulich, N. A., Subramanian, S., Faith, J. J., Gevers, D., Gordon, J. I., Knight, R., et al. 2013. Quality-filtering vastly improves diversity estimates from Illumina amplicon sequencing. Nat. Meth. 10: 57–59.

Briand, J.F., Djeridi, I., Jamet, D., Coupé, S., Bressy, C., Molmeret, M., Le Berre, B., Rimet, F., Bouchez, A., Blache, Y. 2012. Pioneer marine biofilms on artificial surfaces including antifouling coatings immersed in two contrasting French Mediterranean coast sites. Biofouling. 28:453–463.

Briand, J. F., Barani, A., Garnier, C., Réhel, K., Urvois, F., LePoupon, C., et al. 2017. Spatio- temporal variations of marine biofilm communities colonizing artificial substrata including antifouling coatings in contrasted French coastal environments. Microb. Ecol. 74: 585–598.

Brinkmeyer, R. 2016. Diversity of bacteria in ships ballast water as revealed by next generation DNA sequencing. Mar. Pollut. Bull. 107: 277–285.

Bott, T. L. 1996. Algae in microscopic food webs. In Algal Ecology: Freshwater Benthic Ecosystems, ed. R. J. Stevenson, M. L. Bothwell and R. L. Lowe, San Diego, CA: Academic Press, pp. 574–607.

Broda, D.M., Musgrave, D.R., Bell, R.G.2000. Use of restriction fragment length polymorphism analysis to differentiate strains of psychrophilic and psychrotrophic clostridia associated with ‘blown pack’ spoilage of vacuum-packed meats. J. Appl. Microbiol. 88:107– 116.

Campbell, M.L. 2009. An overview of risk assessment in a marine biosecurity context. Pages 353–372 in Rilov G, Crooks J, eds. Marine Bioinvasions: Ecology, Conservation, and Management Perspectives. Springer.

Caporaso, J. G., Kuczynski, J., Stombaugh, J., Bittinger, K., Bushman, F. D., Costello, E. K., et al. 2010. QIIME allows analysis of high-throughput community sequencing data. Nat. Meth. 7: 335–336.

Cassé, F, Swain GW. 2006. The development of microfouling on four commercial antifouling coatings under static and dynamic immersion. Int. Biodeterior. Biodegr. 57: 179– 185

Chiu, J.M.Y., Zhang, R., Wang, H., Thiyagarajan, V., Qian, P.Y. 2008. Nutrient effects on intertidal community: from bacteria to invertebrates. Mar. Ecol. Prog. Ser. 358: 41–50.

Chung, H.C., Lee, O.O., Huang, Y.L., Mok, S.Y., Kolter, R., Qian, P.Y., 2010. Bacterial community succession and chemical profiles of subtidal biofilms in relation to larval settlement of the polychaete Hydroides elegans. ISME J.4: 817–828.

FCUP 102 Chapter 3

De Carvalho, C.C.C.R. 2018. Marine biofilms: A successful microbial strategy with economic implications. Front. Mar. Sci. 5: 126.

Dang, H., and Lovell, C. 2000. Bacterial primary colonization and early succession on surfaces in marine waters as determined by amplified rRNA gene restriction analysis and sequence analysis of 16S rRNA genes. Appl. Environ. Microbiol. 66: 467–475.

Dang, H., Li, T., Chen, M., and Huang, G.2008. Cross- ocean distribution of Rhodobacterales bacteria as primary surface colonizers in temperate coastal marine waters. Appl. Environ. Microbiol. 74: 52–60.

Dang, H., Chen, R., Wang, L., Shao, S., Dai, L., Ye, Y. 2011 Molecular characterization of putative biocorroding microbiota with a novel niche detection of Epsilon- and Zetaproteobacteria in Pacific Ocean coastal seawaters. Environ. Microbiol. 13: 3059 – 74.

Dang, H., Lovell, C. R. 2016. Microbial surface colonization and biofilm development in marine environments. Microbiol. Mo. Bio. Rev. 80: 91–138.

Decelle, J., Romac, S., Stern, R.F., Bendif el, M., Zingone, A., Audic, S., et al. 2015. PhytoREF: a reference database of the plastidial 16S rRNA gene of photosynthetic eukaryotes with curated taxonomy. Mol. Ecol. Resour. 15: 1435–1445.

DeSantis, T.Z., Hugenholtz, P., Larsen, N., Rojas, M., Brodie, EL, Keller, K. et al. 2006. Greengenes, a chimera-checked 16S rRNA gene database and workbench compatible with ARB. Appl. Environ. Microbiol. 72: 5069–5072.

Dobretsov S, Dahms HU, Huang YL, Wahl M, Qian PY. 2007.The effect of quorum sensing blockers on formation of marine microbial communities and larval attachment. FEMS Microbiol Ecol. 60:177–188.

Dobretsov, S.; Teplitski, M.; Paul, C. 2009. Mini-Review: Quorum Sensing in the Marine Environment and Its Relationship to Biofouling. Biofouling 25: 413–27.

Dobretsov S, Abed RMM, Teplitski M. 2013. Mini-review: inhibition of biofouling by marine microorganisms. Biofouling 29:423–441.

Doghri, I., Rodrigues, S., Bazire, A., Dufour, A., Akbar, D., Sopena, V. et al. 2015. Marine bacteria from the French Atlantic coast displaying high forming-biofilm abilities and different biofilm 3D architectures. BMC Microbiol. 15: 231.

Donlan RM. 2002. Biofilms: microbial life on surfaces. Emerg Infect Dis. 8: 881–890.

FCUP 103 Chapter 3

Edgar, R.C. 2010. Search and clustering orders of magnitude faster than BLAST. Bioinformatics 26: 2460–2461.

Emerson, D., Fleming, E. J., and McBeth, J. M. 2010. Iron-oxidizing bacteria: an environmental and genomic perspective. Annu. Rev. Microbiol. 64: 561–583.

Elifantz, H., Horn, G., Ayon, M., Cohen, Y., Minz, D. 2013. Rhodobacteraceae are the key members of the microbial community of the initial biofilm formed in Eastern Mediterranean coastal seawater. FEMS Microbiol. Ecol. 85: 348–357.

Flemming, H.C., 2002. Biofouling in water systems - cases, causes and countermeasures. Applied Microbiology and Biotechnology 59 (6): 629-640.

Fuhrman, J. A., Steele, J. A., Hewson, I., Schwalbach, M. S., Brown, M. V., Green, J. L., et al. 2008. A latitudinal diversity gradient in planktonic marine bacteria. Proc. Natl. Acad. Sci. U.S.A. 105: 7774–7778.

Fuhrman, J. A., Cram, J. A., and Needham, D. M. 2015. Marine microbial community dynamics and their ecological interpretation. Nat. Rev. Micro. 13: 133–146.

Geesey, G. G., B. Wigglesworth-Cooksey, and K. E. Cooksey. 2000. Influence of calcium and other cations on surface adhesion of bacteria and diatoms: A review. Biofouling 15:195– 205.

Giovannoni, S. J., and Stingl, U. 2005. Molecular diversity and ecology of microbial plankton. Nature 437: 343–348.

Hadfield, M.G. 2011. Biofilms and marine invertebrate larvae: what bacteria produce that larvae use to choose settlement sites. Ann. Rev. Mar. Sci. 3: 453–470.

Hallegraeff, G. M. 1998. Transport of toxic dinoflagellates via ships' ballast water: Bioeconomic risk assessment and efficacy of possible ballast water management strategies. Mar. Ecol. Prog. Ser. 168: 297–309.

Hettich, R.L., Sharma, R., Chourey, K., Giannonem R.J. 2012. Microbial metaproteomics: identifying the repertoire of proteins that microorganisms use to compete and cooperate in complex environmental communities. Curr Opin Microbiol. 15: 373–380.

Holm, E. R., Schultz, M. P., Haslbeck, E. G., Talbott, W. J., Field, A. J. 2004. Evaluation of hydrodynamic drag on experimental fouling-release surfaces, using rotating disks. Biofouling 20: 219–226.

FCUP 104 Chapter 3

Huber, J.A., Mark Welch, D., Morrison, H.G., Huse, S.M., Neal, P.R., Butterfield, D.A., Sogin ML. 2007. Microbial population structures in the deep marine biosphere. Science 318: 97–100.

Huggett, M., Williamson, J., Nys, R, Kjelleberg, S, Steinberg, P. 2006. Larval settlement of the common Australian sea urchin Heliocidairs erythrogramma in response to bacteria from the surface of coralline algae. Oecologia 149:604–619.

Hou, B. R., Li, X. G., Ma, X. M., Du, C. W., Zhang, D. W., Zheng, M., et al. 2017. The cost of corrosion in China. npj Mater. Degrad. 1: 4.

Jacques, M., Aragon, V., Tremblay. Y.D.N. 2010. Biofilm formation in bacterial pathogens of veterinary importance. Anim. Health Res. Rev. 11: 97-121.

Jin, J., Wu, G., and Guan, Y. 2015. Effect of bacterial communities on the formation of cast iron corrosion tubercles in reclaimed water. Water Res. 71: 207–218.

Kanehisa, M., Susumu, G. 2000. KEGG: Kyoto encyclopedia of genes and genomes. Nucleic Acids Research. 28:27–30. de Kievit, T.R., and Iglewski, B.H. 2000. Bacterial quorum- sensing in pathogenic relationships. Infect Immun. 68: 4839–4849.

Kim, H. J., Jung, S. W., Lim, D.-I., Jang, M.-C., Lee, T.-K., Shin, K., et al. 2016. Effects of temperature and nutrients on changes in genetic diversity of bacterioplankton communities in a semi-closed bay, South Korea. Mar. Pollut. Bull. 106: 139–148.

Kirchman, D.L. 2002. The ecology of Cytophaga-Flavobacteria in aquatic environments. FEMS Microbiol. Ecol. 39: 91–100.

Korenblum, E., Valoni, É., Penna, M., Seldin, L. 2010. Bacterial diversity in water injection systems of Brazilian offshore oil platforms. Appl Microbiol Biotechnol. 85: 791– 800.

Jones, P., Cottrell, M., Kirchman, D., Dexter, S. 2007. Bacterial community structure of biofilms on artificial surfaces in an estuary. Microb. Ecol. 53: 153–162.

Lai, S., Trembaly, J., Déziel, E. 2009. Swarming motility: a multicellular behaviour conferring antimicrobial resistance. Environ. Microbiol. 11:126–136.

Langille, M.G.I., Zaneveld, J., Caporaso, J.G., McDonald, D., Knights, D., Reyes, J.A., et al. 2013. Predictive functional profiling of microbial communities using 16S rRNA marker gene sequences. Nat. Biotechnol. 31: 814–821.

FCUP 105 Chapter 3

Lawes, J.C., Neilan, B.A.., Brown, M.V., Clark, G.F, Johnston E.L. 2016. Elevated nutrients change bacterial community composition and connectivity: high throughput sequencing of young marine biofilms. Biofouling 32:1, 57-69.

Lee, J.W., Nam J.H, Kim, Y.H, Lee K.H, Lee D.H. 2008. Bacterial communities in the initial stage of marine biofilm formation on artificial surfaces. J Microbiol. 46: 174–182.

Li, B., Yang, Y., Ma, L., Ju, F., Guo, F., Tiedje, J. M., et al., 2015. Metagenomic and network analysis reveal wide distribution and co-occurrence of environmental antibiotic resistance genes. ISME J. 9(11):2490-502.

Li, X. X., Liu, J. F., Yao, F., Wu, W. L., Yang, S. Z., Mbadinga, S. M., et al., 2016. Dominance of desulfotignum, in sulfate-reducing community in high sulfate production- water of high temperature and corrosive petroleum reservoirs. Int. Biodeterior. Biodegradation 114: 45–56.

Li, X. X., Yang, T., Mbadinga, S. M., Liu, J. F., Yang, S. Z., Gu, J. D., et al., 2017. Responses of microbial community composition to temperature gradient and carbon steel corrosion in production water of petroleum reservoir. Front. Microbiol 8:2379.

Liu, J., Liu, X., Wang, M., Qiao, Y., Zheng, Y., Zhang, X.H., 2015. Bacterial and archaeal communities in sediments of the north Chinese marginal seas. Microb. Ecol. 70: 105–117.

Mancuso, F. P., D’Hondt, S., Willems, A., Airoldi, L., and Clerck, O. D. 2016. Diversity and temporal dynamics of the epiphytic bacterial communities associated with the canopy- forming seaweed cystoseira compressa(esper) gerloff and nizamuddin. Front. Microbiol 7:230.

Milici, M., Tomasch, J., Wos-Oxley, M. L., Wang, H., Jauregui, R., Camarinha- Silva, A., et al. 2016. Low diversity of planktonic bacteria in the tropical ocean. Sci. Rep. 6:19054.

Mitbavkar, S., Anil, A.C. 2008. Seasonal variations in the fouling diatom community structure from a monsoon influenced tropical estuary. Biofouling 24:415–426.

McBeth, J.M., Little, B.J., Ray, R.I., Farrar, K.M., Emerson, D. 2011. Neutrophilic iron- oxidizing ‘Zetaproteo- bacteria’ and mild steel corrosion in nearshore marine environments. Appl. Environ. Microbiol. 77: 1405–1412.

McBeth, J.M., Emerson, D. 2016. In Situ Microbial Community Succession on Mild Steel in Estuarine and Marine Environments: Exploring the Role of Iron-Oxidizing Bacteria. Front. Microbiol. 7:767.

FCUP 106 Chapter 3

McCollin, T.A., Hamer, J.P., Lucas, I.A.N. 2000. Transport of phytoplankton via ship’s ballast into ports around England and Wales. In: J. Pederson (Ed.) Marine bioinvasions: Proceedings of the First National Conference, Cambridge, MA: Massachusetts Institute of Technology, MIT Sea Grant College Program, pp. 282–288.

McDougald, D., Rice, S.A., Barraud, N., Steinberg, P.D., and Kjelleberg, S. 2011. Should we stay or should we go: mechanisms and ecological consequences for biofilm dispersal. Nat. Rev. Microbiol. 10: 39–50.

McMurdie, P. J., Holmes, S. 2013. Phyloseq: An R package for reproducible interactive analysis and graphics of microbiome census data. PLoS ONE 8: e61217.

Molino, P.J., Wetherbee, R. 2008. The biology of biofouling diatoms and their role in the development of microbial slimes. Biofouling 24: 365–379.

Moss, J., Nocker, A., Lepo, J.E., Snyder, R.A. 2006. Stability and Change in Estuarine Biofilm Bacterial Community Diversity. Appl. Environ. Microbiol. 72: 5679–88

Muthukrishnan, T., Abed, R.M.M., Dobretsov, S., Kidd, B., Finnie, A.A. 2014. Long-term microfouling on commercial biocidal fouling control coatings. Biofouling. 30:1155–1164.

Needham D.M., Fuhrman, J.A. 2016. Pronounced daily succession of phytoplankton, archaea and bacteria following a spring bloom. Nat. Microbiol. 1: 16005.

Oksanen, J., Blanchet, F. G., Kindt, R., Legendre, P., Minchin, P. R., O'Hara, R. B., et al. 2016. vegan: Community Ecology Package. R package version 2.3-4.

Oliver, J.D. 2005. The viable but nonculturable state in bacteria. J Microbiol. 43:93–125.

Ortega-Morales, B.O., Santiago-García, J., Chan- Bacab, M., Moppert, X., Miranda-Tello, E., Fardeau, M.L., Carrero,J., Bartolo-Pérez,P., Valadéz-González, A., Guezennec, J. 2007. Characterization of extracellular polymers synthesized by tropical intertidal biofilm bacteria, J. Appl. Microbiol. 102: 254–264.

Paarup, M., Friedrich, M. W., Tindall, B. J., and Finster, K. 2006. Characterization of the psychrotolerant acetogen strain syra5 and the emended description of the species acetobacterium carbinolicum. Antonie Van Leeuwenhoek 89: 55–69.

FCUP 107 Chapter 3

Park, S. Y., Lee, S. J., Oh, T. K., Oh, J. W., Koo, B. T., Yum, D. Y. & Lee, J. K. 2003. AhlD, an N-acylhomoserine lactonase in Arthrobacter sp., and predicted homologues in other bacteria. Microbiology 149: 1541–1550.

Patil, J.S., Anil, A.C. 2005. Biofilm diatom community structure: influence of temporal and substratum variability. Biofouling 21:189–206.

Pepe-Ranney C., Hall E. 2015, The effect of carbon subsidies on marine planktonic niche partitioning and recruitment during biofilm assembly. Front. Microbiol. 6:703.

Pollet, T., Berdjeb, L., Garnier, C. et al. 2018. Prokaryotic community successions and interactions in marine biofilms: the key role of Flavobacteriia. FEMS Microbiol. Ecol. 94: fiy083.

Prigent-Combaret, C., Prensier, G., Le Thi TT, Vidal, O., Lejeune, P., Dorel, C. 2000. Developmental pathway for biofilm formation in curli-producing Escherichia coli strains: role of flagella, curli and colonic acid. Environ Microbiol. 2: 450–464.

Renner, L.D., Weibel, D.B. 2011. Physicochemical regulation of biofilm formation, MRS Bull. 36: 347–355.

Qi, Y., Ying, W., Zhu, Z., Wang, X., Li, Z., and Jing, Z. 2016. Bacterial diversity in the surface sediments of the hypoxic zone near the Changjiang estuary and in the east china sea. Microbiology open 5: 323–339.

Railkin, AI. 2004. Marine biofouling – colonization, processes and defenses. Boca Raton, USA: CRC Press.

Rajala, P., Carpén, L., Vepsäläinen, M., Raulio, M., Sohlberg, E., and Bomberg, M. 2015. Microbially induced corrosion of carbon steel in deep ground water environment. Front. Microbiol. 6:647.

Rao, T.S. 2010. Comparative effect of temperature on biofilm formation in natural and modified marine environment. Aquat Ecol. 44:463–478.

Røder, H.L., Sørensen, S.J., Burmølle, M. 2016. Studying Bacterial Multispecies Biofilms: Where to Start? Trends Microbiol. 24:503-13.

Salta, M., Wharton, J.A., Blache, Y., Stokes, K.R., Briand, J.F. 2013. Marine bioﬁlms on artiﬁcial surfaces: structure and dynamics. Environ. Microbiol. 15:2879

FCUP 108 Chapter 3

Sanli, K., Bengtsson-Palme, J., Nilsson, R.H., Kristiansson, E., Rosenblad, M.A., Blanck, H., Eriksson, K. M. 2015. Metagenomic sequencing of marine periphyton: taxonomic and functional insights into biofilm communities. Frontiers Microbiol. 6:1192

Schultz, M.P., Bendick, J.A., Holm, E.R., Hertel, W.M. 2011. Economic impact of biofouling on a naval surface ship. Biofouling 27:87–98.

Shikuma, N.J., and Hadfield, M.G. .2010. Marine biofilms on submerged surfaces are a reservoir for Escherichia coli and Vibrio cholerae. Biofouling 26: 39–46.

Simmons, W.L. and Dybvig, K. 2009. Mycoplasma biofilms ex vivo and in vivo. FEMS Microbiology Letters 295: 77–81.

Slightom, R.N., Buchan, A., 2009. Surface colonization by marine Roseobacters: integrating genotype and phenotype. Appl. Environ. Microbiol. 75:6027–6037.

Stoodley, P., Cargo, R., Rupp, C. J., Wilson, S., Klapper, I. 2002. Biofilm mechanics and shear induced deformation and detachment. J. Ind. Microbiol. Biotechnol. 29: 361–368.

Suh, S. S., Park, M., Hwang, J., Kil, E. J., Jung, S. W., Lee, S., et al. 2015. Seasonal dynamics of marine microbial community in the south sea of Korea. PLOS ONE 10: e 0131633.

Tian, H., Gao, P., Chen, Z., Li, Y., Li, Y., Wang, Y., et al. 2017. Compositions and abundances of sulfate-reducing and sulfur-oxidizing microorganisms in water-flooded petroleum reservoirs with different temperatures in china. Front. Microbiol. 8:143.

Tully, J. G., Bové, J.M., Laigret, F., Whitcom, R.F. 1993. Revised taxonomy of the class Mollicutes: proposed elevation of a monophyletic cluster of arthropod-associated mollicutes to ordinal rank (Entomoplasmatales ord. nov.), with provision for familial rank to separate species with nonhelical morphology (Entomoplasmataceae fam. nov.) from helical species (Spiroplasmataceae), and emended descriptions of the order Mycoplasmatales, family Mycoplasmataceae. Int. J. Syst. Bacteriol. 43:378–385.

Varki, H.H., Freeze, P. Gagneux, Evolution of Glycan Diversity, in: A. Varki, R.D. Cummings, J.D. Esko, H.H. Freeze, P. Stanley, C.R. Bertozzi, G.W. Hart, M.E. Etzler (Eds.) Essentials of Glycobiology, Cold Spring Harbor (NY), 2009.

Videla, H.A., Herrera, L.K. 2005. Microbiologically influenced corrosion: looking to the future. Int. Microbiol. 8,169–180.

Vink, A., Baumann, K.-H., Bockel, B., Esper, O., Kinkel, H., Volbers, A., Willems, H., Zonneveld, K.A.F., 2003. Cocco- lithophorid and dinofagellate synecology in the South and Equatorial Atlantic; improving the paleoecological significance of phytoplanktonic

FCUP 109 Chapter 3

microfossils. In: Wefer, G., Mulitza, S., Ratmeyer, V. (Eds.), The South Atlantic in the Late Quaternary ^ reconstruction of material budgets and Cur- rent Systems. Springer, Berlin.

Wadhams, G. H. and Armitage, J. P. 2004. Making sense of it all: bacterial chemotaxis. Nat. Rev. Mol. Cell Biol.5:1024–1037.

Ward, J.P., J.R. King, A.J. Koerber, J.M. Croft, R.E. Sockett and P. Williams. 2003. Early development and quorum sensing in bacterial biofilms. Journal of Mathematical Biology 47: 23-55.

Webster, N.S., and Negri, A.P. 2006. Site-specific variation in Antarctic marine biofilms established on artificial surfaces. Environ. Microbiol. 8: 1177–1190.

Winfield, M.O., Downer, A., Longyear, J., Dale, M., Barker, G.L.A.2018. Comparative study of biofilm formation on biocidal antifouling and fouling-release coatings using next- generation DNA sequencing. Biofouling. 34(4): 464-477.

Xu, J., Tang, W., Ma, J., and Wang, H. 2017. Comparison of microbial community shifts in two parallel multi-step drinking water treatment processes. Appl. Microbiol. Biotechnol. 101: 5531–5541.

Zargiel KA, Coogan JS, Swain GW. 2011. Diatom community structure on commercially available ship hull coatings. Biofouling 27:955–965.

Zargiel KA, Swain GW. 2014. Static vs dynamic settlement and adhesion of diatoms to ship hull coatings. Biofouling 30:115–129.

Zhang, W. P., Wang, Y., Tian, R. M., Bougouffa, S., Yang, B., Cao, H. L., et al. 2014. Species sorting during biofilm assembly by artificial substrates deployed in a cold seep system. Sci. Rep. 4: 6647.

Yebra, D., Meseguer, S.K., and Kim, D. 2004. Antifouling Technology -Past, Present and Future Steps towards Efficient and Environmentally Friendly Antifouling Coatings. Progress in Organic Coatings 50: 75–104.

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Chapter 4

Research paper: “A multi-bioassay integrated approach to assess the antifouling potential of the cyanobacterial metabolites portoamides” (published in Marine Drugs)

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A Multi-Bioassay Integrated Approach to Assess the Antifouling Potential of the Cyanobacterial Metabolites Portoamides

Jorge Antunes 1,2, Sandra Pereira 1, Tiago Ribeiro 1, Jeffrey E. Plowman 3, Ancy Thomas 3, Stefan Clerens 3,4,5, Alexandre Campos 1, Vitor Vasconcelos 1,2, * and Joana R. Almeida 1, *

1 Centro Interdisciplinar de Investigação Marinha e Ambiental (CIIMAR), Universidade do Porto, Terminal de Cruzeiros do Porto de Leixões, Av. General Norton de Matos s/n 4450-208 Matosinhos, Portugal; [email protected] (J.A.), [email protected] (S.P.); [email protected] (T.R.); [email protected] (A.C.) 2 Departamento de Biologia, Faculdade de Ciências, Universidade do Porto, Rua do Campo Alegre, P 4069-007 Porto, Portugal 3 AgResearch Ltd., 1365 Springs Rd, Lincoln 7674, New Zealand; [email protected] (J.E.P.); [email protected] (A.T.); [email protected] (S.C.) 4 Biomolecular Interaction Centre, University of Canterbury, Christchurch P 8140, New Zealand 5 Riddet Institute, Massey University, Palmerston North P 4442, New Zealand * Correspondence: [email protected] (V.V.); [email protected] (J.R.A.)

Received: 28 December 2018; Accepted: 8 February 2019

Abstract: The cyclic peptides portoamides produced by the cyanobacterium Phormidium sp. LEGE 05292 were previously isolated and their ability to condition microcommunities by allelopathic effect was described. These interesting bioactive properties are, however, still underexplored as their biotechnological applications may be vast. This study aims to investigate the antifouling potential of portoamides, given that a challenge in the search for new environmentally friendly antifouling products is to find non-toxic natural alternatives with the ability to prevent colonization of different biofouling species, from bacteria to macroinvertebrates. A multi-bioassay approach was applied to assess portoamides antifouling properties, marine ecotoxicity and molecular mode of action. Results showed

high effectiveness in the prevention of mussel larvae settlement (EC50 = 3.16 µM), and also bioactivity towards growth and biofilm disruption of marine biofouling bacterial strains, while not showing toxicity towards both target and non-target species. Antifouling molecular targets in mussel larvae include energy metabolism modifications (failure in proton-transporting ATPases activity), structural alterations of the gills and protein and gene regulatory mechanisms. Overall, portoamides reveal a broad-spectrum bioactivity towards diverse biofouling species, including a non-toxic and reversible effect towards mussel larvae, showing potential to be incorporated as an active ingredient in antifouling coatings.

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Keywords: biofouling; antifouling; cyanobacteria; natural products; marine biofilms; mode of action

4.1 Introduction

As soon as a surface is immersed in water, it is rapidly colonized by a wide diversity of organisms in different developmental stages in a process known as marine biofouling [1,2]. Initially, the surface adhesion of organic molecules takes place, followed by the rapid settlement of microorganisms like marine bacteria, protozoans and microalgae [3]. These microfouling communities develop into complex marine biofilms, which provide favourable conditions in which spores of macroalgae and larvae of invertebrates can establish themselves, leading to the attachment of these macrofouling communities [4–6]. Marine biofouling is widely known to cause serious problems for marine industries worldwide and it is considered to be one of the most important issues facing marine technology today [7]. Marine fouling results in an increase of fuel consumption of ships and represents a major vehicle for the spread of invasive marine species [8,9]. Macrofouling organisms such as invertebrates and algae are responsible for a significant percentage of marine fouling biomass and for the diminished hydrodynamic performance of a ship [10]. However, marine biofilms that attach to artificial surfaces are also recognized to be a significant issue for a wide range of submerged engineered structures and are known to have a significant role in the corrosion of surfaces immersed in seawater [11]. The most commonly used strategy to prevent marine biofouling is based on antifouling (AF) paints containing active compounds [12]. However, due to their toxicity towards non- target organisms, some AF compounds have raised many environmental issues [13]. After years of application of toxic antifoulants to control biofouling, the International Maritime Organization (IMO) banned the use of organotin compounds in 2008, making urgent the development of environmentally friendly fouling-resistant coatings [14]. Therefore, large investments are applied yearly in the cleaning and the prevention of biofouling species settlement by using biocide-based antifoulants [15,16]. Marine organisms are a promising rich source of compounds for AF, antibiofilm and antipathogenic purposes, which despite increased research interest in recent years remain relatively underexplored [17,18]. Although

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there was intensive research in the last few years focusing on natural products showing broad and comprehensive AF properties, most of the discovered compounds have a very specific target and bioactivity towards only one or similar species of the marine fouling community [19–21]. In the context of natural products biodiscovery, cyanobacteria are a particularly encouraging source of new natural compounds, as they produce a wide range of secondary metabolites with recognized activity in several different biological responses [22,23], including effective and less toxic alternatives to combat marine biofouling, which might replace the biocide-based coatings currently in use [18]. However, being of natural origin does not guarantee that the AF compounds are non-toxic, and the ideal AF compound should be deterring rather than toxic to the target biofouling species [12]. As recently put forward by the Biocidal Products Regulation (BPR) of the European Union (528/2012), studies to elucidate AF mechanisms are considered to be a prerequisite for the registration and commercialization of any new antifoulant. It is thus of major importance to understand the molecular targets of newly discovered AF compounds because this will also help to identify the molecules or pathways involved in the settlement of biofouling organisms [21] and understand if alterations of these pathways are reversible after exposure. Furthermore, despite the importance of marine biofilms in early stages of biofouling formation [1,2], only a few natural compounds have been isolated with the capacity to disrupt marine biofilms [16,22]. Consequently, more chemical studies and broad-spectrum screening are needed to identify natural products that are active against different targets [23,24]. Portoamides are cyclic dodecapeptides previously isolated in our group from the cyanobacterium Phormidium sp. LEGE 05292 from the Blue Biotechnology and Ecotoxicology Culture Collection (LEGE CC, http://www.ciimar.up.pt/legecc/), based on their allelopathic effect on the microalga Chlorella vulgaris [25]. Portoamides A and B in natural mixtures also showed synergistic allelophatic activity towards other co-occuring microalgae (Ankistrodesmus falcatus and Chlamydomonas reinhardtii) and cyanobacteria (Cylindrospermopsis raciborskii) species [25]. Based on these previously recognized properties of inhibiting the growth of co-occurring microalgae and cyanobacterial species [26], the present work aims to investigate if portoamides are active against diverse marine biofouling organisms to be employed as non-toxic AF agents. The AF potential of portoamides was tested against both micro- and macrofouling species, specifically against relevant marine biofilm-forming bacteria and diatoms and also towards the adhesive larvae of the macrofouling mussel Mytilus galloprovincialis. In addition, the AF molecular targets of these metabolites were studied to determine their mode of action against Mytilus

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galloprovincialis larvae, and their potential marine ecotoxicity was also evaluated. In this study, this multi-bioassay approach will allow for determination of whether portoamides are effective as broad-spectrum and eco-friendly AF compounds and if they follow the new guidelines for the discovery of “clean” natural AF agents.

4.2 Results and Discussion

4.2.1 Mytilus galloprovincialis larvae Antisettlement Activity and Recovery

Portoamides demonstrated a highly significant activity towards the settlement of the mussel Mytilus galloprovincialis plantigrades larvae for all the concentrations tested (p < 0.01) when compared to the negative control, resulting in a complete inhibition of the larval settlement at a concentration of 32 µM, and an 50% response concentration (EC50 ) value of 3.16 µM/4.86 μg·mL−1 (Figure 1).

Figure 1. Dose–response anti-settlement activity of portoamides towards plantigrade larvae

of the mussel Mytilus galloprovincialis. B: DMSO control (0.01%); C: 5 µM CuSO4 as the positive control.

This anti-settlement bioactivity is highly relevant as it is well below the reference value −1 for EC50 (<25 μg·mL ) established by the U.S. Navy program for suitable AF compounds.

This EC50 value is lower than that of some previous AF compounds isolated from marine

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organisms [27–29], and their effectiveness is within the concentration range of synthetic nature inspired AF compounds tested in the same experimental conditions using M. galloprovincialis plantigrades antisettlement responses [30,31]. Considering pure compounds isolated from cyanobacterial strains and tested for antisettlement activity, −1 portoamides are more effective than hantupeptin C (EC50 = 10.6 μg·mL ), but less potent −1 −1 than isomalyngamide A (EC50 = 2.6 μg·mL ), majusculamide A (EC50 = 0.54 μg·mL ), and −1 dolastatin 16 (EC50 = 0.003 μg·mL ), all isolated from the cyanobacterium Lyngbya majuscula [32]. However, these compounds were only tested against Balanus amphitrite cyprids and larval sensitivity among invertebrate species might be distinct. Additionally, dolastatin 16 is very potent against cyprids settlement, but also toxic at low concentrations −1 (LC50 = 20 μg·mL ). The same was found for the compound Maculalactone A isolated from the cyanobacterium Kyrtuthrix maculans, which showed AF potential in field tests, but was −1 found to exert toxicity (LC50 = 4.2 μg·mL ) to B. amphitrite larvae [33]. The portoamides level of effectiveness towards M. galloprovincialis larvae is also higher than that of the ® commercial agent ECONEA , which has a reported EC50 value of 4.01 µM [30]. Despite the high effectiveness in preventing M. galloprovincialis plantigrades settlement, portoamides caused no mortality to the target species at any of the concentrations tested, while the ® −1 commercial AF agent ECONEA was found to exert some toxicity (LC50 = 107.89 μM·mL ) [31]. The lack of toxicity of portoamides towards mussel larvae at the tested concentrations suggests that portoamides have a deterring effect rather than a toxic mechanism towards the mussel larvae. In addition, the settlement recovery bioassay pointed out that after a recovery period of 15 h in fresh seawater, larval settlement responses increased by 35% and 45%, when compared to the responses immediately after portoamides exposure (3 µM and 16 µM, respectively), being also not significantly different from the negative control (Figure 2). This might indicate that the mechanisms involved in larval antisettlement are reversible at least after acute exposure to portoamides.

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Figure 2. Settlement response of plantigrade larvae of the mussel Mytilus galloprovincialis after 15 h of exposure to portoamides followed by 15 h of recovery in filtered seawater. B:

DMSO control (0.01%); C: 5 µM CuSO4 as the positive control.

This implies that portoamides have a potential to be employed as environmentally friendly AF agents against mussel attachment prevention, which is environmentally relevant, as Mytilus sp. are one of most prevalent macrofouling organisms worldwide, representing a significant part of the biofouling biomass [34,35].

4.2.2. Antibacterial and Antimicroalgal Activities

The capacity of portoamides to interfere with microfouling growth was assessed by screening portoamides against a panel of marine bacteria and microalgae species involved in marine biofilm formation. In total, portoamide bioactivity was tested against five marine bacteria and four microalgae strains. The results showed significant activity of portoamides against three of the tested marine bacteria, Cobetia marina (inhibition of 23.3%), Halomonas aquamarina (inhibition of 21.0%) and Pseudoalteromonas atlantica (inhibition of 21.5%) at a concentration of 6.5 µM, but no significant inhibitory activity against Vibrio harveyi or Roseobacter litoralis was detected. In the dose–response assay, portoamides displayed an inhibitory activity of 33.8% against C. marina, 37.1% against H. aquamarina and 38.9% against P. atlantica at the highest tested concentration of 26 µM (Figure 3).

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In this study, the reported EC30 value of portoamides against Halomonas aquamarina (Table 1) was similar to the activity of sphaerocyclamide, a cyanobactin isolated from a Sphaerospermopsis sp. strain [36], and also in line with the activity of nature inspired sulfated compounds [30], which were tested against the same bacterial strain.

Table 1. Bacterial growth inhibition activitiy of portoamides.

Compounds EC30 (M) Halomonas Pseudoalteromonas Cobetia marina aquamarina atlantica Portoamides 14.56 14.83 13.59 (95% CI: 10.21– (95% CI: 10.41– (95% CI: 9.67–21.45) 23.75) 24.66)

EC30, minimum concentration that inhibited 30% of bacterial growth. CI, confidence interval.

Interestingly, the EC30 value of portoamides is lower than the EC30 value of the commercial product ECONEA® (EC30 = 15.31 µM) [31]. Portoamides bioactivity is also similar or stronger than other natural AF compounds tested against other H. aquamarina strains [37,38]. Although the activity of portoamides against C. marina is lower when compared to some reported natural AF compounds or crudes [39,40], portoamides were also more efficient in inhibiting the growth of C. marina than ECONEA®. Portoamides activity against P. atlantica was significanly higher than that of other natural products [35], some of

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which such as eleganediol and eleganolone had EC50 values in the order of 300 to 400 µM

[41], while the portoamides-extrapolated EC50 value is 55.43 µM. Portoamides have already shown a significant inhibitory activity against the microalgae Chlorella vulgaris (IC50 =12.8 −1 µg·mL ) [25], towards the co-occurring microalga Ankistrodesmus falcatus (IC50 = 24.7 −1 −1 µg·mL ) and Chlamydomonas reinhardtii (IC50 = 12.6 µg·mL ), and were also bioactive −1 against a strain of the cyanobacterium Cylindrospermopsis raciborskii (IC50 = 28.4 µg·mL ) species [25]. Those previous results have suggested that portoamides are strongly bioactive secondary metabolites and their underexploited potential was here confirmed by their inhibitory activity towards the growth of several biofilm-forming bacterial strains. No significant inhibitory activity of portoamides was found against diatom growth, except for a slight inhibition of Halomphora sp. growth (<20%) at a concentration of 6.5 µM (Figure 4).

Figure 4. Antimicroalgal activity of portoamides at a concentration of 6.5 µM towards four biofilm-forming marine diatoms Cylindrotheca sp., Halamphora sp., Nitzschia sp. and Navicula sp. (B: 0.1% DMSO); 3.55 µM cycloheximide was used as the positive control (C+).

Diatoms represent a class of particularly relevant microorganisms in biofouling, which attach to artificial surfaces after the bacterial colonization, forming resilient slimy layers on marine surfaces, and are known to be a difficult group to target with any AF strategy [42]. The low inhibitory effect towards the tested microalgae species suggests that the mechanism responsible for the bactericidal activity of the portoamides does not affect these eukaryotic microorganisms.

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4.2.3. Antibiofilm Activity

The antibiofilm screening assay showed a higher activity of portoamides against the same bacterial strains in bacterial growth. Portoamides disrupted the biofilm cells of C. marina (10.2% of inhibition), H. aquamarina (25.1% of inhibition) and P. atlantica (44.5% of inhibition), but no significant inhibitory activity was found against either V. harveyi and R. litoralis biofilms at the tested concentration of 6.5 μM. The bacterial strains for which the most significant biofilm disruption was observed were selected to be further tested at higher and lower concentrations of portoamides (52, 26, 13, 6.5, 3.25,1.63, 0.81 and 0.41 μM). The concentration–response assay displayed a strong antibiofilm activity of portoamides against P. atlantica from the lowest concentration of 0.41 μM (11.3% of growth inhibition) to the highest concentration of 52 μM (71.5% of growth inhibition) (Figure 5), when compared to the negative control (EC50 = 13.54 µM; EC30 = 3.57 µM).

Figure 5. Antibiofilm dose–response activity of portoamides towards the marine bacteria Halomonas aquamarina and Pseudoalteromonas atlantica. B: 0.1% DMSO; C: 4:100 dilution of penicillin-streptomycin-neomycin stabilized solution (Sigma P4083).

Similarly, a significant inhibitory activity of portoamides was observed toward H. aquamarina biofilms from the lowest concentration of 0.41 μM (10.9% of growth inhibition) to the highest concentration of 52 μM (41.0% of growth inhibition) (Figure 5), when compared to the negative control (EC30 = 17.93 µM), but no relevant disruption of C. marina biofilm was found (<10% of inhibition). Taking into consideration this antibiofilm activity

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of portoamides and their inhibitory activity against bacterial growth, it can be hypothesized that these natural compounds have a broad activity against those biofilm bacterial strains at different stages of their life cycle, both at the planktonic stage and at the biofilm stage. The Halomonas genus and Halomonas aquamarina species, in particular, are one of the most common microorganisms constituting marine biofilms, and are particularly prevalent on biofilms attached to the hulls of ships [43,44]. Pseudoalteromonas is among the most preponderant marine bacterial genera recovered from different marine biofilm situations [45−48]. Compounds isolated from a Mediterranean brown alga were found to be disruptive of a Pseudoalteromonas strain D41, and among the tested compounds, the one with highest activity (dictyol C) had a reported EC50 value of 30 μM [41], which is significantly higher than the EC50 obtained for portoamides (EC50 = 13.54 μM). The activity of these compounds refers to the detachment of already adhered bacteria and not to the inhibition of planktonic growth. It is known that inhibitors of biofilm formation can serve as potentially efficient antifoulants, and thus biofilm inhibition and quorum-sensing (QS) inhibition bioassays are often used to assess the AF properties of natural compounds [20]. The Calgary Biofilm Device [49] was used to determine the antibiofilm activity of portoamides, and the Minimal Biofilm Eradication Assay (MBEC) was applied to quantify biofilm eradication as a response of compounds bioactivity [49]. Specifically, with the MBEC assay, it is possible to determine both the minimum biofilm eradication concentration (MBEC), simultaneously with the determination of the minimal inhibitory concentration (MIC) of the planktonic growth for a tested compound. Previous studies which used the MBEC assay reported the disruption of pre-formed biofilms by natural compounds; however, those have focused mostly on pathogenic biofilm strains and not on marine biofilms [50,51]. Other studies have analyzed the response of marine bacteria attachment to surfaces in the presence of natural compounds but have rarely focused on the disruption or on the detachment of already attached biofilms [52], which has importance for AF purposes. Natural compounds, which can disrupt biofilm formation, may be useful for the biotechnological development of an environmentally friendly protection against marine fouling [53]. There has been a lack of studies quantifying the disruption of marine biofilms, but those assays are essential for future AF studies involving new natural compounds as biofilms constitute the initial stage of marine biofouling [21]. So far, only a few natural products derived from marine microorganisms with antibiofilm activity have been reported [20], and most compounds with antimicrofouling activity have been developed to target

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microfouling species in their planktonic stage [16]. However, marine biofilm formation allows bacteria to be more resistant to environmental pressures and are the initial stage of the marine biofouling process [2,3]. Therefore, it is promising to exploit natural compounds with excellent antibiofilm activity as potential antifoulants. Portoamides seem to be particularly effective natural compounds in the eradication of biofilms of marine bacterial strains involved in biofouling and surface corrosion. Portoamides also show higher activity than some previously discovered natural products or some AF commercial biocides. This potential should be confirmed in future studies, namely by testing portoamides against other marine biofilm-forming bacterial strains involved in marine fouling situations.

4.2.4. Quorum-Sensing Inhibition (QSI) Activity

The standard QS disk-diffusion assay with Chromobacterium violaceum was used to determine if portoamides interfered with the QS phenomenon while not interfering with bacterial growth which is an ideal strategy to avoid biofilm formation [2,54]. The QS inhibition screening showed that portoamides at 6.5 µM concentration produced a transparent halo in the Chromobacterium violaceum agar cultures, which is associated with the cellular death of the QS C. violaceum reporter strain. The subsequent assay that quantified the inhibition of growth of the reporter strain, when compared to its production of violacein, further demonstrated a significant inhibition of C. violaceum growth by portoamides (31% of growth inhibition at a concentration of 6.5 µM, and 42% of growth inhibition at a concentration of 13 µM; data not shown) with no significant inhibition violacein production (Figure 6).

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m n

0 2

O /

m 1

7 D

O 0 B C 13 6.5 3.25 Portoamides concentration (mM)

Figure 6. Ratio of the quantification of growth inhibition of Chromobacterium violaceum ATCC 12472 (read at 720 nm) when compared to quantification of the production of violacein by the same bacterium (577 nm). B: 0.1% DMSO; C: 1:100 dilution of penicilin- streptomycin-neomycin stabilized solution (Sigma P4083).

Portoamides inhibitory activity against C. violaceum (a Gram-negative bacteria) further demonstrates the bactericidal/inhibitory activity of these cyanobacterial natural compounds, while not displaying any detectable QS inhibitory activity. Therefore, portoamides do not appear to have potential as quorum-quenching compounds as they were not shown to interfere with AHL-mediated cellular communication in the conducted assays.

4.2.5. Ecotoxicity Assessment

To evaluate the viability of portoamides as AF agents, its potential ecotoxicity was assessed using the standard Artemia salina bioassay. No significant differences in the Artemia salina nauplii mortality rate was found for portoamides at both the tested concentrations (16 µM and 3 µM), when compared with the negative control (p < 0.01) (Figure 7). The absence of toxicity of portoamides against this model toxicity organism [55] suggests the potential of portoamides as suitable AF agents with no expressive hazardous effect on the marine environment. However, the toxicity of portoamides towards organisms of other trophic levels should also be tested to fully interpret the absence of toxicity of these compounds.

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Figure 7. Mortality rate of Artemia salina nauplii after 48 h of exposure to portoamides. B:

1% DMSO in filtered seawater. C: K2Cr2O7 at a concentration of 13.6 µM.

4.2.6. Mode of Action Assessment

The assessment of AF molecular targets of AF candidates represents an important complementary information in the establishment of new AF agents. The challenge for the selection of promising AF candidates should include a mode of action elucidation in target biofouling species permitting the explaination of a broad-spectrum effect or a specific adhesive effect of non-toxic compounds that do not act as biocidal agents. To reach this, the differential expression of proteins in the proteome of M. galloprovincialis plantigrade larvae in response to the portoamides when compared to control larvae was analyzed by label-free shotgun proteomics. The complete information regarding differential protein abundance and the molecular functions of identified proteins is summarized in Table 2. Data correspond to the expression of proteins from ten M. galloprovincialis whole plantigrade larvae from the antisettlement bioassays per replicate, and the mean of the four replicates was considered. Portoamides exposure caused the alteration of several proteins, being the most representative proteins attributed to energetic metabolism processes and structural characteristics (Table 2). The most relevant under-expressed proteins participate in ATP synthesis processes (proton- transporting ATPase activity), indicating an inhibition of ATPases activity, which may result in increasing mitochondrial proton gradient and a decline in ATP levels. This effect was previously described in the colon carcinoma cell line HT-29, after exposure to portoamides and attributed as a molecular mode of action [56].

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Table 2. Differentially expressed proteins (p-value < 0.05) in Mytilus galloprovincialis larvae exposed to portoamides (16 M). Besides the mean log10- transformed abundance values, the names of the identified proteins and the main metabolic processes involved are also listed in the table. Protein accession numbers are from NCBI.

Mean of log10 (x + 1) - Transformed Values p-value Protein Name Metabolic Process Portoamide Control s ATP synthase subunit d, 2.465 0.304 0.004 mitochondrial ATP synthase subunit beta 3.742 3.335 0.011 mitochondrial ATP synthesis H+ ATPase a subunit 3.783 3.494 0.012 Energy mitochondrial Metabolism ATP synthase subunit 3.424 3.059 0.018 gamma 3.455 2.890 0.018 Isocitrate dehydrogenase 3.401 3.023 0.027 Malate dehydrogenase Tricarboxylic Acid cycle Glutamate dehydrogenase 2.825 2.405 0.031 mitochondrial Myosin heavy chain 4.116 3.771 0.050 striated muscle Muscle structural protein; Pedal retractor muscle contractionmovement 3.855 3.469 0.046 myosin heavy chain 4.386 3.882 0.050 Tubulin beta chain 3.857 3.379 0.038 Tubulin alpha-1A chain Cilia- and flagella- 3.166 2.623 0.035 associated protein Cytoskeleton constituent 3.756 3.064 0.021 Myosin heavy chain Structural 3.149 2.888 0.018 Tubulin beta chain 3.497 2.929 0.015 Tubulin beta chain 3.269 2.819 0.004 Myosin heavy chain 2.660 1.971 0.024 Tektin-4 Radial spoke head protein 2.440 1.054 0.027 4 Structural, cell motility Radial spoke head protein 2.343 1.106 0.031 9 3.31471 2.727 0.031 Tektin-3 1.974 0 0.037 Collagen, type VI, alpha 3 Extracellular matrix organization Dolichyl- 2.990 2.673 0.049 diphosphooligosaccharide Gene protein glycosyltransferase Transcription/ Protein modification, PTMs 4.011 3.579 0.040 Arginine kinase Translation Dolichyl- 3.020 2.360 0.016 diphosphooligosaccharide

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protein glycosyltransferase 48 kDa subunit 3.854 3.596 0.048 14-3-3 protein Peptidyl-prolyl cis-trans 3.474 2.958 0.044 isomerase Chaperone, protein activity 3.578 3.200 0.041 Heat shock protein 90 stabilization/regulation Protein disulfide- 2.515 2.191 0.038 isomerase A6 2.431 2.025 0.019 Histone H2A 2.493 2.080 0.027 Histone H2A Leucine-rich repeat Chromatin function, gene 3.618 3.309 0.035 flightless-interacting transcription protein 2 4.961 4.649 0.042 Histone H4 Heterogeneous nuclear 1.968 0 0.024 ribonucleoprotein 87F 2.738 2.356 0.029 60S ribosomal protein L7a 3.071 2.707 0.036 40S ribosomal protein S4 Ribossome constituent/function, 3.332 2.965 0.041 40S ribosomal protein S3 protein translation 3.050 2.675 0.042 40S ribosomal protein S13 3.250 2.727 0.043 40S ribosomal protein S25 3.292 3.038 0.048 60S ribosomal protein L11 1.482 0 0.043 Clathrin heavy chain 1 Vesicle mediated transport 2.565 2.174 0.011 Annexin B9 Transport 3.565 3.113 0.038 ADP, ATP carrier protein Voltage-dependent anion Transporters, carrier proteins 3.667 3.260 0.046 channel 6-phosphogluconate Synthesis of NADPH and 5-carbon 2.595 2.247 0.0468 dehydrogenase sugar precursor Other Processes decarboxylating Glutathione S-transferase Detoxification, response to 3.284 2.942 0.042 sigma 2 oxidative stress Note: See Supplementary Table 1 for a summary of LC-MS/MS protein identification results.

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A set of more than ten proteins related with structural constituents and functions were also found to be downregulated in exposed larvae, particularly proteins responsible for actin filament binding (muscle contraction) and structural constituents of the cytoskeleton, including ciliary microtubules units (tubulin alpha and beta chains, and radial spoke head proteins). The impairment in muscle contraction might compromise the activity of the larva foot that is the major organ responsible for the production of threads and all the steps of the adhesive metabolism that enables attachment [57]. Tubulin isoforms are the units forming microtubules that are the basic structural elements of mussel cilia present in the gills, and radial spoke head proteins are important components of the axoneme that forms the core of cilia. The cilia have the major function of driving water flow through filtration for respiration and feeding along the ventral groove towards the mouth, so failure in the repair and generation of new cilia might be of great importance in the stress condition and health status of mussels [58]. These molecular consequences related to the modulation of ciliary constituents were previously observed in Mytilus sp. in response to warm and cold acclimated conditions [59]. Proteins related with regulatory and transcription/translation processes were also found to be modulated by exposure to portoamides (Table 2). Overall, these molecular alterations in mussel plantigrade larvae exposed to portoamides seem to be related as a drop in ATP production would contribute to a decrease of energetic requirements available for filtration, and thus this contributes to the downregulation of synthesis and repair processes of the gill ciliary epithelia As the observed mode of action seems to be the failure in proton-transporting ATPases activity, it represents a broad-spectrum effect, increasing the relevance of portoamides as an AF agent as it would be effective in a wide range of biofouling species. In addition, considering that the anti-settlement activity is reversible in mussel larvae after a recovery period (Figure 2), this seems to indicate that these molecular targets are subjected to chemical and physiological modulation not causing a permanent damage of the mechanisms responsible for the activity. This might be particularly important in the AF scenario, since the AF active ingredients incorporated into paints should act in a repulsive way and target biofouling species should remain able to colonize alternative substrates. This combination of AF effectiveness, low toxicity and mode of action reversibility is the starting point of the development of suitable AF active ingredients. Some marine natural compounds have also been discovered meeting these characteristics, such as the compound polymeric alkylpyridinium salts (poly-APS) isolated from the Mediterranean marine sponge Haliclona (Rhizoniera) sarai and their synthethic

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analogues which act by neurotransmission blockade/modulation through acetylcholinesterase inhibition, combined with surfactant-like properties that decrease surface tension [60,61]. The observed AF profile of portoamides in this work suggests that the mode of action is dictated by different mechanisms depending on the target groups, and the combination of functionalities of these natural products may serve as an inspiration for the development of new AF agents.

4.3. Materials and Methods

4.3.1. Portoamides Isolation and Purification

The cyanobacterial strain Phormidium sp. LEGE 05292 (LEGE-CC) was cultured and upscaled in the laboratory for the isolation of portoamides A and B, at a proportion of 3:1 (Supplementary Figure S1), as described in [56]. This defined mixture of portoamides, produced naturally, was maintained and used in all the following experiments and hereafter referred to as portoamides.

4.3.2. Antifouling Bioassays

4.3.2.1. Mussel Larvae Antisettlement Bioassays

Mussel (Mytilus galloprovincialis) juveniles (0.5 cm in diameter) were sampled in intertidal pools during low neap tides, at Memória Beach, North Portugal (41°13’59” N; 8°43’28” W) and appropriately transported to the laboratory. Mussel plantigrade larvae (0.5–2 milimeters) were screened among the mussel aggregates in a binocular magnifier (Olympus SZX2-ILLT, Tokyo, Japan), isolated with filtered seawater and gently washed to remove adhered organic particles immediately before the bioassays. The selected M. galloprovincialis plantigrade larvae were

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further screened for typical explored behaviour before bioassays. The anti-settlement bioassays were performed using 5 larvae per replicate in 24-well microplates with 4 replicates per condition during 15 h, at 18 ± 1 °C, in the darkness [30]. Test solutions were prepared in filtered seawater and obtained by serial dilution (2, 4, 8, 16 and 32 µM) of the compounds stock solution (3.65 mM) in DMSO. A negative control (0.01% DMSO) and a positive control (5 μM CuSO4) were included in all bioassays. After exposure, the anti-settlement activity was determined by the presence/absence of efficiently attached byssal threads produced by each individual larva for all the conditions tested, and the semi-maximum response concentration that inhibited 50% of larval settlement (EC50) and the median lethal dose (LC50) was determined, if applicable.

4.3.2.2. Mussel Larvae Settlement Recovery Bioassays

A settlement recovery bioassay was performed to test if the anti-settlement effect observed after acute exposure (15 h) to portoamides is reversible after the same period of time in mussel larvae. The initial exposure bioassay was conducted in the same conditions described in Section 3.2.1. Plantigrade larvae were exposed for 15 h to two portoamides concentrations, 3 and 16 µM, corresponding to the mussel larvae EC50 concentration and the concentration used for the mode of action determination, respectively. Negative (0.01% DMSO) and positive controls (5 μM CuSO4) were included. After exposure, the anti-settlement activity was confirmed by the absence of efficiently attached byssal threads produced by each individual larva when compared to the negative control. Then, the same larvae subjected to portoamides exposure were individually transferred to new test plates with fresh filtered seawater. Controls were included. After 15 h, settlement was evaluated again for all the conditions tested.

4.3.2.3. Antibacterial Bioassays

For antibacterial activity screening, five strains of marine biofilm-forming bacteria Cobetia marina CECT 4278, Vibrio harveyi CECT 525, Halomonas aquamarina CECT 5000,

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Pseudoalteromonas atlantica CECT 570 and Roseobacter litoralis CECT 5395 from the Spanish Type Culture Collection (CECT, Valencia, Spain) were inoculated and incubated for 24 h at 26 °C in Marine Broth (Difco) at an initial density of 0.1 (OD600) in 96-well flat bottom microtiter plates and exposed at the test concentration. Bacterial growth inhibition in the presence of the tested compounds was determined in quadruplicate at 600 nm using a microplate reader (Biotek Synergy HT, Winooski Vermont, USA). Marine broths with 0.1% DMSO and 1:100 penicilin- streptomycin-neomycin stabilized solution (P4083 were used as negative (B) and positive (C) controls, respectively. Statistically significant differences between treatments and control measurements were calculated using a Student’s t-test with a 95% confidence level.

4.3.2.4. Antimicroalgal Bioassays

For the antimicroalgal screening, four strains of benthic marine diatoms, Cylindrotheca sp., Halomphora sp., Nitzschia sp. and Navicula sp. were purchased from the Spanish Collection of Algae (BEA-Banco Español de Algas, Las Palmas, Spain) and inoculated in f/2 medium (with silica at an initial concentration of 2–4 × 106 cells per millilitre, and grown in 96-well flat bottom microtiter plates for 10 days at a temperature of 20 °C. Diatoms growth inhibition in the presence of portoamides at 10 µg/mL was determined in quadruplicate, and quantified by the difference between cell densities among treatments, counted using a Neubauer counting chamber. f/2 medium with 0.1% DMSO and cycloheximide (3.55 µM) were the negative and the positive controls, respectively. Statistically significant differences between treatments and control measurements were calculated using the Student’s t-test with a 95% confidence level. The antibacterial and antimicroalgal activity of a commercial agent (ECONEA®) was also included in the same bioassay conditions (8.7 µM) as a reference AF compound. The compounds which showed significant inhibitory activity in screening assays were selected for further determination of maximal inhibitory concentrations (IC). A serial dilution of the stock solution was used to obtain final concentrations from 13.5 μM to 3 μM.

4.3.2.5. Biofilm Inhibition Assay

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To test the disruption activity of portoamides against marine bacterial biofilms, the MBEC™ (Minimum biofilm eradication concentration) assay system was employed (Innovotech Inc. Edmonton, Canada) following the protocol of Ceri et al. [49]. Marine bacterial biofilm formation was established by adding 200 μL of bacterial inoculum in MB medium (Difco) to MBEC plates and incubating them stagnant at 25 °C for 3 days (Cobetia marina, Vibrio harveyi, Halomonas aquamarina and Pseudoalteromonas atlantica) or 5 days (Roseobacter litoralis). After biofilm formation for each bacterium, the peg lids of each of the MBEC plates were aseptically removed and rinsed twice with sterile PBS. The plate lids with bacterial biofilms were then added to new 96-well flat bottom microtiter plates (Orange Scientific, Braine- L’Alleud, Belgium) with fresh MB medium and incubated overnight at 25 °C for 24 h and exposed to portoamides (concentrations from 0.41 μM to 52 μM). DMSO (0.1%) was used as a negative control, while a 4:100 dilution penicilin-streptomycin-neomycin stabilized solution (P4083) was used as a positive control for biofilm disruption. To determine biofilm disruption, the peg lids of each of the MBEC plates containing bacterial biofilms were washed twice for 1 min with sterile PBS. Afterwards, the lids of the MBEC plates were placed in a new standard 96-well microtiter plate containing 200 μL of sterile of MB medium. The removal of biofilm cells from the pegs of the MBEC plates was achieved by sonication for 5 min at 30 kHz, and vortexing for 30 s. Afterwards, the MBEC lid is replaced with a standard 96-well plate lid and biofilm susceptibility to portoamides was then measured by reading optical density at 600 nm using a microplate reader (Biotek Synergy HT, Winooski Vermont, USA). Bacterial growth and turbidity in this situation correspond exclusively to biofilm cell survival which allows for the determination of MBEC.

4. 3.2.6. Quorum-Sensing Inhibition Assay

4.3.2.6.1 Disk-Diffusion Assay

A standard disk-diffusion assay was used to detect anti-quorum sensing activity of the compounds by using a biomonitor strain of Chromobacterium violaceum (ATCC 12472) following

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the protocol developed by McLean [62] with slight modifications. This reporter bacterium regulates violacein production of the pigment by N-hexanoyl-L-homoserine lactones (C6-HSL) and is inhibited by acylated homoserine lactones (AHLs) analogues and other antagonists. C. violaceum 12472 was grown in Luria-Bertani (LB) broth with 1.2% agar. Five milliliters of molten LB agar (0.3%, w/v) was inoculated with 50 mL of a culture of the C. violaceum 12472 grown overnight in LB. The agar-culture solution was immediately poured over the surface of pre- warmed LB agar plates. Twenty microliters of portoamides (6.5 µM) were pipetted on sterile paper discs, which were then placed on the solidiﬁed agar. The plates were incubated overnight at 30 °C and examined for violacein production. Quorum-sensing inhibition (QSI) was detected by a colourless, opaque, but viable halo around the discs. Growth inhibition representing antibacterial activity was detected by a transparent zone/growth inhibition.

4.3.2.6.2 Quorum-Sensing Inhibition (QSI) Quantitative Assay (Dose–Response Violacein Inhibition Assay)

To quantify the inhibition of QS by the portoamides, the QS reporter strain Chromobacterium violaceum (CECT 494/ ATCC 12472) was selected following the protocol of Martinelli [63] and Gemiarto [64]. This strain was incubated in LB medium (Difco) with 3.25 µM, 6.5 µM and 13 µM of portoamides in 96-well flat bottom microtiter plates. LB medium with sterilized ultra-pure water was used as a negative control, and a 1:100 penicilin-streptomycin-neomycin stabilized solution (P4083) was used as a positive control for the inhibition of cell growth. The cultures were grown overnight (18 h) at 26 °C with a constant shaking of 50 rpm. After the incubation period, the portoamides-treated culture and the controls were used for the determination of OD at 720 nm (to quantify cell density), using a microplate reader (Biotek Synergy HT, Vermont, USA). Afterwards, the plates were dried for 30 min at 60 °C and violacein was re-solubilized by the addition of 200 μL of DMSO and the OD was read at 577 nm (to quantify violacein production). Mean OD720/ OD 577 ratios of the portoamides cultures were compared with the untreated cultures using an unpaired Student’s t-test (p = 0.05) and used to assess the degree of violacein inhibition.

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4.3.3. Ecotoxicity Bioassay

Portoamides were tested for marine ecotoxicity using the Artemia salina nauplii lethality test [30,52]. A. salina eggs were allowed to hatch in nutrient-enriched seawater for approximately 48 h at 25 °C. Test concentrations were 25 and 5 µg/mL and test solutions were prepared in filtered seawater. Bioassays were performed in the darkness in 96-well microplates with eight replicates per condition and 15−20 nauplii per well. K2Cr2O7 (13.6 µM) was included as a positive control and 1% DMSO was used as a negative control. The percentage of mortality was assessed at the end of the exposure period.

4.3.4. Antifouling Mode of Action Towards Mytilus Larvae

4.3.4.1. Sample Preparation

Ten M. galloprovincialis whole plantigrade larvae from the anti-settlement bioassays were used per replicate (four replicates per condition were used) and incubated in lysis buffer with 2% (w/v) SDS, 100 mM Tris-HCl, 0.1 M DTT and protease inhibitors (Roche, 11697498001, Basel, Switzerland), pH 7.6 for 1 h with mixing (450 rpm). Additional denaturation was achieved by sample heating (95 °C, 3 min). All samples were clarified at 16,000× g for 20 min and total protein was estimated based on the absorbance at 280 nm (1 Abs = 1 mg/mL protein) with Nanodrop (Thermo Scientific, Waltham, MA, USA) before storage at −80 °C. Proteins were subsequently digested following filter aided sample preparation (FASP) [65] with modifications. Briefly, protein samples (100 μg) were alkylated with iodoacetamide, incubated with trypsin (Roche, 03708985001) at an enzyme-to protein ratio of 1:50 (w/w) for 16 h at 37 °C, in 30 kDa nominal molecular weight limit (NMWL) centrifugal filter units (MRCF0R030, Millipore, Billerica, MA, USA). Peptide samples were loaded in detergent removal spin columns (Thermo Scientific) and the manufacturer instructions followed to get remove detergent residues from the samples. The

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samples were acidified with formic acid and dried in a Centrivap centrifugal concentrator (Labconco, Kansas City, MO, USA). Finally, the samples were resuspended in 50 μL of mixture composed of 0.1% FA and 2% acetonitrile.

4.3.4.2. LC-MS/MS Analysis

LC-MS/MS was performed on a nanoflow Ultimate 3000 UPLC (Dionex) coupled to an Impact HD mass spectrometer equipped with a CaptiveSpray source (Bruker Daltonik, Bremen, Germany). For each sample, 1 µL of the sample was loaded on a C18 PepMap100 nano-Trap column (300 µm, ID × 5 mm, 5 micron 100 Å) at a flow rate of 3000 nL/min. The trap column was then switched in line with the ProntoSIL C18AQ analytical column (100 µm, ID × 150 mm, 3 micron 200 Å). The reversed phase elution gradient was from 2% to 20% to 45% B over 60 min, total 85 min at a flow rate of 1000 nL/min. Solvent A was LCMS-grade water with 0.1% formic acid; solvent B was LCMS-grade ACN with 0.1% FA. The mass spectrometer was set up in positive ionization MS mode with a mass range of m/z 130–2200. All samples (including biological repeats) were analysed in duplicate. To identify the peptides of interest, a pool from each treatment group was made by combining 5 µL of every sample digest and duplicates were run on the Impact HD mass spectrometer. More specifically, the pooled samples were measured in data-dependent MS/MS mode, where the acquisition speed was 2 Hz in MS and 1–20 Hz in MS/MS mode depending on precursor intensity. Ten precursors were selected in the m/z range of 350–1200, with preference for doubly or triply charged peptides. The analysis was performed in positive ionization mode with a dynamic exclusion of 60 s.

4.3.4.3 Protein Identification and Quantification

Following LC-MS/MS analysis, the QTOF data were searched using the Peaks Studio 8.5 search algorithm (Bioinformatics Solutions, Waterloo, ON, Canada) against an in-house Mytilus sequence database and the uniprot-mollusca-200718-290-109 sequence database. The following

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parameters were used: A precursor mass tolerance of 0.15 Da and a fragment mass tolerance of 0.15 Da were allowed, trypsin was specified as the digestive enzyme, and up to 2 missed cleavages were allowed. Carbamidomethyl (C) was specified as a fixed modification, and oxidation (M) and deamidation (NQ) were chosen as variable modifications. A false discovery rate (FDR) of 1% was used for peptide identification in Peaks. In addition, the Peptide Hit Threshold (−10log_) was set to 30, de novo only 15% of ALC, and only proteins with a minimum of 1 unique peptide identification were included.

4.3.5. Data Analysis

One-way analysis of variance (ANOVA) followed by a Dunnett test against the DMSO control (p < 0.01) was used to analyse anti-settlement, antibacterial and anti-microalgal screening data as well as Artemia salina ecotoxicity data. The semi-maximum response concentration that inhibited

50% of mussel larval settlement (EC50), response concentration that inhibited bacterial growth (IC) and the median lethal dose (LC50) were assessed using Probit regression analysis, when applicable. Pearson Goodness-of-fit (Chi-Square) significance was considered at p < 0.01 for this analysis, and a 95% lower confidence limit (LCL) and a 95% upper confidence limit (UCL) were presented. In proteomic analysis, the mixed model ANOVA was utilized to report protein abundance differences between treatments at a confidence level of 95%. The mixed model ANOVA was performed on log10(x + 1)-transformed values from 4 biological replicates. Unless otherwise stated, the software IBM SPSS Statistics (version 21, SPSS Inc., Chicago, Illinois, USA) was used for statistical analysis.

4.4. Conclusions

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This study elucidates the potential of portoamides as new eco-friendly AF agents, given their high effectiveness in the prevention of the attachment of mussel larvae, enabling the reversibility of the effect after exposure, and their antibacterial properties towards marine biofilm-forming bacteria, while lacking the toxicity towards both target and non-target species. This AF potential is even further enhanced by the fact that the major molecular target of portoamides, in the important macrofouling species Mytilus galloprovincialis, is the failure in ATPase proton- transporting activity, and thus portoamides could be promising as an effective way of controlling the attachment of a wide range of biofouling species.

Supplementary Materials:

The following are available at the Supplementary info (Chapter 7). Summary of LC-MS/MS protein identification results. Figure S1: Relative proportions of portoamides A and B. Absorption spectra (A) obtained by the analytical method with the absorbance as a function of time. The first peak represents portoamide A, while the second peak is portoamide B. The PDA spectrum (B), for each absorbance spectrum, with absorbance at the wavelength of 276.0 nm.

Author Contributions: Scientific and experimental rational conceptualization, J.R.A.; production and isolation of portoamides, T.R.; antifouling bioassays, J.A. (antibacterial, antimicroalgal, antibiofilm, and quorum-sensing inhibition), J.R.A. and S.P. (antisettlement); ecotoxicity bioassays, J.R.A. and S.P.; proteomic analysis, A.C., J.R.A., J.E.P., A.T. and S.C.; writing of the original draft preparation, J.A., T.R., A.C. and J.R.A.; writing of review and editing, VV, JEP, AT, and SC; supervision, J.R.A. and V.V.; funding acquisition, J.R.A. and V.V.

Funding: Funding for this work was provided by the Strategic Funding UID/Multi/04423/2013 through national funds provided by the Foundation for Science and Technology (FCT) and the

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European Regional Development Fund (ERDF), in the framework of the programme PT2020 and by the project NASCEM PTDC/BTA-BTA/31422/2017 (POCI-01-0145-FEDER-031422), financed by the FCT, COMPETE2020 and PORTUGAL2020. J.A. and A.C. also acknowledge the FCT for PhD and Postdoctoral scholarships SFRH/BD/ 99003/2013 and SFRH/BPD/103683/2014, respectively.

Acknowledgments: We wish to thank C.v.K. for statistical analysis.

Conflicts of Interest: The authors declare no conflicts of interest.

6. References

1. Salta, M.; Wharton, J.A.; Blache, Y.; Stokes, K.R.; Briand, J.F. Marine biofilms on artificial surfaces: Structure and dynamics. Environ. Microbiol. 2013, 15, 2879–2893. doi:10.1111/1462-2920.12186. 2. Antunes, J.; Leão, P.; Vasconcelos, V. Marine biofilms: Diversity of communities and of chemical cues. Environ. Microbiol, Rep. 2018, doi:10.1111/1758-2229.12694. 3. Dang, H.; Lovell, C.R. Microbial surface colonization and biofilm development in marine environments. Microbiol. Mo. Bio. Rev. 2016, 80, 91–138, doi:10.1128/MMBR.00037. 4. Qian, P.Y.; Lau, S.C.K.; Dahms, H.U.; Dobretsov, S.; Harder, T. Marine Biofilms as Mediators of Colonization by Marine Macroorganisms: Implications for Antifouling and Aquaculture. Mar. Biotechnol. 2007, 9, 399–410, doi:10.1007/s10126-007-9001-9.

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5. Hadfield, M.G. Biofilms and marine invertebrate larvae: What bacteria produce that larvae use to choose settlement sites. Ann. Rev. Mar. Sci. 2011, 3, 453–470, doi:10.1146/annurev- marine-120709-142753. 6. Mieszkin, S.; Callow, M.E., Callow, J. Interactions between microbial biofilms and marine fouling algae: A mini review. Biofouling 2013, 29, 1097–1113, doi:10.1080/08927014.2013.828712. 7. Schultz, M.P.; Swain, G.W. The influence of biofilms on skin friction drag. Biofouling 2000, 15,129–139, doi:10-1080/08927010009386304. 8. Schultz, M.P. Effects of coating roughness and biofouling on ship resistance and powering. Biofouling 2007, 23, 331–341, doi:10.1080/08927010701461974. 9. Costas, E.; Gonzalez, R.; López-Rodas, V.; Emma Huertas, I. Mutation of Microalgae from Antifouling Sensitivity to Antifouling Resistance Allows Phytoplankton Dispersal through Ships’ Biofouling. Biological. Invasions 2013, 15, 1739–1750, doi:10.1007/s10530-012- 0405-8. 10. Yebra, D.M.; Kiil, S.; Dam-Johansen, K. Antifouling Technology―Past, present and future steps towards efficient and environmentally friendly antifouling coatings. Prog. Org. Coat. 2004, 50, 75–104, doi: 10.1016/j.porgcoat.2003.06.001. 11. De Carvalho, C.C.C.R. Marine biofilms: A successful microbial strategy with economic implications. Front. Mar. Sci. 2018, 5, 126, doi:10.3389/fmars.2018.00126. 12. Chambers, L.D; Wharton, J.A.; Wood, R.J.K.; Walsh, F.C.; Stokes, K.R. Techniques for the measurement of natural product incorporation into an antifouling coating. Prog. Org. Coat. 2014, 77, 473–484, doi: 10.1016/j.porgcoat.2013.11.013. 13. Dafforn, K.A.; Lewis, J.A.; Johnston, E.L. Antifouling strategies: History and regulation, ecological impacts and mitigation. Mar. Pollut. Bull. 2011, 62, 453–465, doi: 10.1016/j.marpolbul.2011.01.012. 14. Callow, J.A.; Callow, M.E. Trends in the development of environmentally friendly fouling resistant marine coatings. Nat. Commun. 2011, 2, 244, doi:10.1038/ncomms1251. 15. Qian, P-Y.; Li, Z; Xu, Y.; Li, Y.; Fusetani, N. Mini-review: Marine natural products and their synthetic analogs as antifouling compounds: 2009–2014. Biofouling 2015, 31, 101–122, doi:10.1080/08927014.2014.997226.

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16. Wang, K.L.; Wu, Z.H.; Wang, Y.; Wang, C.Y.; Xu, Y. Mini-Review: Antifouling natural products from marine microorganisms and their synthetic analogs. Mar. Drugs 2017, 15, 266, doi:10.3390/md15090266. 17. Dahms, H.U.; Ying, X.; Pfeiffer C. Antifouling potential of cyanobacteria: A mini-review. Biofouling 2006, 22, 317–327, doi:10.1080/08927010600967261. 18. Almeida, J.R.; Vasconcelos, V. Natural antifouling compounds: Effectiveness in preventing invertebrate settlement and adhesion. Biotechnol. Adv. 2015, 33, 343–357, doi: 10.1016/j.biotechadv.2015.01.013. 19. Fusetani, N. Biofouling and antifouling. Nat. Prod. Rep. 2004, 21, 94–104, doi:10.1039/B302231P. 20. Fusetani, N. Antifouling marine natural products. Nat. Prod. Rep. 2011, 28, 400–441, doi:10.1039/c0np00034e. 21. Chen, L.; Qian, P-Y. Review on molecular mechanisms of antifouling compounds: An update since 2012. Mar. Drugs 2017, 15, 264, doi:10.3390/md15090264. 22. Dahms, H.U.; Dobretsov, S. Antifouling Compounds from Marine Macroalgae. Mar. Drugs 2017, 15, 265, doi:10.3390/md15090265. 23. Burja, A.M.; Banaigs, B.; Abou-Mansour, E.; Burgess, J.G.; Wright, P.C. Marine cyanobacteria―A prolific source of natural products. Tetrahedron 2001, 57, 9347–9377, doi:10.1016/S0040-4020(01)00931-0. 24. Qian, P.Y.; Chen, L.; Xu, Y. Mini-Review: Molecular Mechanisms of Antifouling Compounds. Biofouling 2013, 29, 381–400, doi:10.1080/08927014.2013.776546. 25. Leao, P.N.; Pereira, A.R.; Liu, W.T.; Ng, J.; Pevzner, P.A.; Dorrestein; P.C.; Konig, G.M.; Vasconcelos, V.M; Gerwick, W.H. Synergistic allelochemicals from a freshwater cyanobacterium. Proc. Natl. Acad. Sci. U. S. A. 2010, 107, 11183-11188, doi:10.1073/pnas.0914343107. 26. Dias, F.; Antunes, J.T.; Ribeiro, T.; Azevedo, J.; Vasconcelos, V.; Leao, P.N. Cyanobacterial Allelochemicals But Not Cyanobacterial Cells Markedly Reduce Microbial Community Diversity. Front. Microbiol. 2017, 8, 1495, doi:10.3389/fmicb.2017.01495. 27. Diers, J.A.; Pennaka, H.K.; Peng, J.; Bowling, J.J.; Duke, S.O.; Hamann, M.T. Structural Activity Relationship Studies of Zebra Mussel Antifouling and Antimicrobial Agents from Verongid Sponges. J. Nat. Prod. 2004, 67, 2117–2120, doi:10.1021/np040097t.

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28. Diers, J.A.; Bowling, J.J.; Duke, S.O.; Wahyuono, S.; Kelly, M.; Hamann, M.T. Zebra Mussel Antifouling Activity of the Marine Natural Product Aaptamine and Analogs. Mar. Biotechnol. 2006, 8, 366–372, doi:10.1007%2Fs10126-005-6055-4. 29. Ortlepp, S.; Pedpradap, S.; Dobretsov, S.; Proksch, P. Antifouling activity of sponge-derived polybrominated diphenyl ethers and synthetic analogues. Biofouling 2008, 24, 201–208, doi:10.1080/08927010802008096. 30. Almeida, J.R.; Correia-da-Silva, M.; Sousa, E.; Antunes, J.; Pinto, M.; Vasconcelos, V.; Cunha, I. Antifouling potential of Nature-inspired sulfated compounds. Sci. Rep. 2017, 7, 42424, doi:10.1038/srep42424. 31. Almeida, J.R.; Moreira, J., Pereira, D.; Pereira, S.; Antunes, J.; Palmeira, A.; Vasconcelos, V.; Pinto, M., Correira-da-Siva, M., Cidade, H. Potential of synthetic chalcone derivatives to prevent marine biofouling. Sci. Total Environ. 2018, 643, 98–106, doi: 10.1016/j.scitotenv.2018.06.169. 32. Tan, L.T; Goh, B.P; Tripathi, A; Lim, M.G.; Dickinson, G.H.; Lee, S.S; Teo, S.L. Natural antifoulants from the marine cyanobacterium Lyngbya majuscula. Biofouling 2010, 2, 685– 695, doi:10.1080/08927014.2010.508343. 33. Brown, G.D.; Wong, H.F.; Hutchinson, N.; Lee, S.C.; Chan, B.K.K.; Williams, G.A. Chemistry and biology of maculalactone A from the marine cyanobacterium Kyrtuthrix maculans. Phytochem. Rev. 2004, 3, 381–400, doi:10.1007/s11101-004-6552-5. 34. Dobretsov, S.; Abed; R.M.M.; Teplitski, M. Mini-review: Inhibition of biofouling by marine microorganisms. Biofouling 2013, 29, 423–441, doi:10.1016/S1387-2656(00)06024-5. 35. Carl, C.; Poole, A.J.; Vucko, M.J.; Williams, M.R.; Whalan, S.; de Nys, R. Optimising settlement assays of pediveligers and plantigrades of Mytilus galloprovincialis. Biofouling 2011, 27, 859–868, doi:10.1080/08927014.2011.605943. 36. Martins, J.; Leikoski, N..; Wahlsten, M.; Azevedo, J.; Antunes, J.; Jokela, J.; Sivonen, K.; Vasconcelos, V.; Fewer, D.P.; Leão, P. Sphaerocyclamide a prenylated Cyanobactin from the cyanobacterium Sphaerospermopsis sp. LEGE 00249. Sci. Rep. 2018, 8, 14537, doi:10.1038/s41598-018-32618-5. 37. Hanssen, K.Ø.; Cervin, G.; Trepos, R.; Petitbois, J.; Haug, T.; Hansen, E.; Andersen, J.H.; Pavia, H.; Hellio, C.; Svenson, J. The Bromotyrosine Derivative Ianthelline Isolated from the

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Arctic Marine Sponge Stryphnus fortis Inhibits Marine Micro- and Macrobiofouling. Mar. Biotechnol. 2014, 16, 684−694, doi:10.1007/s10126-014-9583-y. 38. Moodie, L.W.K.; Trepos, R.; Cervin, G.; Larsen, L.; Larsen, D.S.; Pavia, H.; Hellio, C.; Cahill, P.; Svenson, J. Probing the structure-activity relationship of the natural antifouling agent polygodial against both micro and macrofoulers by semisynthetic modification. J. Nat. Prod. 2017, 80, 515–525, doi: 10.1021/acs.jnatprod.6b01056. 39. Maréchal, J.P.; Culioli, G.; Hellio, C.; Thomas-Guyon, H.; Callow, M.E.; Clare, A.S.; Ortalo- Magne, A. Seasonal variation in antifouling activity of crude extracts of the brown alga Bifurcaria bifurcata (Cystoseiraceae) against cyprids of Balanus amphitrite and the marine bacteria Cobetia marina and Pseudoalteromonas haloplanktis. J. Exp. Mar. Biol. Ecol. 2004, 313, 47–62, doi: 10.1016/j.jembe.2004.07.016. 40. Tsoukatou, M.; Marechal, J.P.; Hellio, C.; Novakovic, I.; Tufegdzie S.; Sladic, D.; Gasic, M.J.; Clare, A.S., Vagias, C.; Roussis, V. Evaluation of the activity of the sponge metabolites avarol and avarone and their synthetic derivatives against fouling micro- and macroorganisms. Molecules 2012, 12, 1022–1034, doi:10.3390/12051022. 41. Viano, Y.; Bonhomme, D.; Camps, M.; Briand, J.; Ortalo-Magne, A.; Blache, Y.; Piovetti, L.; Culioli, G. Diterpenoids from the Mediterranean Brown Alga Dictyota sp. Evaluated as Antifouling Substances against a Marine Bacterial Biofilm. J. Nat. Prod. 2009, 72, 1299– 1304, doi:10.1021/np900102f. 42. Molino, P.J.; Wetherbee, R. Mini-review: The biology of biofouling diatoms and their role in the development of microbial slimes. Biofouling 2008, 24, 365–379, doi:10.1080/08927010802254583. 43. Inbakandan, D.P.; Sriyutha Murthy, R.; Khan, S.J. 16S rDNA Sequence Analysis of Culturable Marine Biofilm Forming Bacteria from a Ship’s Hull. Biofouling 2010, 26, 893– 899, doi:10.1080/08927014.2010.530347. 44. Liaqat, I.; Bachmann, R.T.; Edyvean, R.G.J. Type 2 quorum sensing monitoring, inhibition and biofilmformation in marine microrganisms. Curr. Microbiol. 2014, 68, 342–351, doi:10.1007/s00284-013-0484-5. 45. Melander, C.; Moeller, P.D.R.; Ballard, T.E.; Richards, J.J.; Huigens III, R.W.; Cavanagh, J. Evaluation of dihydrooroidin as an antifouling additive in marine paint. Int. Biodeter. Biodegr. 2009, 63, 529–532, doi: 10.1016/j.ibiod.2008.08.009.

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46. Bernbom, N.; Ng, Y.Y.; Kjelleberg, S.; Harder, T.; Gram, L. Marine bacteria from danish coastal waters show antifouling activity against the marine fouling bacterium Pseudoalteromonas sp strain S91 and zoospores of the green alga Ulva australis independent of bacteriocidal activity. Appl. Environ. Microbiol. 2011, 77, 8557–8567, doi:10.1128/AEM.06038-1. 47. Bernbom, N.; Ng, Y.Y.; Olsen, S.M.; Gram, L. Pseudoalteromonas spp. serve as initial bacterial attractants in mesocosms of coastal waters but have subsequent antifouling capacity in mesocosms and when embedded in paint. Appl. Environ. Microb. 2013, 79, 6885–6893, doi:10.1128/AEM.06038-11. 48. Brian-Jaisson, F.; Ortalo-Magné A.; Guentas-Dombrowsky, L.; Armougom, F.; Blache Y.; Molmeret, M. Identification of Bacterial Strains Isolated from the Mediterranean Sea Exhibiting different Abilities of Biofilm Formation. Microb. Ecol. 2004, 68, 94–110, doi:10.1007/s00248-013-0342-9. 49. Ceri, H.; Olson, M.E.; Stremick, C.; Read, R.R.; Morck, D.; Buret, A. The Calgary Biofilm Device: New technology for rapid determination of antibiotic susceptibilities of bacterial biofilms. J. Clin. Microbiol. 1999, 37, 1771–1776. 50. Dusane, D.H.; Damare, S.R.; Nancharaiah, Y.V.; Ramaiah, N.; Venugopalan, V.P.; Kumar, A.R.; Zinjarde, S.S. Disruption of microbial biofilms by an extracellular protein isolated from epibiotic tropical marine strain of Bacillus licheniformis. PLoS ONE 2013, 8, e64501, doi: 10.1371/journal.pone.0064501. 51. Richards, J.J.; Melander, C. Controlling Bacterial Biofilms. ChemBioChem 2009, 10, 2287– 2294, doi:10.1002/cbic.200900317. 52. Salta, M.; Wharton, J.A.; Dennington, S.P.; Stoodley, P.; Stokes, KR. Anti-biofilm performance of three natural products against initial bacterial attachment. Int. J. Mol. Sci. 2013, 14, 21757–21780, doi:10.3390/ijms141121757. 53. Armstrong, E.; Boyd, K.G.; Burgess, J.G. Prevention of marine biofouling using natural compounds from marine organisms. Biotech. Ann. Rev.2000, 6, 221–241, doi:10.1016/S1387- 2656(00)06024-5. 54. Dobretsov, S.; Teplitski, M.; Paul, C. Mini-Review: Quorum Sensing in the Marine Environment and Its Relationship to Biofouling. Biofouling 2009, 25, 413–427, doi:10.1080/08927010902853516.

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55. Meyer, B.N.; Ferrigni, N.R.; Putnam, J.E.; Jacobsen, L.B.; Nichols, D.E.; McLaughlin, J.L.; Brine shrimp: a convenient general bioassay for active plant constituents. Planta Med. 1982, 45, 31–34, doi:10.1055/s-2007-971236. 56. Ribeiro, T.; Lemos, F.; Preto, M.; Azevedo, J.; Sousa, M.L.; Leao, P.N.; Vitorino, R.; Vasconcelos, V.; Urbatzka, R. Cytotoxicity of portoamides in human cancer cells and analysis of the molecular mechanisms of action. PloS ONE 2017, 12, doi: 10.1371/journal.pone.0188817. 57. DeMartini, D.G.; Errico, J.M.; Sjoestroem, S.; Fenster, A.; Waite, J.H. A cohort of new adhesive proteins identified from transcriptomic analysis of mussel foot glands. J. R. Soc. Interface 2017, 14, 131, doi:10.1098/rsif.2017.0151. 58. Dral, A.D.G. Movements and Co-ordination of the Latero-frontal Cilia of the Gill Filaments of Mytilus. Nature 1966, 210, 1170–1171, doi:10.1038/2101170a0. 59. Fields, P.A.; Zuzow, M.J.; Tomanek, L. Proteomic responses of blue mussel (Mytilus) congeners to temperature acclimation. J. Exp. Biol. 2012, 215, 1106–1116, doi:10.1242/jeb.062273. 60. Sepčić K.; Turk T. 3-Alkylpyridinium Compounds as Potential Non-Toxic Antifouling Agents. In Antifouling Compounds. Marine Molecular Biotechnology; Fusetani, N., Clare, A.S., Eds.; Springer: Berlin, Heidelberg, 2006. 61. Piazza, V.; Dragić , I.; Sepčić, K.; Faimali, M.; Garaventa, F.; Turk, T.; Berne, S. Antifouling activity of synthetic alkylpyridinium polymers using the barnacle model. Mar. Drugs 2014, 12, 1959–1976, doi:10.3390/md12041959. 62. McLean, R.J.; Pierson, L.S., III; Fuqua, C. A Simple screening protocol for the identification of quorum signal antagonists. J. Microbiol. Methods 2004, 58, 351–360, doi: 10.1016/j.mimet.2004.04.016. 63. Martinelli, D.; Grossmann, G.; Sequin, U.; Brandl, H.; Bachofen, R. Effects of natural and chemically synthesized furanones on quorum sensing in Chromobacterium violaceum. BMC Microbiol. 2004, 4, doi:10.1186/1471-2180-4-25. 64. Gemiarto, A.; Ninyo, N.N.; Lee, W.L.; Logis, J.; Fatima, A.; Chan, E.W.C.; Lim, C.S.Y. Isoprenyl caffeate, a major compound in manuka propolis, is a quorum-sensing inhibitor in Chromobacterium violaceum. Antonie Van Leeuwenhoek. 2015, 108, 491–504, doi:10.1007/s10482-015-0503-6.

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65. Wiśniewski, J.R.; Zougman, A.; Nagaraj, N.; Mann, M. Universal sample preparation method for proteome analysis. Nat. Methods 2009, 6, 359–362.

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Chapter 5: Search for cyanobacterial compounds with antimicrofouling and quorum sensing inhibition activity

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Search for cyanobacterial compounds with antimicrofouling and quorum sensing inhibition activity

Jorge Antunes1,2, Joana Azevedo1,2, Pedro Leão1, Vitor Vasconcelos1,2

2Departamento de Biologia, Faculdade de Ciências, Universidade do Porto, Rua do Campo Alegre, P 4069-007 Porto, Portugal

Abstract

Due to their bioactive potential, cyanobacterial metabolites have the potential to be used as environmentally friendly antifouling compounds and to be a less toxic alternative to the biocide- based coatings currently in use. Against this backdrop, cyanobacteria from the LEGE culture collection, which harbors several hundreds of strains, was screened for their potential antifouling activity. In specific, 61 cyanobacterial crude extracts from the LEGE CC (freshwater, brackish and marine) were selected to be tested for the inhibition of growth of microfouling organisms (bacteria and diatoms), against macroalgae spore attachment, as well as for the inhibition of quorum sensing (QS). Our screening assays demonstrated that the unidentified cyanobacterium strain LEGE 00060, Dolichospermum sp. LEGE 00246, Cuspidothrix issatschenkoi strain LEGE 00247, and Synechococcus sp. strain LEGE 11248 were particularly bioactive and were therefore selected to be grown in higher scale, in order to allow for the purification of their active constituent(s). Bioassay-guided isolation showed that the strain LEGE 11248, a marine subtidal cyanobacterium collected in Portugal, had the highest antifouling potential taking in consideration all the conducted assays. This strain was therefore selected for the attempt of the isolation and identification of bioactive compounds for that activity. This effort resulted on several potent bioactive sub-fractions of the marine strain LEGE with anti-microfouling and quorum-inhibition potential that will require further characterization.

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Keywords: Antifouling; biofilms; cyanobacteria metabolites; natural products; antifouling; quorum-sensing

5. 1. Introduction

Marine biofouling causes serious problems for marine industries worldwide, particularly the shipping industry, and raises both economic and environmental problems (Yebra et al., 2004). Macrofouling organisms such as invertebrates and macroalgae are responsible for a significant percentage of marine fouling biomass and for the diminished hydrodynamic performance of a ship (Schultz, 2007). However, marine biofilms and microfouling organisms (such as bacteria and diatoms) that attach to artificial surfaces are also recognized to be a significant issue for a wide range of submerged engineered structures and are known to have a significant role in the corrosion of surfaces immersed in seawater (Salta et al., 2013; Antunes et al., 2018). The most commonly used strategy to prevent marine biofouling is based on antifouling (AF) coatings containing bioactive compounds (Wu et al., 2017). After years of application of toxic antifoulants to control marine biofouling, the International Maritime Organization (IMO) banned the use of organotin compounds in 2008, making urgent the development of environmentally friendly fouling-resistant coatings (Qian et al., 2013). The demand of AF coatings incorporating non-toxic biocides has therefore increased, and they are now subject to restrictive legislations (e.g., the European Biocidal Products Directive, which requires several risk studies before registration and marketing authorization) (Hellio et al., 2002). One of the alternatives has been the utilization of antifouling products of natural origin, particularly searching for natural compounds among marine organisms (Fusetani, 2004, 2011). Marine microorganisms in particular are a good source of anti-fouling compounds, as they can be cultivated in large scales, and genomic tools can be used to identify the genes responsible for biosynthesis of the active metabolites (Dahms et al., 2006; Dahms and Dobretsov, 2017). In the context of natural products biodiscovery, cyanobacteria are therefore a particularly encouraging source of new natural compounds, as they produce a wide range of secondary

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metabolites with recognized activity on several different biological responses (Burja et al., 2001; Leão et al., 2012; Dahms and Dobretsov, 2017) including effective and less toxic alternatives to combat marine biofouling and replace the biocide-based coatings currently in use (Almeida and Vasconcelos, 2015; Wang et al., 2017). Cyanobacteria can easily be cultured and produce compounds much more rapidly and in large amounts when compared to invertebrates and algae, while being amenable to growth in mass culture, and manipulated to achieve optimal production of bioactive substances, sustaining yields on an industrial scale (Bhadury and Wright, 2004; Dahms et al., 2006). In spite of this potential of cyanobacteria for the production of AF products, cyanobacteria are still underexplored as a source of potential AF compounds, with few compounds from cyanobacteria been identified so far (Dahms et al., 2006; et al., 2017; Antunes et al., 2019). Our laboratory maintains a culture collection (LEGE Culture Collection) of over 400 strains of cyanobacteria, including diverse marine, brackish water and freshwater strains, which have been collected mostly in the Portuguese aquatic environments, either from the continent, as well as some strains from the Madeira and Azores archipelagos (Ramos et al., 2018). Several of these strains have shown interesting bioactive profiles using other marine organisms - bacteria, microalgae and metazoans) as targets, as well having anti-cancer, anti-viral, anti-microbial, or anti- biofouling properties (Leão et al., 2013; Freitas et al., 2015, Ribeiro et al., 2017; Antunes et al., 2019). Therefore, it is predicted that these diverse cyanobacterial strains harbor a largely unknown chemical diversity and bioactive potential, which will form the basis for discovery of anti-fouling compounds among the LEGE CC cyanobacterial strains.

5.2 Materials and methods

5.2.1. Cyanobacteria growth and primary screening

An initial primary screening was conducted with an extensive panel of crude extracts obtained from 61 cyanobacterial strains (Table 1) both from freshwater, brackish and the marine

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environment. The selected cyanobacterial strains are part of the LEGE – CIIMAR culture collection ( http://www.ciimar.up.pt/legecc/), and were cultured in an appropriate medium (Z8 medium (Kotai, 1972) for freshwater strains; the same Z8 medium was used for marine strains but supplemented with 25 g/L NaCl and 10 µg/mL vitamin B12, for enrichment). Culturing was carried out in 6 L flasks (with 4 L medium each) under a temperature of 25°C with a light–dark cycle of 14:10 h and 25 μE m−2 s −1 of photon irradiance.

Table 1 - LEGE culture collection strains for which biomass extraction has been carried out.

LEGE code Strain ID Location Environment

LEGE 00038 Synechocystis salina Portugal Marine

LEGE 00041 Synechocystis salina Portugal Marine

LEGE 00049 Unidentified Oscillatoriales Morocco Freshwater

LEGE 00052 Unidentified Synechoccales Morocco Freshwater

LEGE 00053 Unidentified Synechoccales Morocco Freshwater

LEGE 00055 Phormidium cf. irriguum Morocco Freshwater

LEGE 00060 Unidentified cyanobacterium Morocco Freshwater

LEGE 00065 Unidentified cyanobacterium Morocco Freshwater

LEGE 00246 Dolichospermum sp. Portugal Freshwater

LEGE 00247 Cuspidothrix issatschenkoi Portugal Freshwater

LEGE 00259 Dolichospermum sp. Portugal Freshwater

LEGE 03273 Oscillatoria sp. Portugal Freshwater

LEGE 03278 Dolichospermum sp. Portugal Freshwater

LEGE 04288 Nodularia sp. Portugal Freshwater

LEGE 04357 Nostoc aff. commune Morocco Freshwater

LEGE 05292 Phormidium sp. Portugal Freshwater

LEGE 06013 cf. Romeria sp. Portugal Marine

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LEGE 06014 Nodosilinea sp. Portugal Marine

LEGE 06020 Nodosilinea sp. Portugal Marine

LEGE 06077 Nostoc sp. Portugal Brackish

LEGE 06105 Plectonema cf. radiosum Portugal Marine

LEGE 06109 Cyanobium sp. Portugal Marine

LEGE 06110 Nodosilinea sp. Portugal Marine

LEGE 06113 Cyanobium sp. Portugal Marine

LEGE 06114 Plectonema cf. radiosum Portugal Marine

LEGE 06116 Unidentified Synechoccales Portugal Marine

LEGE 06118 Unidentified Synechoccales Portugal Marine

LEGE 06119 Nodosilinea sp. Portugal Marine

LEGE 06130 Cyanobium sp. Portugal Marine

LEGE 06134 Cyanobium sp. Portugal Marine

LEGE 06135 Cyanobium sp. Portugal Marine

LEGE 06137 Cyanobium sp. Portugal Marine

LEGE 06140 Cyanobium sp. Portugal Marine

LEGE 06141 cf. Oculatella sp. Portugal Marine

LEGE 06143 aff. Nodosilinea sp. LEGE 06148 Portugal Marine

LEGE 06148 aff. Nodosilinea sp. LEGE 06148 Portugal Marine

LEGE 06174 Chroococcidiopsis sp. Portugal Marine

LEGE 06223 Planktothrix mougeotii Portugal Freshwater

LEGE 06224 Planktothrix mougeotii Portugal Freshwater

LEGE 06225 Planktothrix mougeotii Portugal Freshwater

LEGE 06361 Leptolyngbya sp. Portugal Freshwater

LEGE 07075 unidentified Synechococcales Portugal Brackish

LEGE 07173 Cyanobium sp. Portugal Marine

LEGE 07197 Tychonema sp. Portugal Freshwater

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LEGE 07198 Tychonema sp. Portugal Freshwater

LEGE 07221 Tychonema sp. Portugal Freshwater

LEGE 07229 Planktothrix mougeotii Portugal Freshwater

LEGE 08334 Sphaerospermopsis sp. Portugal Freshwater

LEGE 09399 Cyanobium gracile Portugal Freshwater

LEGE 11428 Synechococcus sp. Portugal Marine

LEGE 12432 Unidentified Synechoccales Portugal Freshwater

LEGE 14444 Tychonema borneti Colombia Freshwater

LEGE 14445 Tolypothrix sp. Colombia Freshwater

LEGE 91093 Microcystis aeruginosa Portugal Freshwater

LEGE 91342 Microcystis aeruginosa Portugal Freshwater

LEGE 91351 Microcystis aeruginosa Portugal Freshwater

LEGE X- 002 Anabaena sp. Finland Freshwater

LEGE XX061 Unidentified cyanobacterium Portugal Freshwater

LEGE XX280 Planktothrix sp. Portugal Freshwater

LEGE X-357 Unidentified cyanobacterium Portugal Freshwater

Col.1 HB Unidentified cyanobacterium Portugal Freshwater

5.2.2. Extraction, purification and identification of antifouling compounds

Cyanobacterial biomass (0.5 g) was extracted with a 2:1 (v/v) mixture of CH2Cl2/MeOH at ambient temperature, followed by three extractions with the same mixture of solvents. The slurry was centrifuged (4495  g). Supernatants were pooled together, the solvent completely evaporated, and yield calculated (4-27%). All extracts were dissolved in DMSO at 30 or 10 μg/mL

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concentration, serving as stock test solutions for the antibacterial assays and anti-microalgal assay. Freeze-dried biomass samples were extracted accordingly to Edwards et al., (2004). The strains which yielded bioactive crude extracts were re-inoculated and cultivated at a larger scale (~120 L) their biomass harvested, freeze-dried and re-extracted. The large-scale organic extracts were then fractionated using vacuum liquid chromatography (VLC), using Silica gel 60, with a gradient system of solvents from the non-polar hexanes to the more polar methanol. In specific, fractions were obtained using a solvent gradient from 90% hexane to 100% EtOAC to 100% MeOH, resulting in nine fractions (A-I). Further purification steps included different types of column chromatography and/or semi-preparative HPLC-PDA and the resulting fractions were monitored by 1H NMR and/or LC-HRMS so as to evaluate the progress of the purification. For size exclusion chromatography 6.5 g of Sephadex were swollen for 3 hours to give a 20 mL column. The chromatography proceeded by gravity flow at approximately 0.55 mL/min with a 3:7 mixture of MeOH: H2O as eluent. A portion of each chromatographic fraction was dissolved at 10 mg/mL in DMSO for future use in bioassays. Each purification step led to a new set of fractions to be tested in the anti-microfouling and quorum sensing bioassays that guided the isolation until spectroscopically pure compounds were obtained. Solid Phase Extraction (SPE) was done in Strata®Silica, 5 g / 20 mL cartridges and fraction monitoring by Thin Layer Chromatography (TLC) aluminum sheets in silica gel 60, F254. HPLC fractionation was carried out using a Waters Alliance e2695 HPLC coupled with a Photo Diode Array (PDA) 2998 equipped with an Aeris Peptide C18 chromatographic column from Phenomenex (4.6 mm by 150 mm; particle size, 3.6 µm). Empower 2 Chromatography Data Software was used for reporting peak information. All solvents used were HPLC gradient grade or LC-MS grade for HPLC-PDA and LC-MS fractionation/analysis and ACS grade for extraction, VLC, SPE and TLC. LC-HRMS analysis of fractions from LEGE 11248 were acquired in an Ultimate 3000 HPLC (Thermo Scientific, Germany) monitored at 230 and 254 nm. The system, coupled with a Q Exactive Focus Orbitrap spectrometer and controlled by Xcalibur 2.1.0, operated in switch ion mode. The chromatography column used was an Aeris Peptide C18 (Phenomenex; 4.6 mm by 150 mm; particle size 3.6µm). Capillary voltage of source ESI was set at 3.8 KV and the tube lens voltage 50 V. The capillary temperature was 320⁰C. Sheath gas and auxiliary gas flow rate were at 35 and 10. Full scan was done from 200 to 3000 m/z. 1H NMR data was acquired

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on a 400 MHz Bruker Avance III. The samples were dissolved in deuterated water, methanol or chloroform purchased from Alfa Aesar and BDH Prolabo (VWR).

5.2.3 Anti-microfouling bioassays

5.2.3.1 Bioassay with fouling marine bacteria

Five known marine fouling bacteria strains were purchased from the Spanish Type Culture Collection (CECT): Cobetia marina CECT 4278, Vibrio harveyi CECT 525, Roseobacter litoralis CECT 5395, Halomonas aquamarina CECT 5000, Pseudoalteromonas atlantica CECT 570. The bacteria were inoculated in Marine Broth (MB) medium (Difco) at an initial density of 0.1 (OD600) and grown in 96 well flat-bottom microtiter plates (Orange Scientific) for 24 hours at a temperature of 26 ºC. Bacterial growth inhibition in the presence of cyanobacterial extracts of (3µg/mL and 30µg/mL), cyanobacterial fractions (10µg/mL) or compounds (10µg/mL) was determined in quadruplicate by reading of optical density at 600 nm using a microplate reader (Biotek Synergy HT, Vermont, USA) Marine Broth medium with ultra-pure water, and penicilin-streptomycin- neomycin stabilized solution (Sigma), negative and positive controls respectively. Statistically significant differences between treatments and control measurements were calculated using Student ’s t test with 95% confidence level.

5.2.3.2. Bioassay with fouling diatoms

For the anti-microalgal screening, four strains of benthic marine diatoms, Cylindrotheca BEA0501B sp., Halomphora sp. BEA0050, Nitzschia sp. BEA0497 and Navicula sp. BEA0055

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were purchased from the Spanish Collection of Algae (BEA-Banco Español de Algas, Las Palmas, Spain) and inoculated in f/2 medium (Sigma) with silica at an initial concentration of 2–4 × 106 cells per millilitre, and grown in 96-well flat bottom microtiter plates for 10 days at a temperature of 20 °C. Diatom inhibition in the presence of cyanobacterial extracts of (3µg/mL and 30µg/mL), cyanobacterial fractions (10µg/mL) or compounds (10µg/mL) was determined in quadruplicate quantified by the difference in cell densities among treatments. Cell densities were determined using a Neubauer counting chamber. f/2 medium with 0.1% DMSO, and cycloheximide (3.55 µM) were the negative and the positive controls, respectively. Statistically significant differences between treatments and control measurements were calculated using the Student’s t-test with a 95% confidence level.

5.2.4.1. Quorum sensing (QS) inhibition test

Standard disk–diffusion assay was used to detect anti-quorum sensing activity of the compounds by using the biomonitor strain Chromobacterium violaceum ATCC 12472 following the protocol developed by McLean et al., (2004) with slight modifications. A freshly inoculated culture of C. violaceum 12472 was grown overnight in Luria-Bertani (LB) broth with 1.2% agar. Five milliliters of molten LB agar (0.3% w/v) was inoculated with 50 mL of C. violaceum 12472 culture grown overnight in LB broth. The agar-culture solution was immediately poured over the surface of pre-warmed LB agar plates. Fifteen-microliters of the cyanobacterial extracts or fractions (10 µg/mL) to be tested was then pipetted on 6 mm sterile paper disks (Oxoid, Spain) which were placed on the solidiﬁed agar. The plates were incubated overnight at 30 ºC and examined for violacein production. Quorum sensing inhibition was detected by a colorless, opaque, but viable halo around the discs. Growth inhibition representing antibacterial activity towards C. violaceum was detected by a transparent zone/growth inhibition (Figure 1). DMSO was used as negative control and penicilin-streptomycin-neomycin stabilized solution (Sigma) was used as positive control for growth inhibition.

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Figure 1. LB agar plate representative of a quorum-sensing inhibition bioassay. The assay uses Chromobacterium violaceum ATCC 12742 and depicts both quorum-sensing inhibition with non- pigmented zones (4), and antimicrobial activities with translucent zones (1 and 3). Translucent zones show inhibition of bacterial growth, while opaque zones represent inhibition of violacein production with viable bacterial growth. (1) denotes positive control, (2) negative control, (3) crude extract from LEGE strain 91342 and (4) crude extract from LEGE strain 06014, respectively.

5.2.4.2. Quorum-sensing inhibition quantitative assay (Dose-response violacein inhibition assay)

To quantify the inhibition of quorum-sensing, the quorum sensing (QS) reporter strain Chromobacterium violaceum (ATCC 12472) was selected following the protocol of Martinelli et al., 2004 and Gemiarto et al., 2011. This strain was incubated in LB medium (Difco) with 3.25, µM, 6.5 µM and 13 µM of the tested crudes and inoculated in 96 well flat-bottom microtiter plates, while LB medium with sterilized ultra-pure water was used as negative control, and 1:100 penicilin-streptomycin-neomycin stabilized solution (Sigma P4083) was used as a positive control for the inhibition of cell growth. The cultures inoculated with the tested compounds were grown

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overnight (18 hours) at 26ºC with constant shaking of 50 rpm. After the incubation period, the compound treated culture and the controls were used for the determination of OD at 720 nm to quantify cell density, using a microplate reader (Biotek Synergy HT, Vermont, USA). Afterwards, the plates were dried for 30 minutes at 60ºC and violacein was re-solubilized by the addition of 200 μl of DMSO and the OD was read at 577 nm, to quantify violacein production. Mean OD720/ OD 577 ratios of the cultures with the tested compound were compared with the untreated cultures using Student’s unpaired T-test and used to assess the degree of violacein inhibition.

5.2.5 Antibacterial screening susceptibility assay

Two gram-positive bacterial strains (Staphylococcus aureus ATCC 25923, and Bacillus subtilis ATCC 6633), and two negative strains (Escherichia coli ATCC 25922, and Pseudomonas aeruginosa ATCC 27853) were selected to determine the antibacterial potential of the cyanobacterial crudes and fractions. These bacteria were grown in Mueller-Hinton agar medium (MH – BioKar diagnostics, France) from stock cultures, and MH plates were incubated at 37°C prior to obtain fresh cultures for each in vitro bioassay. Afterwards, bacterial pure colonies were picked from overnight cultures in MH (with a swab) and suspended in 5 mL of buffered peptone water (Oxoid, England), The turbidity of the inoculum is adjusted in order to equal a 0.5 McFarland standard and MH plates are seeded with that inoculum. Blank discs (6 mm in diameter - Oxoid, England) are placed in the inoculated plates and, then, impregnated with 15 µl of a 1mg/mL solution (in DMSO) of each tested compound (DMSO is use as a negative control). Plates are left 30 min at room temperature and then are incubated overnight at 37°C, and antibacterial activity is recorded when a halo around the disc is visible.

5.2.6 Assay with macroalgae

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5.2.6.1. Induction of algae spore release

Ulva lactuca was selected as a target organism in our project because it is a common fouling alga and it is used commonly as a fouling test species Fletcher (1989). The bioassay with this macroalgae spores was conducted according to Hattori and Shizuri (1996). Specimens of Ulva lactuca were collected from the intertidal region of Praia da Memória, Matosinhos, and fresh fertile fragments of the algae frond were washed with seawater and placed in 6 well-plates. In one treatment condition, the algae fragments were immersed in filtered seawater, and in another treatment condition the algae fragments were immersed in 10mL/l of PES medium (Provasoli enriched seawater medium). The 6-well plates were incubated at 15ºC under a 12:12 h light /dark cycle and the plates were observed daily with a magnifying lens (20X) for the production of spores. At the fourth day of incubation the algae immersed in seawater started producing spores (Figure 2b-second row), and there was no production of spores in the algae fragments in the enriched seawater medium. This response was observed by checking the release of spores from the algae fragments in an inverted microscope (Figure 2a and 2 b). A hemocytometer was used to determine the concentration of algal zoospores, and a concentration of around of 2x 105 spores mL-1 was selected for the initial inoculum of the spores.

Figure 2. a) Ulva lactuca thallus after the release of spores, in the fourth day of incubation in the wells with filtered seawater b) Ulva lactuca spores in the wells with filtered seawater.

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5.2.6.2. Bioassay with macroalgae spores

The bioassay with macroalga spores was conducted in flat-bottom 96-well plates as described by Chambers et al., 2006 with slight modifications. Wells were filled with 200 l of filtered seawater plus the tested compound and 50 μl of spore inoculum (around 1x 105 spores mL- 1) was added. DMSO (0.1%) was used as the negative control, while copper sulfate (0.15) was used as the positive control, Plates were then incubated at 15ºC for 6 days, and after the incubation period, the bottom of each well was examined for the presence of germinated spores with an inverted microscope (20x magnification). A spore was considered as germinated when the germ tube was visible (See Figure 3a). The medium was removed from each well and replaced with 10% formalin to stop further germination. Counts were made in 3 different fields of view in three different wells of the same concentration, and results expressed as a percentage germination. Wells were then washed with filtered seawater and the attached spores were counted using an inverted microscope (20× magnification).

Figure 3: a) Ulva spores after immersion in 6 days in artificial seawater with visible germination tubes; b) Ulva spores after immersion in copper sulfate without the presence of germination tubes.

5.3. Results and discussion

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5.3.1. Screening results for the bioassay with microfouling organisms and quorum-sensing inhibition

The inhibitory activity of 61 of the crude extracts at a concentration of 3 and 30 μg/mL were determined against a panel of known marine fouling bacteria (Supplementary Table 2). The results showed that 57% of the cyanobacterial extracts inhibited the growth of at least one fouling bacteria, while 35% of the extracts inhibited the growth of two of more bacteria. These results strengthen the already recognized potential of cyanobacterial compounds for antifouling applications (Dahms et al., 2006; Dobretsov et al., 2006). The inhibitory activity of the cyanobacterial extracts was also determined against well-known fouling diatoms (Supplementary Table 2), and the results demonstrated that 56% of the 61 cyanobacterial extracts inhibited the growth of at least one microalga, while 16% of the extracts inhibited the growth of two, or more of the tested microalgae. Taking in consideration these results with the assays with the marine fouling bacteria we can conclude that there is a significant potential of cyanobacterial metabolites to be employed against the growth of microfouling organisms (Burja et al., 2003; Dahms and Dobretsov, 2017). Previous research has demonstrated that Nostocarboline for instance, a metabolite isolated from the freshwater cyanobacterium Nostoc sp., and its synthetic analogs were highly algicidal and antimicrobial, indicating that they are potential antifoulants, though their effects on macrofoulers were not examined (Hender et al., 2008). Among the antifouling compounds isolated from marine microorganisms, Hong and Cho isolated two compounds belonging to the butelonide class from a seaweed epibiotic bacterium Streptomyces violacoruber SCG-09 which had activity against diatom Navicula annexa (Hong and Cho, 2013). Tan et al., 2010 isolated a series of compounds with mixed polyketide-polypeptide structures from the marine cyanobacterium Lyngbya majuscula. Among them, dolastatin and majusculamide A inhibited larval settlement of the barnacle Balanus amphitrite. A lactone with a 2-furanone ring - maculalactone A - was the most abundant secondary metabolite from a marine cyanobacterium Kyrtuthrix maculans, which showed a toxic effect on the naupliar larvae of the barnacles B. amphitrite (Tan et al., 2010). None

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of these cyanobacterial compounds, however, demonstrated activity against the main marine fouling groups (both microfouling and macrofouling). Among the 61 cyanobacterial crude extracts tested, 23 displayed quorum sensing inhibition activity, while 6 crude extracts showed inhibitory activity towards C. violaceum growth which was later confirmed by growth inhibition on 96 wells flat-bottom microtiter plates (Supplementary Table 2). These results suggest a relatively broad QS inhibition activity among cyanobacterial extracts. A previous study showed that cyanobacterial strains isolated from a hypersaline microbial mat inhibited QS-dependent production of violacein by C. violaceum CV017 (Abed et al., 2013) and the same was observed with cyanobacterial mats from hot springs (Dobretsov et al., 2011) which had strong quorum-sensing inhibition activity. A few isolated compounds have been reported from cyanobacteria with abilities to inhibit QS gene expression. Malyngolide and lyngbyoic acid isolated from Lyngbya majuscula for instance, were able to inhibit violacein production in C. violaceum (Kwan et al., 2011). Two other compounds — malyngamide C and 8- epi-malygamide isolated from L. majuscula were also able to inhibit QS phenomenon (Kwan et al., 2010). Furthermore, honaucins produced by cyanobacterium Leptolyngbya crossbyana are known to inhibit bacterial QS bioluminescence in V. harveyi BB120 (Choi et al., 2012). In general, it has been demonstrated that secondary metabolites from cyanobacteria can inhibit QS and therefore may have a potential application in controlling the growth of biofouling communities (Dobretsov et al., 2011). In the marine environment, a large number of marine bacteria (especially those that induce settlement of fouling macro-foulers) communicate with each other by QS and form biofilms, which has an influence on marine macroorganism adhesion (Antunes et al., 2018). Therefore, compounds with excellent anti-biofilm or QS inhibition activity have the potential to be used as antifoulants. However, until now there are few compounds from marine microorganisms with broad antifouling potential against both microfouling and macrofouling organisms. Some compounds isolated from cyanobacteria, like maculalactone (Brown et al., 2004) and portoamides (Antunes et al., 2019) have demonstrated broad antifouling activity against those different targets from microfouling to macrofouling organisms. Still, until recently, assays against biofilm growth or against biofilm disruption have seldom been used with antifouling compounds derived from marine organisms. The formation of marine biofilms has been increasingly recognized as an important factor in influencing fouling of marine macro-biofoulers (Salta et al., 2013; Carvalho, 2018), but the existing knowledge of the interaction between biofilms and settling

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propagules is quite limited. Marine biofilms are perhaps much more serious than macro-fouling for the marine industry and for aquaculture, yet so far, only several marine-derived antibiofilm compounds have been isolated and identified (Dahms and Dobretsov, 2017). Overall, marine cyanobacteria have been recognized as a rich source of secondary metabolites with AF properties (Burja et al., 2001; Dobretsov et al., 2013; Kwan et al., 2011). The still relatively low number of AF compounds isolated from microorganisms is primarily due to the fact that it is much easier to collect sponges, corals, seaweeds, and other macro-organisms from the natural environment (Dahms et al., 2006). The production of compounds by microalgae can be manipulated by changing the growth conditions, availability and concentration of nutrients in the culture medium, light intensity, temperature and/or pH (Volk and Furkert, 2006), and microbes offer more promising sources of bioactive compounds due to the bioactive potential of their secondary metabolites (Fusetani 2011, Wang et al., 2017).

5.3.2. Results for the assay with marine macroalgae

To optimize the macroalgae assay, it was difficult to determine the conditions which resulted in the release of algae spores in sufficient densities to conduct the attachment assay. Only one fraction was therefore selected to be tested in this bioassay, the fraction E17163I which did not to have a significant effect in the attachment of Ulva spores at an initial concentration of 10 g/mL (Figure 4). Higher concentrations should be tested to fully determine the anti-macroalgae attachment potential of these compounds. Other eventually isolated compounds in the conducted screening should also be tested in the anti-macroalgae settlement assay in the future. Regarding anti-macroalgae compounds derived from marine organisms, lobocompactol an active AF diterpene, isolated from a marine-derived actinomycete Streptomyces cinnabarinus PK209 exhibited significant AF activity against the macro-alga Ulva pertusa and the diatom Navicula

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annexa, and inhibited the growth of the fouling bacteria Pseudomonas aeruginosa KNP-5. (Cho et al., 2012). Regarding anti-macroalgae compounds of cyanobacterial origin Leucothrix mucor was found to inhibit the settlement of the spores of U. pertusa as well the attachment of the biofilm forming bacterial strains P. aeruginosa KNP-3 (Cho et al., 2012). The cyanobacteria Scytonema hofmanni produces the secondary metabolite cyanobacterin (Mason, 1986). Cyanobacterin is a strong inhibitor of eukaryotic algae and higher plants but not of photosynthetic bacteria (Gleason, 1990), but its effect on fouling macroalgae is was not determined.

U lv a s p o r e g e r m in a tio n

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Figure 4: Percentage of spore germination of Ulva lactuca spores exposed respectively to artificial seawater (negative control), copper sulfate (positive control) and fraction E17163I.

5.3.3. Bioactivity-guided isolation of antifouling compounds

Taking in consideration the primary screening results, both from the anti-microfouling assay and the QS-inhibition bioassay, combined with anti-macrofouling assays being conducted in another projects conducted in our lab resulted on the selection of 11 cyanobacterial strains with

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the most interesting potential for a broad anti-fouling control (Table 2) which were thus selected to be grown in large scale (ca. 120 L). The LEGE strains XX002, XX280, 00060, 00246, 00247, 03273, 06134, 07075, 07221, 91342 and 11428 were therefore selected for large scale growth. Of those strains, LEGE strains XX280, 00060, 00246, 00247, 03273, 07075, 07221, 91342 and 11428, were subjected to VLC fractioning for the isolation of their bioactive compounds (Table 3).

Table 2: Cyanobacterial strains selected for mass culturing and subsequent extraction and fractioning.

Anti- Anti- Quorum- LEGE Strain and origin Microphotographs bacterial diatom sensing Code assay assay inhibition assay

Anabaena sp. Finland x-002 Fresh water + + +

Planktothrix sp. XX280 Freshwater + - -

LEGE Unidentified cyanobacterium 00060 Morocco Freshwater + + +

LEGE Dolichospermum sp. 00246 Portugal Freshwater + + +

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LEGE Cuspidothrix issatschenkoi 00247 Portugal Freshwater + - -

LEGE Oscillatoria sp. Portugal 03273 Freshwater + + +

LEGE Cyanobium sp. Portugal 06134 Sabellaria sp. reef, intertidal, - - + epipsamic

LEGE 07221 Tychonema sp. Portugal Freshwater + - +

LEGE Unidentified filamentous 07075 Synechococcales + - - Portugal Brackish water LEGE Microcystis aeruginosa 91342 Portugal Fresh water + + - (non-MC producing strain)

LEGE Synechococcus sp. Portugal 11428 Marine + - +

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Table 3: Cyanobacterial strains grown at large-scale and VLC extracted

LEGE CC Code Extraction code Biomass (g) Crude extract VLC A-I fractions yield (%) Yield (%) LEGE 00060 E16156 16.3 22 89 LEGE 00247 E17159 11.2 14 76 LEGE 00246 E17162 17.4 17 98 LEGE 11428 E17163 31.6 22 80 LEGE 03273 E17166 24.1 13 96 LEGE XX280 E 17178 31.3 38 41 LEGE 07075 E 18180 14.0 27 52

The crude biomasses of LEGE 00060, LEGE 00246, LEGE 00247, LEGE 03273, LEGE 11428 were obtained by centrifugation and freeze-dried in order to be VLC (vacuum liquid chromatography) fractioned. After VLC fractioning, each of the obtained fractions were tested against the growth of marine fouling bacteria and diatoms for assay guided compound isolation of the bioactive compounds. (See Supplementary tables 3 to 9). Subsequent work focused on re- fractioning fractions with higher bioactivity through SPE, in order to conduct subsequent assays aiming at the structural elucidation of those compounds. The unidentified filamentous cyanobacterium strain LEGE 00060 (extraction code E16156) was the first to be tested for their antimicrofouling activity. Among the VLC fractions, the 16156_E fraction had a particular strong bioactive against the growth of marine bacteria growth, inhibiting also the quorum inhibition phenomenon (See supplementary table 3). This fraction was therefore further purified by normal phase gravity chromatography using a gradient from 75% hexane to 25% EtOAC, resulting in six fractions (E1-E6). The new fractions were tested against the same panel of antimicrofouling strains and two sub-fractions, E 16156_E3 and E 16156_E4, (eluted with 50% EtOAC: 50% Hexane) inhibited strongly the growth of the tested marine bacteria growth, particularly Cobetia marina and Pseudoalteromonas atlantica (Figure 5). Those same fractions also inhibited quorum-sensing phenomenon, and also inhibited the growth of the pathogenic gram-negative and gram-positive bacteria E. coli and S. aureus (See

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supplementary table 3). The 1H NMR spectrum of these fractions showed the presence of chlorophylls and/or their derivatives (characteristic resonances in the δ 8-10 regions) and of other metabolites (Supplementary figures 1 and 2), as well likely the presence of phthalate, which are plasticizers, substances added to plastic to increase their flexibility, transparency, durability, and longevity which could be responsible for the observed bioactivity. Further work would be needed to elucidate the structure of these compounds, however as both fractions E 16156_E3 and E 16156_E4 yielded masses below 10 mg, the isolation of pure compounds for structural characterization was unlikely to be achieved, and therefore these samples were not processed any further.

P s e u d o a lte r o m o n a s a tla n tic a g r o w th C o b e tia m a r in a g r o w th

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l l 3 4 l l 3 4 o o E E o o E E r r r tr t t 6 6 t 6 6 n n 5 5 n n 5 5 o o 1 1 o o 1 1 6 6 C C 6 6 C C 1 1 1 1 e e e e E E v v E iv tv E ti it t i a s a s g o g o e P e P N N Figure 5a and 5b: Antibacterial activity of fractions 16156 E3 and 16156 E4 against the growth of Cobetia marina (Figure 5a) and Pseudoalteromonas atlantica (Figure 5b). B: 0.1% DMSO; C: 1:100 dilution of penicilin-streptomycin-neomycin stabilized solution (Sigma P4083). T-test results: * p-value <0.05, ** p-value<0.01; *** p-value <0.001.

Among other tested VLC fractions, the fractions obtained from the cyanobacteria strain LEGE 00247, Cuspidothrix issatschenkoi (E17159), and particularly the fractions E 17162_A, and E 17162_B obtained from the strain LEGE 00246, a freshwater Dolichospermum sp. had activity against the growth of marine fouling bacteria, as well against diatoms (See Figure 6 and supplementary table 5). Another cyanobacterial strain, the Oscillatoria sp. strain LEGE 03273

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(E17666), also produced several bioactive fractions, particularly the fractions E17166_D, E17166_F and E17166_G (See Figure 7 and supplementary table 7).

V ib r io h a r v e y i g r o w th H a lo m o n a s a q u a m a r in a g r o w th

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l l A l l A o o o o r r 2 r r 2 t t 6 t t 6 n n 1 n n 1 o o 7 o o 7 C C 1 1 C C e e E e e E iv tv v v t i ti it a s a s g o g o e P e P N N

P s e u d o a lte r o m o n a s a tla n tic a g r o w th R o s e o b a c te r lito r a lis g r o w th

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l l D l l D o o o o r r 6 r r 6 t t 6 t t 6 n n 1 n n 1 o o 7 o o 7 1 C 1 C C C e e E e e E v v iv tv ti it t i a s a s g o g o e P e P N N

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Furthermore, the fraction E17163_I, obtained from the strain LEGE11428, a marine Synechococcus sp. was particularly active against bacteria and diatoms, and also the against pathogenic bacteria E. coli, S. aureus and Salmonella. (See Figure 8 and supplementary table 8). All of these fractions were analyzed by 1H NMR (Supplementary figures 3 to 8).

N a v ic u la s p . g r o w th C y lin d r o th e c a s p . g r o w th

1 5 0

o w

r *** **

o *** * 1 0 0

g r

1 0 0

a i

D 5 0

% %

0 0

l l I l l I o o o o r r 3 r r 3 t t 6 t t 6 n n 1 n n 1 o o 7 o o 7 1 C C 1 C C E E e e e e v v iv tv ti it t i a s a s g o g o e P e P N N Figure 8a and 8b: Anti-diatom activity of fraction 17163 I against the growth of Cylindtrotheca sp. (Figure 8a) and Navicula sp. (Figure 8b). B: 0.1% DMSO. Cycloheximide (3.55 µM) was used as the positive control (C+). T-test results: * p-value <0.05, ** p-value<0.01; *** p-value <0.001.

Fraction E17163_I was selected to proceed other chromatographic purification methods due to the following reasons: it formed a white crystal a large mass was obtained (~1.8 grams, 25%) and its 1H NMR spectrum contained diagnostic chemical shifts characteristic of the functional group amino (characteristic exchangeable resonances in the δ 8-8.5 regions), the presence of aromatic groups (characteristic resonances in the δ 6.5-7.5 regions) and probable sugar protons (characteristic resonances in the δ 3.5-4 regions), as well as probable acetyl groups

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(characteristic resonances in the δ 3.5-4 regions δ 2.0 to δ 2.1), anomeric protons (characteristic resonances in the δ 3.5-4 regions δ 4.4 to δ 5.5) (Figure S6). The fraction E 17163_I was then submitted to normal phase SPE following a stepwise gradient of Methanol: H2O (10%-75% MeOH) and EtoAC: MeOH (10%-50% EtoAC), and 15 fractions were collected. Those fractions were concentrated by rotavapor and pooled according to their TLC (Silica Gel TLC) profiles, which reduced the number of fractions to nine i.e. E17163_I_1-9. Those fractions were submitted to the bacteria and diatom assays as well to other parallel macrofouling assays. As the most interesting bioactivity was found on the fraction I_7, with the combination of the NMR profile spectra, this fraction was chosen to be further separated in a Sephadex LH-20 column (MeOH 35% at 0.5 mL/min). 10 sub-fractions were therefore obtained: I_7_A-J, and the 1H NMR showed the cleaner Sephadex fraction, I_7_D, which was further purified by reversed phase HPLC-PDA (Figure 9.).

2.00 1.659

1.50 2.082

AU 1.00 8.801

0.50

5.353

8.546

7.729 9.705

0.00

2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 10.00 Minutes

300.00

250.00 nm

200.00

2.00 3.00 4.00 5.00 6.00 7.00 8.00 9.00 Minutes

Figure 9. HPLC-PDA Chromatogram and PDA contour of E17163_I7_D.

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The fraction E17163_I7 (1.6 g, 73%) that was also purified by Sephadex LH-20 resulting in 11 fractions. Fraction E17163_I7_D (237 mg, 35%) formed a white crystalline solid after being evaporated, however, it became amorphous when a later crystallography analysis attempt was made. This fraction had strong activity against all the tested biofilm-forming marine bacteria. This fraction was thus further purified by HPLC-PDA (Figure 5). Chromatographic purification resulted in a total of nine fractions (E17163_I7_D (1-9) with a very low yield of 17%. The entire HPLC run and additional washing eluent were collected what made us suppose that a large amount of the injected fraction was not 100 % soluble or some compounds bonded irreversibly with the chromatographic sorbent column. Nevertheless, we were able to obtain a much less convoluted 1H NMR (Figure S8) spectra of the HPLC fraction E17163_I7_4_D6 (~0.42 mg, 10%) which was also analyzed by NMR (Supplementary figure 6). Both spectra indicate the presence of an aromatic moiety substituted in a para fashion (characteristic double duplet resonances in the δ 6.5-7.5 regions) and showing still the characteristic signal for an amino group, which could indicate the presence of a tyrosine residue. Overall, the 1H NMR and HRMS data signatures for the major compounds obtained from E17163I sub-fractions seem to be consistent with either glysosidic and peptidic moieties, or both (e.g. glycopeptides). Various factors, such as the presence of compounds labile to easy degradation or the very small amount of fraction to continue purification did not allow the so wanted full structure elucidation of some of the bioactive fractions. For all those reasons we will pursue in the future the purification and structure elucidation of these compounds from Synechococcus sp. LEGE 11248.

5.4. Main conclusions

The bioassay-guided screening of the tested LEGE CC cyanobacterial strains demonstrated the strong antifouling potential of cyanobacterial crudes against the growth of microfouling species associated to marine biofilms, as well to inhibit the quorum-sensing phenomenon. In general, it

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was shown that cyanobacterial natural products can be a promising strategy for the control of marine biofilm growth and against marine biofouling species in general. In particular, crudes from strains LEGE 00060, LEGE 00246, LEGE 00247, LEGE 11248 were particularly bioactive and their VLC fractions further inhibited the panel of microfouling species. A particularly promising fraction was obtained from the strain LEGE 11428, which seem to be consistent with glysosidic and peptidic moieties, or both (e.g. glycopeptides). The very small amount of this fraction did not allow the so wanted for further purification and full structure elucidation of this fraction, which will be however be continued in the future.

5.5 References

Abed, R.,Dobretsov, S., Marwan, A.F., Sarath, P.G., Sudesh, K., Paul, V.J. 2013. Quorum- Sensing Inhibitory Compounds from Extremophilic Microorganisms Isolated from a Hypersaline Cyanobacterial Mat. Journal of Industrial Microbiology and Biotechnology 40: 759–72.

Almeida, J.R., Vasconcelos, V. 2015. Natural antifouling compounds: effectiveness in preventing invertebrate settlement and adhesion. Biotechnol Adv. 33: 343-57.

Antunes, J.T., Leão, P.N., Vasconcelos, V.M. 2018. Marine biofilms: diversity of communities and of chemical cues. Environmental Microbiology Reports. url: https://doi.org/10.1111/1758- 2229.12694.

Antunes, J., Pereira, S., Ribeiro T., Plowman J.E., Thomas, A., Clerens, S., Campos A., Vasconcelos V., Almeida, J.R. 2019. A multi-bioassay integrated approach to assess the antifouling potential of the cyanobacterial metabolites portoamides. Mar. Drugs 17(2): 111.

Barberousse,H., Lombardo, R.J., Tell, G., Coute, A. 2006. Factors involved in the colonisation of building facades by algae and cyanobacteria in France. Biofouling 22:79 – 90.

Bhadury,P., Wright, P.C. 2004. Exploitation of marine algae: biogenic compounds for potential antifouling applications. Planta 219: 561–578.

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Brown, G., Wong, H., Hutchinson, N., Lee, S., Chan, B., Williams, G. 2004. Chemistry and biology of maculalactone A from the marine cyanobacterium Kyrtuthrix maculans. Phytochem Rev 3:381–400.

Burja, A.M., Banaigs, B., Abou-Mansour, E., Burgess, J.G., Wright, P.C. 2001 Marine cyanobacteria - a prolific source of natural products. Tetrahedron 57(46): 9347-77.

De Carvalho, C.C.C.R. 2018. Marine biofilms: A successful microbial strategy with economic implications. Front. Mar. Sci. 5: 126.

Cho, J.Y., Kang, J.Y., Hong, Y.K., Baek, H.H., Shin, H.W., Kim, MS. 2012. Isolation and structural determination of the antifouling diketopiperazines from marine-derived Streptomyces praecox 291-11. Biosci Biotechnol Biochem. 76: 1116–1121.

Dahms, H.U., Xu, Y., Pfeiﬀer, C. 2006. Antifouling potential of cyanobacteria. Biofouling 22:317 327.

Dahms, H.U.; Dobretsov, S. 2017. Antifouling Compounds from Marine Macroalgae. Mar Drugs 15(9):265.

Dobretsov, S., Teplitski, M., Bayer, M., Gunasekra, S., Proksch, P., Paul, V.J. 2011. Inhibition of marine biofouling by bacterial quorum sensing inhibitors. Biofouling. 27:893–905.

Dobretsov, S., Abed; R.M.M., Teplitski, M. 2013.Mini-review: Inhibition of biofouling by marine microorganisms. Biofouling. 29(4): 423–441.

Edwards, D. J., Marquez, B.L., Nogle, L.M., McPhail, K., Goeger, D.E., Roberts, M.A, Gerwick, W.H. 2004. Structure and biosynthesis of the jamaicamides, new mixed polyketide-peptide neurotoxins from the marine cyanobacterium Lyngbya majuscula. Chem Biol. 11: 817-833.

Fletcher, R.L. 1989. A bioassay technique using the marine fouling green alga Enteromorpha. Int Biodeterior. 25: 407–422.

Freitas, S., Martins, R., Costa, M., Leão, P, Vitorino, R, Vasconcelos, V. 2016. Hierridin B Isolated from a Marine Cyanobacterium Alters VDAC1, Mitochondrial Activity, and Cell Cycle Genes on HT-29 Colon Adenocarcinoma Cells. Mar Drugs 214: 158.

Fusetani, N. 2004. Biofouling and antifouling. Nat Prod Rep 21:94–104.

Fusetani, N. 2011 Antifouling marine natural products. Nat Prod Rep 28: 400–410.

Gemiarto, A., Ninyo, N.N., Lee, W.L., Logis, J., Fatima, A., Chan, E.W.C., Lim, C.S.Y. 2015 Isoprenyl caffeate, a major compound in manuka propolis, is a quorum-sensing inhibitor in Chromobacterium violaceum. Antonie Van Leeuwenhoek. 108: 491–504.

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Gleason, F.K., Case, D.E., Sipprell, K.D., Magnuson, T.S. 1986. Effect of the natural algicide cyanobacterin on a herbicide-resistant mutant of Anacystis nidulans R2. Plant Sci 46:5 – 10.

Hattori, T., Shizuri, Y., 1996. A screening method for antifouling substances using spores of the fouling macroalga Ulva conglobata Kjelleman. Fisheries Science 62: 955e-58.

Hellio, C., Berge, J.P., Beaupoil, C., Le Gal, Y., Bourgougnon, N. 2002.Screening of marine algal extracts for anti-settlement activities against microalgae and macroalgae. Biofouling18: 205-215.

E. Hender, M. Sjogren, S., Hodzic, R., Andersson, U., Goransson, P. R., Jonsson and L. Bohlin, J.2008. Antifouling activity of a dibrominated cyclopeptide from a marine sponge Geodia barretti. Nat. Prod. 71: 330–333.

Hong, Y.K., Cho, J.Y. 2013. Effect of seaweed epibiotic bacterium Streptomyces violaceoruber SCH-09 on marine fouling organisms. Fish Sci. 79:469–475.

Jelic-Mrcelic, G., Sliskovic, M., Antolic, B. 2006. Biofouling communities on test panels coated with TBT TBT-free copper based antifouling paints. Biofouling. 22:293–302.

Kotai, J. 1972. Instructions for the preparation of modified nutrient solution Z8 for algae. Oslo: Norwegian Institute for Water Research Blindern NIVA B-11/69.

Kwan, J.C., Meickle, T., Ladwa, D., Teplitski, M., Paul, V., Luesch, H. 2011 Lyngbyoic acid, a “tagged” fatty acid from marine cyanobacterium, disrupts quorum sensing in Pseudomonas aeruginosa. Mol Biosyst. 7: 1205–16.

Kwan JC, Teplitski M, Gunasekara SP, Paul VJ, Luesch H. 2010. Isolation and biological evaluation of 8-epi-malyngamide C from the Floridian marine cyanobacterium Lyngbya majuscula. J Nat Prod. 73:463–6.

Leao, P.N., Costa, M., Ramos, V.; Pereira, A.R.; Fernandes, V.C., Domingues, V.F., Gerwick, W.H., Vasconcelos, V.M., Martins, R. 2013. Antitumor activity of hierridin B, a cyanobacterial secondary metabolite found in both filamentous and unicellular marine strains. PLoS One 8 e69562

Martinelli, D., Grossmann, G., Sequin, U., Brandl, H., Bachofen, R. 2004.Effects of natural and chemically synthesized furanones on quorum sensing in Chromobacterium violaceum. BMC Microbiol. 4: 25.

Mason, C.P., Edwards, K.R., Carlson, R.E., Pignatello, J., Gleason, F.K., Wood, J.M. 1982. Isolation of chlorine containing antibiotic from the freshwater cyanobacterium Scytonema hofmani. Science 215:400 – 402.

McLean, R.J., Pierson, L.S., III; Fuqua, C. 2004. A Simple screening protocol for the identification of quorum signal antagonists. J. Microbiol. Methods 58: 351–360.

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Qian, P.Y., Chen, L., Xu, Y. 2013. Mini-Review: Molecular mechanisms of antifouling compounds. Biofouling 29: 381–400.

Ramos, V., Morais, J., Castelo-Branco, R., Pinheiro, A., Martins, J., Regueiras, et al. 2018. Cyanobacterial diversity held in microbial biological resource centers as a biotechnological asset: the case study of the newly established LEGE culture collection. J Appl Phycol. 30:1437–51.

Ribeiro, T., Lemos, F., Preto, M., Azevedo, J., Sousa, M.L., Leao, P.N., Vitorino, R., Vasconcelos, V., Urbatzka, R. 2017. Cytotoxicity of portoamides in human cancer cells and analysis of the molecular mechanisms of action. PloS ONE 12 (12): e018817.

Salta, M., Wharton, J.A., Blache, Y., Stokes, K.R., Briand J-F. 2013. Marine bioﬁlms on artiﬁcial surfaces: structure and dynamics. Environ Microbiol. 15:2879

Schultz, M.P. 2007. Effects of coating roughness and biofouling on ship resistance and powering. Biofouling. 23: 331–41.

Tan, L.T., Goh, B.P.L. 2010. Chemical ecology of marine cyanobacterial secondary metabolites: a mini review. J Coastal Dev. 13:1–9

Tan, L.T., Goh, BPL, Tripathi, A., Lim, M.G., Dickinson, G.H., Lee, S.S.C., Teo, S.L.M. 2010. Natural antifoulants from the marine cyanobacterium Lyngbya majuscula. Biofouling. 26:685– 695.

Volk, R-B., Furkert, F.H. 2006. Antialgal, antibacterial and antifungal activity of two metabolites produced and excreted by cyanobacteria during growth. Microbiol Res. 161:180–186.

Wang, K.L., Wu, Z.H., Wang, Y., Wang, C.Y.; Xu, Y. 2017. Mini-Review: Antifouling natural products from marine microorganisms and their synthetic analogs. Mar Drugs. 15: 266.

Whitton, B. A. & Potts, M. (2000) in The Ecology of Cyanobacteria, eds. Whitton, B. A. & Potts, M. (Kluwer, Dordrecht, The Netherlands), pp. 1–11.

Yebra, D.M., Søren K., Dam-Johansen, K. 2004. Antifouling Technology - Past, present and future steps towards efficient and environmentally friendly antifouling coatings. Progress in Organic Coatings 50: 75–104.

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6- Main Conclusions

The importance of marine biofilms in the marine fouling process was critically discussed along this PhD thesis. The diversity of the marine biofilm communities was discussed, as well the influence of environmental parameters involved in their formation (Chapter 2). The description of chemical cues, particularly quorum sensing (QS), involved in biofilm formation or involved in the relation with macrofouling settlement was also critically reviewed. Lastly, since the formation of a marine biofilm is considered to be an initial, QS-dependent step in the development of marine fouling events, QS inhibition is discussed on its potential as a tool for anti-biofouling control in marine settings. The analysis of 16S rRNA amplicon sequencing data demonstrated that the biofilm community structure succession and their functional profile significantly changed in the sampled seasons (Chapter 3). The marine biofilm communities seem therefore to experience strong temporal variations, which have been suggested by previous authors but has not been thoroughly analyzed in previous studies. In specific, the dominant biofilm taxa sampled during spring and during winter were distinct, particularly in the first sampled days (day 1 to day 4), and also during the last sampled day (day 30). During the day 30 of biofilm growth in spring there is already a strong dominance (more than 70%) of autotrophic organisms (mostly diatoms), while during winter those group of organisms were not abundant (less than 5%). This makes us conclude that the marine biofilms in warmer seasons stabilize and mature significantly earlier than biofilms sampled in cold waters, a conclusion obtained also from the functional analysis of the marine biofilm communities. Previously unreported pathogenic taxa in biofilm was detected in some of the samples, and their presence seems to be strongly correlated to temporal variations. These findings lend further support to the role of marine biofilms as vectors of harmful taxa, particularly in big ports with access to effluents. Additionally, taxa associated to the corrosion of the steel surfaces was also detected among the pioneer colonizing bacteria proving that those taxa are present in the steel surfaces since a very early period. Biofilm communities growing in steel was significantly distinct both from bacterial community present in the seawater, and from the one

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growing in steel surfaces covered with anti-corrosion paint. Overall, we suggest that the observed temporal succession of early marine biofilm communities should be taken in consideration for future antifouling applications, as well for the effective control of pathogenic and steel-corroding associated bacterial taxa. Those strategies should take in consideration that pioneer colonizing bacteria of artificial surfaces may change along the season of the year and that the transition from dominant heterotrophic to autotrophic groups and the shift towards macrofouling settlement may have occur in very different temporal settings. This in turn may be responses to broader temporal cycles. In Chapter 4 we determined the potential of the cyanobacterial metabolites portoamides as potential new eco-friendly antifouling agents with comprehensive bioactivity. This was demonstrated by their high effectiveness in eradicating the growth of marine bacteria and diatom growth and also by their disrupting potential of biofilms of marine fouling bacteria. In order to test the anti-biofilm assay, we employed the MBEC assay which was not previously used with marine biofilm forming bacteria and we argue that this assay should be used in the future to effectively screen bioactive compounds against attached biofilm disruption. Previous studies testing new antifouling compounds rarely test those compounds in anti-biofilm assays which are crucial because that is the initial stage of marine fouling and bacteria in the biofilm stage has a considerable different physiology and resistance compared to bacteria in the planktonic stage. The antifouling potential of the cyanobacterial metabolites portoamides was further strengthened in the employed multi-bioassay approach, as they demonstrated activity against macroinvertebrate settlement (Mytilus sp.) while displaying no apparent toxicity against target and non-target organisms. The study of the antifouling molecular targets of portoamides in mussel larvae demonstrated energy metabolism modifications (failure in proton-transporting ATPases activity), as well structural alterations of the gills and protein and gene regulatory mechanisms. Portoamides showed a broad- spectrum bioactivity towards diverse biofouling species, including a non-toxic and reversible effect towards mussel larvae, showing their potential to be incorporated as active ingredients in antifouling coatings with no apparent ecotoxicity. However, portoamides should be further tested against marine organisms of other trophic groups to fully infer about their eventual toxicity. The observed AF activity of portoamides in this work also suggests that their mode of action is dictated by different mechanisms depending on the target groups, and the combination of functionalities of these natural products may serve as an inspiration for the development of new AF agents. This is

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particularly important as the elucidation of AF mechanisms are nowadays considered to be a prerequisite for the registration and commercialization of any new antifoulant. Chapter 5 demonstrated the strong potential of the tested cyanobacterial bioactive extracts for microfouling control, anti-biofilm and for quorum sensing inhibition, proving the potential of cyanobacteria as a source of these products. Following the results of the primary screening, five promising cyanobacterial bioactive strains were selected to be grown in higher scale, in order to obtain higher amount of crude biomass to be further VLC fractionated: the unidentified strain LEGE 00060, Cuspidothrix issatschenkoi strain LEGE 00247, Dolichospermum sp. strain LEGE 00246, Synechococcus sp. strain LEGE 11248. The bioassay-guided isolation showed that LEGE 11248 was the most active strain and was therefore selected for the attempt of the isolation and identification of bioactive compounds responsible for antifouling activity and quorum sensing inhibition activity. This resulted in a strongly bioactive sub-fraction originating from this marine cyanobacterium’s crude extract, which will be the target of further studies aimed at revealing the identity of the bioactive constituent, which was not yet possible due to an apparent low stability of molecule(s). Overall, this PhD project strengthens the importance of characterizing marine biofilms in different environmental settings and along different time periods, as biofilm community diversity and functionality may vary strongly along with time. Besides the ecological relevance of these findings, this knowledge should have importance for an efficient antifouling strategy that can control biofouling formation by acting efficiently against the main taxa of early marine biofilms in different settings. At the same time, the antifouling screening and the anti-biofilm assay here developed, further demonstrates the potential of marine natural products from cyanobacteria, as an alternative, “cleaner” strategy to control biofilm growth, and subsequently to control marine fouling development from an early period

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7- References

Abarzua, S., Jakubowski, S., Eckert, S., Fuchs, P. 1999. Biotechnological investigation for the prevention of marine biofouling. II. Blue-green algae as potential producers of biogenic agents for the growth inhibition of microfouling organisms. Bot Mar. 42: 459–465.

Antunes, J., Leão, P., Vasconcelos, V. 2018. Marine biofilms: Diversity of communities and of chemical cues. Environ. Microbiol, Rep. doi:10.1111/1758-2229.12694.

Antunes, J., Pereira, S., Ribeiro, T., Plowman J.E., Thomas, A., Clerens, S., Campos A., Vasconcelos V., Almeida, J.R. 2019. A multi-bioassay integrated approach to assess the antifouling potential of the cyanobacterial metabolites portoamides. Mar. Drugs. 17(2): 111.

Barberousse, H., Lombardo, R.J., Tell, G, Coute, A. 2006. Factors involved in the colonization of building facades by algae and cyanobacteria. Biofouling. 22(1-2): 69-77.

Battin, T. J., Sloan, W. T., Kjelleberg, S., Daims, H., Head, I. M., Curtis, T. P., 2007. Microbial landscapes: new paths to bioﬁlm research. Nat. Rev. Microbiol. 5: 76–81.

Bhadury, P., Wright, P. C. 2004. Exploitation of marine algae: biogenic compounds for potential antifouling applications. Planta 219: 561–578.

Besemer, K., Peter, H., Logue, J.B., Langenheder, S., Lindstorm, E.S., Tranvik, L.J. 2012. Unraveling assembly of stream biofilm communities. ISME J. 6: 1459–6.

Blomster, J., Hoey, E.M., Maggs, C.A., Stanhope, M.J. 2000. Species- specific oligonucleotide probes for macroalgae: molecular discrimination of two marine fouling species of Enteromorpha (Chlorophyceae). Molecular Ecology 9: 177–186.

Bossenbroek, J.M., Kraft, C.E., Nekola, J.C.2001. Prediction of long- distance dispersal using gravity models: Zebra mussel invasion of inland lakes. Ecological Applications 11: 1778–1788.

Boudouresque, C.F., Verlaque, M. 2002. Assessing scale and impact of ship-transported alien macrophytes in the Mediterranean Sea. In: Briand F (ed) Alien marine organisms introduced by ships in the Mediterranean and Black seas. Workshop Monographs n8 20. CIESM, Monaco, pp 53–63.

Brady, R. F., Singer, I. L. 2000. Mechanical factors favoring release from fouling release coatings. Biofouling 15: 73–81.

FCUP Chapter 7 179

Bressy, C., Lejars, M. 2014. Marine fouling: an overview. J Ocean Technol. 9:19–28.

Briand, J.F., Jacquet, S., Bernard, C., Humbert, J.F., 2003. Health hazards for terrestrial vertebrates from toxic cyanobacteria in surface water ecosystems. Vet. Res. 34: 1–17.

Briand, J.F. 2009. Marine Antifouling Laboratory Bioassays: An Overview of Their Diversity. Biofouling 25: 297–311.

Briand J.F., Djeridi, I., Jamet, D., Coupé, S., Bressy, C., Molmeret, M., Le Berre, B., Rimet F, Bouchez A, Blache Y, 2012, Pioneer marine biofilms on artificial surfaces including antifouling coatings immersed in two contrasting French Mediterranean coast sites. Biofouling 28: 453–463.

Briand, J.F., Barani, A., Garnier, C., Réhel, K., Urvois, F., LePoupon, C., Bouchez, A, Debroas, D., Bressy, C. 2017. Spatio-Temporal Variations of Marine Biofilm Communities Colonizing Artificial Substrata Including Antifouling Coatings in Contrasted French Coastal Environments. Microb Ecol. 74:585-598.

Brian-Jaisson, F., Ortalo-Magné, A., Guentas-Dombrowsky, L., Armougom, F., Blache,Y. and Molmeret, M. 2014. Identification of Bacterial Strains Isolated from the Mediterranean Sea Exhibiting Different Abilities of Biofilm Formation. Microb Ecology. 68: 94–110.

Burgess, J.G., Boyd, K.G., Armstrong, E., Jiang, Z., Yan, L.M., Berggren, M., May, U., Pisacane, T., Granmo, A., Adams, D.R., 2003. The development of a marine natural product-based antifouling paint. Biofouling 19: 197–205.

Callow, M. E., Callow, J. A., Pickett-Heaps, J. D., Wetherbee, R. 1997 Primary adhesion of Enteromorpha (Chlorophyta, Ulvales) propagules: quantitative settlement studies and video microscopy. J. Phycol. 33: 938–947.

Callow, J.A., Callow, M.E. 2011. Trends in the development of environmentally friendly fouling- resistant marine coatings. Nat Commun. 2:244.

Camps, M., Barani,A. Gregori,G. Bouchez, A., le Berre, B., Bressy, C., Blache, Y., Briand. J.F. 2014. Antifouling Coatings Influence Both Abundance and Community Structure of Colonizing Biofilms: A Case Study in the Northwestern Mediterranean Sea.. Appl. Environ. Microbiol 80: 4821–31.

Cao, Y., He, S., Zhou, Z. Zhang, Z., Mao, Q., Zhang, H. 2012. Orally Administered Thermostable N-Acyl Homoserine Lactonase from Bacillus sp. Strain AI96 attenuates Aeromonas hydrophila Infection in Zebrafish, Appl. Environ. Microbiol. 78:1899–1908.

Cassé, F., Swain, G.W. 2006. The development of microfouling on four commercial antifouling coatings under static and dynamic immersion. Int Biodeterior Biodegradation 53: 179–185.

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Chambers, L.D., Wharton, J.A., Wood, R.J.K., Walsh, F.C., Stokes, K.R. 2014 Techniques for the measurement of natural product incorporation into an antifouling coating. Prog Org Coat. 77: 473–484.

Cresswell, T, Richards, J.P., Gleg, G.A., Readman, J.W. 2006. The impact of legislation on the usage and environmental concentrations of Irgarol 1051 in UK coastal waters. Mar. Pollut. Bull. 52:1169–1175.

Dafforn KA, Lewis JA, Johnston EL. 2011. Antifouling strategies: history and regulation, ecological impacts and mitigation. Mar Pollut Bull. 62: 453 – 465.

De Carvalho, C.C.C.R. 2018. Marine biofilms: A successful microbial strategy with economic implications. Front Mar Sci. 5: 126.

Chen, L., Qian, P-Y. 2017. Review on molecular mechanisms of antifouling compounds: An update since 2012. Mar. Drugs. 15: 264.

Choi, H., Mascuch, S.J., Villa, F.A., Byrum, T., Teasdale, M.E., Smith, J.E., et al. 2012 Potent inhibitors of inflammation and bacterial quorum sensing: synthetic derivatives and structure– activity relationships. Chem Bio. 19: 589–98.

Chiu, J., Zhang, R., Wang, H., Thiyagarajan, V., and Qian, P. 2008. Nutrient effects on intertidal community: from bacteria to invertebrates. Mar Ecol Prog Ser. 358: 41–50.

Costas, E., Gonzalez, R., López-Rodas, V., Emma Huertas, I. 2013. Mutation of microalgae from antifouling sensitivity to antifouling resistance allows phytoplankton dispersal through ships Biofouling. Biological Invasions 15: 1739–50.

Dahms, H.U., Ying, X.; Pfeiffer C. 2006. Antifouling potential of cyanobacteria: A mini-review. Biofouling 22: 317–327.

Dahms, H.U., Dobretsov, S. 2017.Antifouling Compounds from Marine Macroalgae. Mar Drugs15(9): 265.

Dang, H., Lovell, C. R. 2016 Microbial surface colonization and biofilm development in marine environments. Microbiol Mo Bio Rev. 80: 91–138.

Davies, D.G., Parsek, M.R., Pearson, J.P., Iglewski, B.H., Costerton, J.W., Greenberg, E.P. 1998 The involvement of cell-to-cell signals in the development of a bacterial biofilm. Science 280: 295–8.

FCUP Chapter 7 181

Dobretsov, S., Dahms, H.U., Qian, PY 2006. Inhibition of biofouling by marine microorganisms and Their Metabolites. Biofouling 22 (1): 43–54.

Dobretsov, S., Dahms, H.U., Yili, H., Wahl, M., Qian, P.Y. 2007. The effect of quorum-sensing blockers on the formation of marine microbial communities and larval attachment. FEMS Microbiol Ecol. 60:177–188.

Dobretsov, S., Teplitski, M., Paul, C. 2009 Mini-Review: Quorum-sensing in the marine environment and its relationship to biofouling. Biofouling 25: 413–27.

Dobretsov, S., Teplitski, M., Bayer, M., Gunasekra, S., Proksch, P., Paul, V.J. 2011. Inhibition of marine biofouling by bacterial quorum sensing inhibitors. Biofouling 27:893–905.

Dobretsov, S., Abed, R.M.M., Teplitski, M. 2013. Mini-review: Inhibition of biofouling by marine microorganisms. Biofouling 29: 423–441.

Dong, Y.H., Wang, L.H., Xu, J.L. et al. 2001. Quenching quorum-sensing-dependent bacterial infection by an N-acyl homoserine lactonase. Nature 411: 813-817.

Dürr, S., and Thomason, J. C. 2010. Biofouling. Oxford: Blackwell Publishing Ltd.

Fusetani, N. 2004. Biofouling and antifouling. Nat Prod Rep. 2: 94–104.

Fusetani, N. 2011 Antifouling marine natural products. Nat Prod Rep. 28: 400–410.

Hall-Stoodley, L., Costerton, J. William, Stoodley, Paul. 2004 Bacterial bioﬁlms: from the natural environment to infectious diseases. Nat. Rev. Microbiol. 2: 95–108.

Hellio C, De La Broise D, Dufosse L, Le Gal Y, Bourgougnon, N. 2001, Inhibition of marine bacteria by extracts of macroalgae: potential use for environmentally friendly antifouling paints. Mar Environ Res. 52: 231-247.

Hellio, C., Berge, J.P., Beaupoil, C., Le Gal, Y., Bourgougnon, N. 2002. Screening of marine algal extracts for anti-settlement activities against microalgae and macroalgae. Biofouling 18: 2052-15.

Holm, E. R., Schultz, M. P., Haslbeck, E. G., Talbott, W. J., Field, A. J. 2004 Evaluation of hydrodynamic drag on experimental fouling-release surfaces, using rotating disks. Biofouling 20: 219–226.

Howell, D., Behrends, B. 2006. A review of surface roughness in antifouling coatings illustrating the importance of cutoff length. Biofouling 22:401 – 410.

FCUP Chapter 7 182

Huggett, M.J., Nedved B.T., Hadfield M.G. 2009. Effects of initial surface wettability on biofilm formation and subsequent settlement of Hydroides elegans. Biofouling 25:387–399.

Hunsucker, K.Z., Vora, G.J., Hunsucker, J.T., Gardner, H., Leary, D.H., Kim, S., Lin, B., Swain, G. 2018. Biofilm community structure and the associated drag penalties of a groomed fouling release ship hull coating. Biofouling 34:162–172.

Jelic-Mrcelic, G., Sliskovic, M., Antolic, B. 2006. Biofouling communities on test panels coated with TBT free copper based antifouling paints. Biofouling 22:293–302.

Flemming, H.C., Wingender, J. 2010. The Biofilm Matrix. Nature Review Microbiology 8 (9): 623–33.

Freitas, S., Martins, R., Costa, M., Leão, P., Vitorino, R., Vasconcelos, V., et al. 2016 Hierridin B Isolated from a Marine Cyanobacterium Alters VDAC1, Mitochondrial Activity, and Cell Cycle Genes on HT-29 Colon Adenocarcinoma Cells. Mar Drugs. 14: 158.

Fuqua, C., Parsek, M. R., Greenberg, E. P. 2001. Regulation of gene expression by cell-to-cell communication: acyl-homoserine lactone quorum sensing. Annu Rev Genet. 35: 439–468.

Givskov, M., de Nys, R., Manefield, M., Gram, L., Maximilien, R., Eberl, L., Molin, S., Steinberg, P. & Kjelleberg, S. 1996 Eukaryotic interference with homoserine lactone-mediated prokaryotic signalling. J. Bacteriol. 178: 6618–6622.

Gollasch, S. 2002. The importance of ship hull fouling as a vector of species introduction in the North Sea. Biofouling 18: 105–121.

Hattori, T., Shizuri, Y., 1996. A screening method for antifouling substances using spores of the fouling macroalga Ulva conglobata Kjelleman. Fisheries Science 62: 955-958.

Hellio, C. and Yebra, D. Advances in marine antifouling coatings and technology, Woodhead Publishing Limited, Cambridge, 2009.

Hellio, C., Berge, J.P., Beaupoil, C., Le Gal, Y., Bourgougnon, N. 2002.Screening of marine algal extracts for anti-settlement activities against microalgae and macroalgae. Biofouling 18:2052-15.

Hayden, H.S., Blomster, J., Maggs, C. A., Silva, P.C., Stanhope, M.J., Waaland, JR. 2003. Linnaeus was right all along: Ulva and Enteromorpha are not distinct genera. Eur J Phycol. 38: 277 – 294.

Joint, I., Callow, M.E., Callow, J.A., and Clarke, K.R. 2000. The attachment of Enteromorpha zoospores to a bacterial biofilm assemblage. Biofouling 16: 151–158.

FCUP Chapter 7 183

Joint I., Tait, K., Callow, M.E., Callow, J.A., Milton, D., Williams, P., Cámara, M. 2002. Cell-to- Cell communication across the Prokaryote–Eukaryote boundary. Science 298: 1207.

Kolar, C.S., Lodge, D.M. 2001. Progress in invasion biology: predicting invaders. Trends Ecol Evol. 16:199–204.

Kotrikla, A., 2009. Environmental management aspects for TBT antifouling wastes from the shipyards. Journal of Environmental Management 90: S77–S85.

Lawes,J.C.., Neilan, B.A.., Brown M.V., Clark, G.F, Johnston E.L. 2016. Elevated nutrients change bacterial community composition and connectivity: high throughput sequencing of young marine biofilms. Biofouling 32:1: 57-69.

Leão, P. N., Engene, N., Antunes, A., Gerwick, W. H., and Vasconcelos, V. 2012. The chemical ecology of cyanobacteria. Nat. Prod. Rep. 29: 372–391.

Leão, P.N., Costa, M., Ramos, V., Pereira, A.R., Fernandes, V.C., Domingues, V.F., Gerwick, W.H., Vasconcelos, V.M., Martins, R. 2013.Antitumor activity of hierridin B, a cyanobacterial secondary metabolite found in both filamentous and unicellular marine strains. PLoS ONE 8: e69562.

Lebret, K., Thabard, M., Hellio, C. 2009. Algae as marine fouling organisms: adhesion damage and prevention. In: Hellio C, Yebra DM, editors. Advances in marine antifouling coatings and technologies. Cambridge (UK): Woodhead Publishing Ltd & CRC Press. p. 80–112.

Lejars M., Margaillan A., Bressy C. 2012.Fouling release coatings: a nontoxic alternative to biocidal antifouling coatings. Chem. Rev. 112:4347–4390 28.

Leray, M., Knowlton, N. 2015 DNA barcoding and metabarcoding of standardized samples reveal patterns of marine benthic diversity. Proc. Natl. Acad. Sci. USA 112: 2076–2081.

Lockwood, J.L., Hoopes, M.F., Marchetti, M.P. 2007. Invasion ecology. Oxford: Blackwell; p. 466.

Maki, J., Mitchell, R. 2002. Biofouling in the marine environment. In: Bitton H, editor. Encyclopedia.

Manefield, M.T, Rasmussen, B., Henzter, H, Andersen, J.B., Steinberg, P., Kjelleberg, S., and Givskov, M. 2002. Halogenated Furanones Inhibit Quorum Sensing through Accelerated LuxR Turnover. Microbiology 148 (4): 1119–27.

Maréchal, J.-P., Hellio, C. 2009 Challenges for the development of new non-toxic antifouling solutions. Int. J. Mol. Sci. 10: 4623–4637.

FCUP Chapter 7 184

Mieszkin, S., Callow, M.E., Callow, J. 2013. Interactions between microbial biofilms and marine fouling algae: a mini review. Biofouling. 29: 1097 – 1113.

Mineur, F., Johnson, M.P., Maggs, C.A.2008. Macroalgal Introductions by hull fouling on recreational vessels: seaweeds and sailors. Environ. Manage. 42: 667-676.

Molino, P.J., Campbell, E., Wetherbee, R. 2009. Development of the initial diatom microfouling layer on antifouling and fouling-release surfaces in temperate and tropical Australia. Biofouling. 25:685–694. de Nys, R., Guenther, J. 2009. The impact and control of biofouling in marine finfish aquaculture. In: Hellio C, Yebra D, editors. Advances in marine antifouling coatings and technologies. Cambridge (UK): Woodhead Publishing Ltd. p. 177–221.

Nunnery, J.K.; Engene, N.; Byrum, T.; Cao, Z.; Jabba, S.V.; Pereira, A.R.; Matainaho, T.; Murray, T.F.; Gerwick, W.H. 2012. Biosynthetically intriguing chlorinated lipophilic metabolites from geographically distant tropical marine cyanobacteria. J. Org. Chem. 77: 4198–4208.

Omae, I. 2003. General aspects of tin-free antifouling paints. Chem. Rev. 103: 3431–3448.

Ortega-Morales, B.O., Santiago-García, J., Chan- Bacab, M., Moppert, X., Miranda-Tello, E., Fardeau, M.L., Carrero, J., Bartolo-Pérez, P., Valadéz-González, A., Guezennec, J. 2007. Characterization of extracellular polymers synthesized by tropical intertidal biofilm bacteria. J. Appl. Microbiol. 102: 254–264.

Paerl, H.W., Paul, V. 2011. Climate change: links to global expansion of harmful cyanobacteria. Water. Res. 46:1349–1363.

Parsek, M.R. Greenberg, EP. 2005. Sociomicrobiology: The Connections between Quorum Sensing and Biofilms. Trends in Microbiology 13 (1): 27–33.

Patel, P., Callow, M.E., Joint, I. Callow, J. 2003. Specificity in the Settlement–modifying Response of Bacterial Biofilms towards Zoospores of the Marine Alga Enteromorpha. Environ Microbiol. 5: 338–49.

Penesyan, A., Marshall-Jones, Z., Holmström, C., Kjelleberg, S., Egan, S. 2009. Antimicrobial activity observed among cultured epiphytic bacteria reflects their potential as source of new drugs. FEMS Microbiol Ecol. 69:113–124.

Pollet, T., Berdjeb, L., Garnier, C. et al. 2018. Prokaryotic community successions and interactions in marine biofilms: the key role of Flavobacteriia. FEMS Microbiol. Ecol. 94: fiy083

FCUP Chapter 7 185

Qian, P. Y., S. C K Lau, H. U. Dahms, S. Dobretsov, Harder, T. 2007. Marine biofilms as mediators of colonization by marine macroorganisms: implications for antifouling and aquaculture. Marine Biotechnology 9: 399–410.

Qian, P.Y., Xu, Y., Fusetani, N., 2010. Natural products as antifouling compounds: recent progress and future perspectives. Biofouling 26: 223-234.

Qian, P.Y., Chen, L., Xu, Y. 2013 Mini-Review: Molecular mechanisms of antifouling compounds. Biofouling 29: 381–400.

Qian, P-Y.; Li, Z; Xu, Y.; Li, Y.; Fusetani, N. 2015. Mini-review: Marine natural products and their synthetic analogs as antifouling compounds: 2009–2014. Biofouling 31: 101–122.

Railkin, A.I. 2004. Marine biofouling – colonization, processes and defenses. Boca Raton, USA: CRC Press.

Rao, D., Webb, J. S., Holström, C., Case, R., Low, A., Steinberg, P., and Kjelleberg, S. 2007. Low densities of epiphytic bacteria from the marine alga Ulva australis inhibit settlement of fouling organisms. Appl. Environ. Microbiol. 73: 7844–7852.

Rasmussen, T.B. and Givskov, M. 2006. Quorum-sensing inhibitors as anti-pathogenic drugs. Int. J. Med. Microbiol. 296: 149–161.

Salta, M., Wharton, J., Stoodley, P., Dennington, S., Goodes, L., Werwinski, S., et al. 2010. Designing biomimetic antifouling surfaces. Philos Trans R Soc Lond A. 368: 4729–4754.

Salta, M., Wharton, J.A., Blache, Y., Stokes, K.R., Briand, J.F. 2013. Marine biofilms on artificial surfaces: structure and dynamics. Environ. Microbiol. 15:2879.

Scardino, A.J., and de Nys, R. 2011. Mini review: biomimetic models and bioinspired surfaces for fouling control. Biofouling 27: 73-86.

Schultz, M.P., Swain, G.W. 2000. The influence of biofilms on skin friction drag. Biofouling 15: 129–139.

Schultz, M.P. 2007. Effects of coating roughness and biofouling on ship resistance and powering. Biofouling 23: 331–41.

Schultz, M.P., Bendick, J.A., Holm, E.R., Hertel, W.M. 2011. Economic impact of biofouling on a naval surface ship. Biofouling 27:87–98.

Sivonen, K., Jones, G,1999. Cyanobacterial toxins. In: Chorus I, Bartram J (eds) Toxic cyanobacteria in water: a guide to their public health consequences, monitoring and management. E & FN Spon, London, p. 41.

FCUP Chapter 7 186

Sonak, S., Pangam, P., Giriyan, A., Hawaldar, K. 2009. Implications of the ban on organotins for protection of global coastal and marine ecology. J Environ Manage. 90(Suppl 1): S96–S108.

Sunda, W.G., Granéli, E., Gobler, C. J. 2006. Positive feedback and their development of ecosystem disruptive algal bloom. J. Phycol. 42: 963-974.

Tait, K., Joint, I., Daykin, M., Milton, D. L., Williams, P., Cámara, M. 2005. Disruption of quorum sensing in seawater abolishes attraction of zoospores of the green alga Ulva to bacterial biofilms. Environ. Microbiol. 7: 229–240.

Tait, K., Williamson, H., Atkinson, S., Williams, P., Cámara, M., Joint, I., 2009. Turnover of quorum sensing signal molecules modulates cross-kingdom signaling. Environ. Microbiol. 11: 1792–1802.

Thomas, K.V., Brooks, S. 2010. The environmental fate and effects of antifouling paint biocides. Biofouling 26:73–88.

Townsin, R.L. 2003. The ship hull fouling penalty. Biofouling 19: 9–15.

Voulvoulis, N., Scrimshaw, M.D., Lester, J.N. 1999. Alternative antifouling biocides. Appl. Organometal Chem. 13:135–143.

Wahl, M. 1989. Fouling and Antifouling - Some Basic Aspects. Mar Ecol Prog Ser. 58: 175–89.

Wang, K.L., Wu, Z.H., Wang, Y., Wang, C.Y., Xu, Y. 2017.Mini-Review: Antifouling natural product from marine microorganisms and their synthetic analogs. Mar Drugs. 15: 266.

Whitton, B. A., Potts, M. 2000. in The Ecology of Cyanobacteria, eds. Whitton, B. A. & Potts, M. (Kluwer, Dordrecht, The Netherlands), pp. 1–11

Xu, J., Tang, W., Ma, J., Wang, H. 2017. Comparison of microbial community shifts in two parallel multi-step drinking water treatment processes. Appl. Microbiol. Biotechnol. 101: 5531–5541.

Yamada, H. 2007.Behaviour, occurrence, and aquatic toxicity of new antifouling biocides and preliminary assessment of risk to aquatic ecosystems. Bull Fish Res Agen. 21:31–45.

Yebra, D., Meseguer, S.K., Dam-Johansen, K. 2004. Antifouling Technology - Past, Present and Future Steps towards Efficient and Environmentally Friendly Antifouling Coatings. Prog. Org. Coat. 50: 75–104.

Zargiel, K.A., Coogan, J.S., Swain, G.W.2011. Diatom community structure on commercially available ship hull coatings. Biofouling. 27:955–965.

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Zhang, S. P., Huang, R., Li, F. F., Wei, H. X., Fang, X. W., Xie, X. S., 2016. Antiviral anthraquinones and azaphilones produced by an endophytic fungus Nigrospora sp. from Aconitum carmichaeli. Fitoterapia 112: 85–89.

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8- Publications, conference proceedings and attended courses

Publications (acknowledging PhD scholarship SFRH/BD/99003/2013):

Antunes JT, Azevedo, J., Rego, A., Sousa, A., Leão PN, Vasconcelos VM. 2019. Distinct profiles of early marine biofilms in two seasons in a Portuguese Atlantic Port. Manuscript to be submitted.

Antunes JT, Leão PN, Vasconcelos VM. 2018. Marine biofilms: diversity of communities and of chemical cues. Environmental Microbiology Reports. url: https://doi.org/10.1111/1758- 2229.12694

Almeida, J.R., Moreira, J., Pereira, D., Pereira, S. Antunes, J., Palmeira, A., Vasconcelos, V., Pinto, M., Correia-da-Silva, M., Cidade, H. 2018 Potential of synthetic chalcone derivatives to prevent marine biofouling. Science of the Total Environment 643: 98–106. url: https://doi.org/10.1016/j.scitotenv.2018.06.169

Almeida, J.R., Correia-Da-Silva, M., Sousa, E., Antunes, J., Pinto, M., Vasconcelos, V., Cunha, I., 2017. Antifouling potential of nature-inspired sulfated compounds. Scientific Reports 7, 42424. doi: 10.1038/srep42424 url: https://www.nature.com/articles/srep42424

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Conference proceedings (acknowledging PhD scholarship SFRH/BD/99003/2013):

Jorge Antunes, Joana Azevedo, Vitor Vasconcelos and Pedro Leão. Search for cyanobacterial bioactive compounds towards natural antifouling development. Abstract accepted to poster communication at Bio IberoAmérica, held on Salamanca, Spain. 5-8 June 2016.

Joana Azevedo, Jorge Antunes, Pedro Leão and Vitor Vasconcelos. Anti-fouling potential from the LEGE culture collection (LEGE-CC). Poster communication at the 30th International Symposium on the Chemistry of Natural Products and the 10th International Congress on Biodiversity (ISCNP30 & ICOB10), held on Athens, Greece, 25-29 November 2018.

Jorge Antunes, Sandra Pereira, Tiago Ribeiro, Jeffrey E. Plowman, Ancy Thomas, Stefan Clerens Alexandre Campos, Vitor Vasconcelos and Joana R. Almeida. Evaluation of the anti-fouling potential of the cyanobacterial metabolites portoamides using a multi-bioassay approach. Oral presentation at the BIOPROSP_19, held on Tromso, Norway, 25-27 February 2019.

Attended courses:

Marine Metagenomics Workshop, May 7-11, 2018. IGC, Oeiras, Portugal.

Bioactivities Course: Cellular effects of marine natural compounds. November 13-16, 2017. Matosinhos, Portugal.

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9- Appendix

9.1 From Chapter 3 “Distinct profiles of early marine biofilms in two different seasons in a Portuguese Atlantic Port” (manuscript to be submitted)

Supplementary figure 1: Rarefaction plot of observed OTUs for all the 66 samples.

9.2 From Chapter 4 “A multi-bioassay integrated approach to assess the antifouling potential of the cyanobacterial metabolites portoamides”

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Protein name Accession Significanc Cover #Pept #Unique PTM Avg. e age ides peptides Mass (%)

Energy metabolism

ATP synthase subunit d, XP_009028970.1 14.86 7 1 1 N 20040 mitochondrial ATP synthase subunit beta XP_021356377.1 57.57 30 13 13 Y 56361 mitochondrial H+ ATPase a subunit ABJ51956.1 151.76 22 12 12 N 59490 mitochondrial ATP synthase subunit gamma XP_009064140.1 68.99 11 3 3 N 3230659490

Isocitrate dehydrogenase AFI56365.1 32.31 12 5 5 N 50501

Malate dehydrogenase AAF27650.1 25.87 7 2 2 N 35955

Glutamate dehydrogenase XP_022314664.1 8.96 2 1 1 N 60325 mitochondrial

Structural

Myosin heavy chain striated muscle XP_022317649.1 200 25 47 41 Y 228093

Pedal retractor CAB64663.1 153.53 33 21 19 N 87460 muscle

Tubulin beta chain XP_014664190.1 200 28 9 5 Y 49949

Tubulin alpha-1A chain XP_021370666.1 50.58 39 13 2 Y 38578

Cilia- and flagella- XP_022332848.1 associated protein 23.01 8 2 2 N 33218

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Myosin heavy chain KFB49247.1 139.38 39 6 5 Y 20888

Tubulin beta chain NP_001292292.1 37.69 23 6 2 Y 33218

Tubulin beta chain XP_009029528.1 9.32 30 5 1 N 27930

Myosin heavy chain XP_021350592.1 65.88 5 8 8 Y 230478

Tektin-4 XP_011416098.1 24.6 6 2 2 N 52138

Radial spoke head protein 4 XP_021370563.1 8.73 4 1 1 N 50579

Radial spoke head protein 9 XP_021363337.1 8.56 4 1 1 N 36000

Tektin-3 XP_011450983.1 49.81 13 7 7 Y 62040

Collagen, type VI, alpha 3 XP_021375365.1 18.34 7 1 1 N 15965

Protein activity regulation

Dolichyl- diphosphooligosaccharide protein XP_021341263.1 27.49 8 4 4 N 68234 glycosyltransferase

Arginine kinase AKS48144.1 132.06 20 6 6 Y 42147

Dolichyl- diphosphooligosaccharide protein XP_021367901.1 22.72 4 2 2 N 48797 glycosyltransferase 48 kDa subunit

14-3-3 protein XP_018019106.1 200 26 6 4 Y 29977

Peptidyl-prolyl cis- trans isomerase XP_011444269.1 26.14 9 2 2 N 29564

heat shock protein CAJ85741.1 82.08 12 8 8 N 83125 90

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Protein disulfide- isomerase A6 XP_011446200.1 8.85 2 1 1 N 47641

Gene transcription/translation

Histone H2A Q6WV66.3 8.82 37 4 1 N 13360

Histone H2A XP_021377317.1 8.79 34 4 1 Y 14212

Leucine-rich repeat flightless-interacting XP_021363857.1 9.33 3 1 1 N 45577 protein 2

Histone H4 XP_014590042.1 200 50 6 6 Y 11367

Heterogeneous nuclear ribonucleoprotein XP_012945445.1 18.28 3 1 1 N 36578 87F 60S ribosomal protein L7a XP_021369622.1 8.9 5 1 1 N 30513

40S ribosomal protein S4 XP_022290789.1 24.47 7 2 2 N 29583

40S ribosomal protein S3 XP_011438308.1 33.34 9 2 2 N 26632

40S ribosomal protein S13 XP_011441368.1 8.94 8 1 1 N 17356

40S ribosomal protein S25 XP_021370373.1 27.7 18 2 2 N 16837

60S ribosomal protein L11 XP_013068480.1 30.49 13 2 2 N 20256

Transport

Clathrin heavy chain 1 XP_021354511.1 11.55 1 1 1 N 192108

Annexin B9 XP_022317867.1 8.76 5 1 1 N 35854

ADP, ATP carrier SCN46548.1 77.98 15 4 4 Y 33267 protein

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Voltage-dependent ADI56517.1 anion channel 75.71 12 3 3 N 30686

Other processes

6-phosphogluconate dehydrogenase XP_022343934.1 8.76 2 1 1 N 53103 decarboxylating

Glutathione S- transferase sigma 2 AFQ35984.1 22.18 11 2 2 N 23359

9.3 From Chapter 5 “Search for cyanobacterial compounds with antimicrofouling and quorum sensing inhibition activity”

Supplementary table 2. Screening results for the crude cyanobacterial extracts against bacteria and diatom growth and against quorum-sensing inhibition

Strain Concentra C. V. H. R. P. Navicula Cylindro Halom Nitzschi QS tion marina harvey aquam litoralis atlantica theca phora a i arina

LEGE 3 µg/mL - - - - 27% - - - - - 08332 30 µg/mL LEGE 3 µg/mL ------+ 00060 30 µg/mL 22% 26 % 62% 72% LEGE 3 µg/mL 28% 13% 65% ------+ 03273 30 µg/mL 39% 24% 25% 65% LEGE 3 µg/mL 15% 18% 9% 25% 15% - - - - + 07075 21% 22% 11% 31% 12% 30 µg/mL

LEGE 3 µg/mL - - 23% ------+ 07221 30 µg/mL 24% 48% LEGE 3 µg/mL - 27% 39% ------09399 30 µg/mL 37% 32% 80% XX 0061 3 µg/mL - - 31% - 16% - - - - + 30 µg/mL - - 27% 24% 15% 42%

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82% - Inhibitio LEGE 3 µg/mL ------n of 00049 growth 30 µg/mL 86% LEGE 3 µg/mL - - 29% - 20% 33% 63% - - + 03278 30 µg/mL 40% 60% 83% LEGE 3 µg/mL - - 23% - - - - - Inhibitio 06223 34% n of growth 30 µg/mL - X-357 3 µg/mL - - 23% - 23% - - - + 30 µg/mL 20% - - 72% X-002 3 µg/mL - - 4% - - - - - 31% + 30 µg/mL 35% - - 82% 68% LEGE 3 µg/mL - - 13% - 9% - - - - - 00247 30 µg/mL 61% 15% LEGE 3 µg/mL ------00055 30 µg/mL LEGE 3 µg/mL ------37% + 00053 30 µg/mL 87% 47% LEGE 3 µg/mL ------+ 06134 30 µg/mL LEGE 3 µg/mL 27% ------+ 00065 30 µg/mL 21% - LEGE 3 µg/mL - - 33% - - - - - 37% + 00052 30 µg/mL 29% 57% LEGE 3 µg/mL - 16% ------Inhibition 07197 27% 26% of growth 30 µg/mL LEGE 3 µg/mL ------03275 30 µg/mL LEGE 3 µg/mL ------07148 30 µg/mL - LEGE 3 µg/mL ------+ 05292 30 µg/mL - LEGE 3 µg/mL 25% ------51% - - 06141 30 µg/mL - - - 22% 86% LEGE 3 µg/mL 19% - - - 13% - - - 45% + 06014 30 µg/mL 15% 27% - - 26% 52% 49% 63% 30 µg/mL LEGE 3 µg/mL 20% - - - - - 75% - Inhibition 91342 22% 30% 15% 23% of growth 30 µg/mL 85% - 37% LEGE 3 µg/mL ------75% - - 06020 30 µg/mL 85% 3 µg/mL ------

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LEGE 30 µg/mL 06225 LEGE 3 µg/mL ------04288 30 µg/mL 78% LEGE 3 µg/mL ------+ 06224 30 µg/mL LEGE 3 µg/mL 19% 14% ------+ 00041 30 µg/mL - 21% - LEGE 3 µg/mL ------+ 11428 30 µg/mL 29% 29% - 25% 47% Col. 1 3 µg/mL 15% - 40% ------+ HB 30 µg/mL 25% 71% LEGE 3 µg/mL ------+ 07229 30 µg/mL 65% LEGE 3 µg/mL 17% ------+ 06119 30 µg/mL 30% 34% LEGE 3 µg/mL - 33% - - - - - Inhibition 06361 - 50% of growth 30 µg/mL - 55% 54% LEGE 3 µg/mL - - 39% ------06116 30 µg/mL 46% 48% LEGE 3 µg/mL 19% - 40% - - - - 45% - + 00038 30 µg/mL 26% 72% 61% LEGE 3 µg/mL - - 54% ------06148 30 µg/mL 55% - LEGE 3 µg/mL 31% ------06113 30 µg/mL 56% LEGE 3 µg/mL 25% - 30% - - - - 91093 30 µg/mL 34% 55% - - 65% - LEGE 3 µg/mL - - 25% - - - 60% - - - 06013 53% 30 µg/mL 87% LEGE 3 µg/mL ------36% - + 00259 30 µg/mL 90% LEGE 3 µg/mL - - - - 27% - 56% - - 06114 30 µg/mL 22% - 78% LEGE 3 µg/mL ------06137 30 µg/mL - - - 80% 87%

LEGE 3 µg/mL ------76% - - 00065 30 µg/mL - 23% LEGE 3 µg/mL ------42% - - 12432 30 µg/mL - 48% 47% 67% LEGE 3 µg/mL - - 42% ------06130 30 µg/mL 37% - 3 µg/mL - - - - 27% - - - - -

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LEGE 30 µg/mL - 12168 LEGE 3 µg/mL ------66% - - 91351 30 µg/mL 60% 61% 46% LEGE 3 µg/mL ------06143 30 µg/mL LEGE 3 µg/mL ------06118 30 µg/mL 71% LEGE 3 µg/mL 14% - - 15% ------XX280 30 µg/mL 23% 10% 38% 12% LEGE 3 µg/mL ------00141 30 µg/mL LEGE 3 µg/mL - - - - 25% - - - - - 06105 30 µg/mL 20% 27% - LEGE 3 µg/mL ------06135 30 µg/mL LEGE 3 µg/mL - - - - 28% - 32% - - - 06109 30 µg/mL 12% 67% LEGE 3 µg/mL ------06140 30 µg/mL LEGE 3 µg/mL ------07173 30 µg/mL 54% LEGE 3 µg/mL - - 24 % - - - - 37% 00246 20% - + 30 µg/mL 49%

Note: Values represent the average of inhibition of growth of the bacteria or diatom species in the presence of the crude of the cyanobacteria when compared to growth in MB medium (negative control). “+” Represents inhibition of quorum sensing activity in the presence of cyanobacterial extract. “-“. Represents the absence of inhibition of quorum-sensing activity in the presence of cyanobacterial extract.

Supplementary table 3: Activity for fractions and subfractions of LEGE 00060 (E16156)

C. V. H. aquam P. R. Cylindro- Halom- Navicula Nitzschia QS Fraction marina harveyi arina atlantica litoralis theca phora

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E17159 12% 18% 22% 26% 44% 27% 45% - 23% - A E17159 - - 19% - 50% - - - - - B E17159 - - 12% - 38% - - - - - C E17159 ------D E17159 ------E E17159 ------F E17159 ------G E17159 ------H E17159 - - - - 13% - - - - - I

Supplementary table 4: Activity for fractions of LEGE 00247 (E 17159)

C. V. H. aquam P. R. Cylindro- Halom- Navicula Nitzschia QS Fraction marina harveyi arina atlantica litoralis theca phora

E16156 18% - - - 18% - - - - - A E16156 - - 19% - 50% - - - - - A1 E16156 - 21% ------A2 E16156 - 15% 23% 23% ------A3 E16156 18% - 23% 24% - - 26% 32% - - A4 E16156 ------

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B E16156 ------B1 E16156 ------B2 E16156 ------B3 E16156 ------B4 E16156 ------E E16156 - - 30% 30% ------E1 E16156 ------E2 E16156 12% 48% 12% 12% - - - - 35% 21% E3 E16156 14% - 11% 12% ------E4

Supplementary table 5: Activity for fractions of LEGE 00246 (E17162)

C. V. H. P. R. Cylindro- Halom- Fraction marina harveyi aquama atlantica litoralis theca phora Navicula Nitzschia QS rina E17162 - 20% 30% 22% ------A E17162 - - - 20% - - - - - B

E17162 ------C E17162 ------D E17162 ------E E17162 ------F E17162 ------G

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E17162 ------H E17162 14% ------I

Supplementary table 6: Activity for fractions of LEGE 11428 (Subfractioning of E17163)

C. V. H. P. R. Cylindro- Halom- Fraction marina harveyi aquamarina atlantica litoralis theca phora Navicula Nitzschia QS n E17163 37% - 12% 12% 37% - - - - - A E17163 - - - - 19% - - - - - B E17163 ------C E17163 - 21% ------D E17163 14% - 23% 24% 24% 25% - 32% - - E E17163 - - 24% 23% ------F E17163 ------G E17163 ------H E17163 - 25% - 26% - 12% - 25% - + I

Supplementary table 7: Activity for fractions of LEGE 03273 (E17166)

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C. V. H. aqua P. R. Cylindro- Halom- Navicula Nitzschia Fraction marina harveyi marina atlantica litoralis theca phora QS

E17166 ------A E17166 13% 13% - - 13% - - - - - B

E17166 - - - - 15% - - - - - C E17166 16% 15% 14% 20% 16% - - - - + D E17166 ------E E17166 - 20% 20% 20% 20% 20% - - - + F E17166 - 10% 11% 10% 10% 10% - - - - G E17166 ------H E17166 I - - 9% ------

Supplementary table 8: Activity for fractions of LEGE 11428 (Subfractioning of E17163)

C. V. H. aquam P. R. Cylindro- Halom- Navicula Nitzschia Fraction marina harveyi arina atlantica litoralis theca phora QS

E17163 7% - - 17% ------E3A E17163 ------E3B E17163 ------E3C E17163 ------E3D E17163 12% ------E3E

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Supplementary table 9: Activity for fractions of LEGE 11428 (Subfractioning of E17163)

C. V. H. P. R. Cylindro- Halom- Navicula Nitzschia Fraction marina harveyi aquamarina atlantica litoralis theca phora QS

E17163 ------I1 E17163 ------I2 E17163 ------I3 E17163 ------I4 E17163 12% ------I5 E17163 12% 19% 12% 30% 24% 15% 17% - 21% + I6 E17163 - - - 20% 27% 32% - - - - I7 E17163 - 9% - 19% ------I8 E17163 11% 11% 11% 9% - 21% 15% - 17% + I9

Note: Values represent the average of inhibition of growth of the bacteria or diatom species in the the presence of the crude of the cyanobacteria when compared to growth in MB medium (negative control). “+” Represents inhibition of quorum sensing activity in the presence of cyanobacterial extract. “-“. Represents the absence of inhibition of quorum-sensing activity in the presence of cyanobacterial extract.

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Supplementary figure 3: 400MHZ 1H NMR spectrum of fraction E 16156_E3.

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Supplementary figure 4: 400MHZ 1H NMR spectrum of fraction E 16156_E4.

Supplementary figure 5. 400MHZ 1H NMR spectrum of fraction E 17159_A.

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Supplementary figure 6. 400MHZ 1H NMR spectrum of fraction E 17162_A.

Supplementary figure 7. 400MHZ 1H NMR spectrum of fraction E 17166_D.

Supplementary figure 8: 400MHZ 1H NMR spectrum of fraction E 17163_I.

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