characterization of cyanobacteria isolated from the

D

Characterization of

cyanobacteria isolated

from the Portuguese

coast: diversity and

potential to produce

bioactive compounds

Ângela Marisa Oliveira Brito Programa Doutoral em Biologia de Plantas Departamento de Biologia

2015

Orientador Paula Tamagnini, Professora Associada

Faculdade de Ciências, Universidade do Porto

Coorientador Vitor M. Vasconcelos, Professor Catedrático

Faculdade de Ciências, Universidade do Porto

Coorientador Marta Vaz Mendes, Investigadora Auxiliar

IBMC - Instituto de Biologia Molecular e Celular

Aos meus pais

One should not pursue goals that are easily achieved. One must develop an instinct for

what one can just barely achieve through one’s greatest efforts.

Albert Einstein

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast vii

Acknowledgments

The work presented here is the result of a close collaboration of several people

(involved directly or indirectly) to whom I would like to express my gratitude.

First, I want to acknowledge my supervisor, Prof. Paula Tamagnini, for this

opportunity and for the extraordinary support, guidance and patience (especially during

the thesis writing) always expressed over all these years (not only during the PhD).

Thank you for sharing your knowledge, for always keeping your door open and for the

encouragement throughout this work (not always an easy path).

I would like to acknowledge my co-supervisor Marta Vaz Mendes, for all the help,

support, and valuable teachings. Thank you for spending time to teaching and helping

me with the “PKS and NRPS world”.

I would also like to thank my co-supervisor Prof. Vitor Vasconcelos for the

opportunity. Thank you for the valuable advices, support and availability always

expressed.

I want to express my gratitude to Professor William H. Gerwick at the Scripps

Institution of Oceanography, University of California San Diego, for giving me the

opportunity to spend some time in his lab. It was a pleasure to work and learn so much

on cyanobacteria secondary metabolites. Thank you so much for welcoming me in your

lab and for showing me the wonderful world of marine natural products.

A huge thank you to Vitor Ramos for the precious help and advices that were crucial

for the achievement of the results presented in this thesis. Thank you for your support

and friendship.

I am particularly grateful to all co-authors that contributed to this work. Without you

this work would not be possible.

Prof. Arlete Santos, as well as Rui Fernandes and Francisco Figueiredo (Histology

and Electron Microscopy facility, IBMC), for all the help with the transmission electron

microscopy.

I would like to thank Cristina Vieira and Jorge Vieira for assistance with the

phylogenetic analysis.

To all present and former members of the Bioengineering and Synthetic

Microbiology (BSM) Group, a big thank you for the wonderful work environment, the

viii FCUP

Characterization of cyanobacteria isolated from the Portuguese coast

sharing of ideas, help, support, friendship, tips and laughs. Sharing my lab time with

you all was a fun and constructive experience.

Special thanks to Rita Mota. Thanks for all the help (not only in the work), patience,

and for constantly showing me the things in a different way. Thanks for your friendship,

and for being with me in all moments (good and bad ones). Everything would be harder

without you!

To Sara Pereira. Thank you for your help, support and friendship. Thanks for being

always there. I will never forget that my “first steps” in the lab were with you! Thanks for

everything!

To Meritxell Notari. Thanks for your good mood and help either in the lab and at

home. I think that you deserve the award for “best housemate”, because I know that it

was not easy to share almost 24h per day with me. Thanks for your patience,

friendship and all good moments shared.

To Filipe Pinto. Thank you for your support and for being always available to help

me (and anyone who asks). Thank you for helping me with images, computer problems

and I could continue... (quem tem um Filipe tem tudo!).

To Catarina Pacheco (thanks for the help and support over all these years), Paulo

Oliveira (thanks for constructive discussions about the work and for your help with the

GC), Steeve Lima, Eunice Ferreira, Carlos Flores, Marina Santos and José Pereira for

the good work environment, the sharing of ideas and help. Thank you all.

To the present and former members of the MCA and RCS groups. Thank you for

your support and friendship.

To all members of the Gerwick Laboratory, especially Evgenia Glukhov, Ozlem

Demirkiran, Karin Kleigrewe and Enora Briand. Thank you for your precious help.

To all members of LEGE, CIIMAR. Thank you for all the help, support and advices.

To all my friends, especially Juliana Oliveira (thank you for being by my side and for

your support that encouraged me to never look back) and all members of the “team”:

Rita Mota, Meritxell Notari, Sara Pereira, Filipe Pinto, Ricardo Celestino (thanks for

your help with the thesis, thank you for everything…except the spiders!), Joana

Gaifem, Sérgio Ferreira, Nádia Gonçalves and Nelson Ferreira. Thank you for all the

good moments spent together, dinners, parties, laughs. Thank you for being always

there (in the good and bad times).

Special thanks to Goody and Brian. Thank you for welcoming me in your house

during my stay in San Diego, for your care and for all the good moments. You made

everything easier; all the words won’t be enough to thank you.

I would also like to acknowledge the Fundação para a Ciência e Tecnologia (FCT)

for the PhD. Grant SFRH/BD/70284/2010.

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast ix

And for last, but for sure not least…

Quero agradecer à minha família, em especial aos meus pais (os meus pilares),

pois sem eles nada disto teria sido possível. Não há palavras que descrevam tudo o

que são para mim. Muito obrigada por estarem sempre comigo e apoiarem sempre as

minhas decisões (mesmo por vezes não estarem totalmente de acordo). Obrigada pelo

amor, compreensão, dedicação e pelos valores transmitidos.

À minha irmã Sónia (desde sempre a minha melhor amiga) e ao meu cunhado

Filipe. Obrigada pelo vosso apoio incondicional, por toda ajuda e por estarem sempre

comigo. Vocês foram e são fundamentais para aquilo que sou hoje.

Finalmente, como não poderia deixar de ser, aos meus sobrinhos Tiago Filipe e

Ana Carolina (que nasceu mesmo na fase final de escrita da tese) que são sem dúvida

o melhor do Mundo, são o meu orgulho.

x FCUP


This work was funded by FEDER funds through the Operational Competitiveness

Programme - COMPETE and by national funds through FCT - Fundação para a Ciência e a

Tecnologia under the projects FCOMP-01-0124-FEDER-010600 (PTDC/MAR/102258/2008),

FCOMP-01-0124-FEDER-010568 (PTDC/MAR/099642/2008), FCOMP-01-0124-FEDER-

028314 (PTDC/BIA-MIC/2889/2012) and the PhD grant SFRH/BD/70284/2010 (POPH-QREN).

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xi

List of publications

(Ao abrigo do disposto no número 2 do Artigo 8º do decreto-Lei nº 388/70)

This dissertation is based on the following publications and some unpublished results:

I - Brito Â., Ramos V., Seabra R., Santos A., Santos C.L., Lopo M., Ferreira S.,

Martins A., Mota R., Frazão B., Martins R., Vasconcelos V., Tamagnini P. 2012.

Culture-dependent characterization of cyanobacterial diversity in the intertidal zones of

the Portuguese coast: a polyphasic study. Systematic and Applied Microbiology 35:

110-119.

II - Brito Â., Mota R., Ramos V., Lima S., Santos A., Vieira J., Vieira C.P., Vasconcelos

V.M., Tamagnini P. 2015. Isolation of cyanobacterial strains from the Portuguese

Atlantic coast disclosures novel diversity. (Manuscript).

III - Brito Â., Gaifem J., Ramos V., Glukhov E., Dorrestein P.C., Gerwick W.H.,

Vasconcelos V.M., Mendes M.V., Tamagnini P. 2015. Bioprospecting Atlantic

cyanobacteria for bioactive secondary metabolites reveals untapped chemodiversity.

Algal Research 9: 218-226.

Other publications:

Pinho L.X., Azevedo J., Brito Â., Santos A., Tamagnini P., Vilar V.J.P., Vasconcelos V.

M., Boaventura R.A.R. 2015. Effect of TiO2 photocatalysis on the destruction of

Microcystis aeruginosa cells and degradation of cyanotoxins microcystin-LR and

cylindrospermopsin. Chemical Engineering Journal 268: 144-152.

Publications (Chapter II and IV) are reproduced with the permission from the copyright holders.

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xiii

Abstract

Cyanobacteria are photosynthetic prokaryotes with a wide geographical distribution

and are present in a broad spectrum of environments, including extreme ones. They

play an important role in the global carbon cycle as primary producers and some

strains are able to fix atmospheric nitrogen, characteristic that confers them an

advantage in nutrient-poor and nitrogen-depleted environments like the marine ones.

Cyanobacteria are also recognized as a prolific source of secondary metabolites

with biological activities. Some of these compounds are toxic to a wide array of

organisms, including humans, but others exhibit promising activities with interest in

biotechnological applications, namely in the pharmacological, cosmetic and agricultural

fields. Despite the ecological importance of cyanobacteria and their biotechnological

potential, little is known about the diversity of these organisms on the Portuguese

continental coast.

In this work, the diversity of cyanobacteria on the intertidal zones of the Portuguese

coast was assessed. For this purpose nine beaches were sampled and 60 of the

retrieved isolates were characterized using a polyphasic approach. Based on

morphology, 35 morphotypes were identified being the most representative group the

non-heterocystous forms, with a predominance of the filamentous

Oscillatoriales/Subsection III. The 16S rRNA gene was partially sequenced for all the

isolates and phylogenetic analyses were performed. For the majority of the isolates

there was a broad congruence between the phenotypic and genotypic based

identifications. Moreover, the 16S rRNA gene sequences of nine strains had less 97%

similarity compared the ones available in the databases revealing novel cyanobacterial

diversity. To further characterize these strains, the remaining part of 16S rRNA gene,

the internal transcribed spacer (ITS) and the 23S rRNA gene were sequenced and

additional phylogenetic analyses were carried out. Together with the morphological and

ultrastructural data, this analysis showed that the unicellular isolate LEGE 07459 is

related with the genera Geminocystis and Cyanobacterium (Chroococcales/Subsection

I), but presents distinct characteristics indicating that could be a representative of a

new genus. The isolate LEGE 07179, previously identified as Hyella sp., clusters with

other Pleurocapsales/Subsection II forming a distinct branch. However, since this is the

first sequence available for members of this genus it was not possible to further infer

phylogenetic relationships. Concerning the filamentous strains, the new data indicated

xiv FCUP


that the isolate LEGE 06188 can, indeed, be assigned to the genus Leptolyngbya,

while the strains LEGE 07167, 07176, 07157 and 06111 (previously assigned to L.

fragilis, L. mycoidea, and Phormidium/Leptolyngbya, respectively) constitute a distinct

cluster that probably should be assigned to a new genus, firstly isolated from the

Portuguese coast.

The screening for putative diazotrophs indicated that approximately 30% of our

isolates are potential N2-fixers, suggesting that these cyanobacteria may play an

important role in N2 fixation in the intertidal zones. In the PCR screening for toxin

producers (presence of the genes encoding proteins involved in the production of

microcystin, nodularin and saxitoxin), no amplicons were obtained. However, the

primers used were designed based on sequences from freshwater cyanobacteria

therefore may not be the most appropriate for marine strains. Moreover, previous

studies demonstrated that extracts of cyanobacterial strains also isolated from the

Portuguese coast were toxic to marine invertebrates. To further evaluate the potential

of these isolates to produce other secondary metabolites, a PCR screening for the

presence of genes encoding polyketide synthases (PKS) and non-ribosomal peptide

synthetases (NRPS) targeting the ketosynthase (KS) and adenylation (A) domains

respectively, was performed. The PKS and NRPS genes were present in the majority

of the isolates, with the PKS genes being more ubiquitous than the NRPS genes. The

sequences obtained were used in an in silico prediction for PKS and NRPS systems.

The sequences encoding for KS and A-domains led to the identification of biosynthetic

gene clusters that code for 13 or 10 different compounds, respectively. In general, the

identities were low or had no correspondence indicating that the corresponding

biosynthetic gene clusters are probably not yet characterized. In addition, RT-PCR

analyses revealed that both PKS and NRPS genes were transcribed under routine

laboratory conditions in the majority of the isolates tested.

To complement the previous results on the potential of our isolates to produce

secondary metabolites, crude extracts of 57 of our isolates were analysed by LC-

MS/MS. Their MS/MS spectra together with the MS/MS spectra of reference

cyanobacterial compounds were used to generate a network. From this analysis it was

possible to infer that our samples contained compounds related with naopopeptin and

antanapeptin, an analogue of dolastatin 16 as well as malyngamide - type molecules. A

connection between microcystin, nodularin and nodes from different isolates was also

observed, indicating that some of our strains might be able to produce compounds

related with hepatotoxins. Moreover, there were clusters composed by nodes from our

samples only, highlighting that our strains are able to produce novel compounds.

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xv

Overall, this work provided new insights on the knowledge of cyanobacteria from the

Portuguese Atlantic coast revealing novel biodiversity and highlighting their potential to

fix N2. It was also demonstrated that these isolates are an untapped source of

secondary metabolites, stressing the importance of exploring the potential of

cyanobacterial strains from the temperate regions for the production of novel

compounds.

Keywords: biodiversity; cyanobacteria; nitrogen fixation; NRPS; phylogeny; PKS;

Portuguese coast; secondary metabolites.

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xvii

Resumo

As cianobactérias são procariontes fotossintéticos com uma ampla distribuição

geográfica e que se encontram numa grande diversidade de habitats, incluindo os

ambientes extremos. As cianobactérias desempenham um papel importante no ciclo

global do carbono como produtores primários e algumas estirpes são capazes de fixar

o azoto atmosférico, característica que lhes confere uma vantagem em ambientes

pobres em nutrientes e deficientes em compostos azotados como é o caso dos

ambientes marinhos.

As cianobactérias são também reconhecidas como uma fonte prolífica de

metabolitos secundários com atividades biológicas. Alguns destes compostos são

tóxicos para uma grande variedade de organismos, incluindo o Homem, enquanto

outros têm aplicações biotecnológicas promissoras, em áreas como a farmacologia, a

indústria de cosméticos e a agricultura. Apesar da importância das cianobactérias a

nível ecológico e do seu potencial biotecnológico, sabe-se muito pouco sobre a sua

diversidade na costa continental portuguesa.

Neste trabalho, foi avaliada a diversidade de cianobactérias nas zonas intertidais da

costa portuguesa. Com este propósito, foram amostradas nove praias e de entre os

organismos isolados, 60 foram caracterizados utilizando uma abordagem polifásica.

Com base na morfologia foram identificados 35 morfotipos, sendo as formas não-

heterocísticas o grupo mais representativo, com predomínio das cianobactérias

filamentosas (Oscillatoriales/Subseção III). Para todos os isolados foi sequenciado

parcialmente o gene do 16S rRNA e foi realizada a respetiva análise filogenética. Para

a maioria dos isolados houve uma grande congruência entre a identificação fenotípica

e genotípica. Para além disso, os resultados obtidos mostraram que as sequências do

gene do 16S rRNA de nove estirpes tinham menos de 97% de similaridade com as

sequências disponíveis nas bases de dados, revelando assim nova diversidade. Para

complementar a caracterização molecular destas estirpes foi ainda sequenciado a

restante parte do gene do 16S rRNA, o espaçador interno transcrito (ITS) e o gene do

23S rRNA, e foram realizadas análises filogenéticas adicionais. Em conjunto com o

estudo da morfologia e ultraestrutura, esta análise mostrou que o isolado unicelular

LEGE 07459 está relacionado com os géneros Geminocystis e Cyanobacterium

(Chroococcales/ Subseção I), apresentando contudo características distintas que

indicam tratar-se de um novo género. O isolado LEGE 07179, anteriormente

xviii FCUP


identificado como Hyella sp., agrupa com outras Pleurocapsales/Subseção II, contudo

forma um ramo separado. Como esta é a primeira sequência de um membro do

género Hyella a estar disponível nas bases de dados, não foi possível inferir mais

sobre as relações filogenéticas. No que diz respeito às estirpes filamentosas, os

resultados indicam que o isolado LEGE 06188 pertence ao género Leptolyngbya,

enquanto as estirpes LEGE 07167, 07176, 07157 e 06111 (anteriormente identificados

como L. fragilis, L. mycoidea, e Phormidium/Leptolyngbya, respetivamente) constituem

um grupo distinto, provavelmente um novo género, isolado pela primeira vez do litoral

português.

O rastreio de possíveis estirpes diazotróficas mostrou que aproximadamente 30%

das cianobactérias isoladas são potenciais fixadoras de azoto atmosférico, o que

sugere que estes organismos podem desempenhar um papel importante na fixação de

N2 nas zonas intertidais. Os resultados negativos obtidos no rastreio para possíveis

produtoras de toxinas (PCR para presença de genes que codificam proteínas

envolvidas na produção de microcistina, nodularina e saxitoxina) parecem indicar a

ausência de estirpes tóxicas. No entanto, os oligonucleótidos utilizados foram

desenhados com base em sequências de cianobactérias de água doce logo, podem

não ser os mais apropriados para as estirpes marinhas. Além disso, estudos anteriores

demonstraram que extratos de cianobactérias, também isoladas a partir da costa

portuguesa, eram tóxicos para os invertebrados marinhos. Para continuar a avaliar o

potencial destes isolados para a produção de outros metabolitos secundários, foi

realizado um rastreio por PCR para a presença de genes que codificam as sintases de

policétidos (PKS) e sintetases de péptidos não ribossomais (NRPS), tendo

respetivamente como alvo os domínios de ketosintase (KS) e adenilação (A). Os

resultados revelaram que os genes dos PKS e NRPS estão presentes na maioria dos

isolados, sendo os genes dos PKS mais ubíquos que os genes dos NRPS. As

sequências obtidas foram usadas numa predição in silico dos sistemas de PKS e

NRPS. As sequências que codificam os domínios KS e A levaram à identificação de

agrupamentos de genes cujos produtos podem sintetizar respetivamente 13 ou 10

compostos diferentes. Regra geral, os valores de identidade foram baixos ou não

havia mesmo uma correspondência com as bases de dados indicando que estes

agrupamentos de genes provavelmente ainda não foram caracterizados. Análises de

RT-PCR revelaram também que os genes dos PKS e dos NRPS são transcritos em

condições laboratoriais de rotina na maioria dos isolados testados.

Para complementar os resultados obtidos relativamente à produção de metabolitos

secundários, foram analisados por LC-MS/MS os extratos brutos de 57 dos isolados.

Os espectros obtidos, bem como os espectros de compostos de referência, foram

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xix

utilizados para a construção de uma rede molecular (molecular networking). A partir

desta análise, foi possível inferir que as nossas amostras continham compostos

relacionados com naopopeptina e antanapeptina, um análogo da dolastatina 16, bem

como compostos relacionados com malingamidas. Foi também possível observar uma

ligação entre a microcistina e a nodularina e os pontos de conexão de diferentes

isolados, o que indicam que algumas destas cianobactérias marinhas isoladas da

costa portuguesa podem produzir compostos relacionados com hepatotoxinas. Na

rede molecular foram, também, observados grupos formados exclusivamente por

pontos de conexão relativos aos nossos isolados, o que indica a sua capacidade para

produzir novos compostos.

Em suma, este trabalho possibilitou a aquisição de novos conhecimentos sobre as

cianobactérias da costa atlântica portuguesa, revelando nova biodiversidade e

realçando o potencial de algumas estirpes para fixar azoto atmosférico. Foi também

demonstrado que estes isolados são uma fonte inexplorada de metabolitos

secundários, salientando a importância que a investigação de estirpes de

cianobactérias das regiões temperadas pode ter na descoberta e produção de novos

compostos.

Palavras-chave: biodiversidade; cianobactérias; costa portuguesa; filogenia; fixação

de azoto; metabolitos secundários; NRPS; PKS.

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xxi

Table of Contents

Acknowledgements vii

List of publications xi

Abstract xiii

Resumo xvii

Table of Contents xxi

List of Figures xxv

List of Tables xxix

List of Abbreviations xxxi

Chapter I

General introduction 1

1. Introduction 3

1.1. Cyanobacteria 3

1.2. Taxonomy of cyanobacteria 5

1.3. Nitrogen fixation 8

1.4. Cyanobacterial secondary metabolites 10

1.4.1. Prediction of natural product biosynthetic gene clusters 17

1.4.2. Mass spectrometry (MS) based metabolomics: molecular networking 19

1.5. Cyanobacterial diversity in Portuguese waters and secondary metabolites

production

20

1.6. Objectives 21

Chapter II

Culture-dependent characterization of cyanobacterial diversity in

the intertidal zones of the Portuguese coast: a polyphasic study 23

Abstract 25

Introduction

25

xxii FCUP


Material and Methods

26

Sampling sites 26

Cyanobacteria sampling, isolation, and culturing 26

Light and transmission electron microscopy (TEM) and morphological

identification 26

DNA extraction, purification, PCRs and sequencing 27

Nucleotide sequence aceession numbers 27

Phylogenetic analysis 27

Results and Discussion 27

Morphological characterization 27

DNA sequence and phylogenetic analysis 28

Screening for putative N2 fixers and toxin producers 33

Conclusions 33

Acknowledgments 33

References 33

Supplementary Material I 35

Chapter III

Characterization of novel cyanobacterial strains isolated from the

Atlantic Portuguese coast 43

Abstract 46

Introduction 47

Material and Methods 49

Cyanobacterial strains and culture conditions 49

Light microscopy 49

Transmission electron microscopy (TEM) 49

Scanning electron microscopy (SEM) 50

DNA extraction, PCR amplification, cloning and sequencing 50

Nucleotide sequence accession numbers 51

Phylogenetic analysis 51

Secondary structure of ITS 52

Pigment analysis 52

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xxiii

Nitrogenase activity 53


Isolate belonging to Chroococcales (Subsection I) 54

Isolate belonging to Pleurocapsales (Subsection II) 56

Isolate belonging to Oacillatoriales (Subsection III) 58

Capacity to fix atmospheric nitrogen and to produce secondary metabolites 65

Conclusions 65

Acknowledgments 66

References 67

Supplementary Material II 73

Chapter IV

Bioprospecting Atlantic cyanobacteria for bioactive secondary

metabolites reveals untapped chemodiversity

79

Abstract 81

Introduction 81

Material and Methods 82

Cyanobacterial isolates and culture conditions 82

DNA extraction, PCR amplification and sequencing 82

Sequence analysis and in silico prediction of PKS and NRPS systems 82

Nucleotide sequence accession numbers 82

RNA extraction and RT-PCR 82

Extraction, LC-MS analysis and molecular networking generation 82


PCR screen for PKS and NRPS encoding genes 83

In silico prediction of PKS and NRPS systems 84

Transcriptional analysis 84

LC-MS and molecular networking studies 85

Conclusions 88

Acknowledgments 88

References 88

Supplementary Material III 91

xxiv FCUP


Chapter V

General Discussion, Conclusions and Future Perspectives 99

5.1. General Discussion and Conclusions 101

5.2. Future perspectives 105

General References

References 107

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xxv

List of Figures

Chapter I

Figure 1 Schematic representation of nitrogenase enzymatic complex. 9

Figure 2 Schematic representation of enzymatic domains in polyketide

synthases and nonribosomal peptide synthetases. 11

Figure 3 Examples of chemical structures of cyanobacterial metabolites. 13

Figure 4 Chemical structures of microcystin-LR, nodularin and saxitoxin. 14

Figure 5 Structural organization of the microcystin biosynthetic gene cluster

in Microcystis aeruginosa. 15

Figure 6 Structural organization of the nodularin biosynthetic gene cluster in

Nodularia spumigena. 15

Figure 7 Structural organization of the saxitoxin biosynthetic gene cluster in

Cylindrospermopsis raciborskii. 16

Figure 8 Graphical representation of a traditional network generated with

cyanobacteria extracts and reference compounds. 20

Chapter II

Figure 1 Distribution of the isolates per taxonomic group. 28

Figure 2 Light micrographs illustrating the diversity of cyanobacterial

morphotypes isolated from the intertidal zones of nine beaches on

the Portuguese coast. 30

Figure 3 Transmission electron micrographs of selected isolates from the

sampling sites. 31

Figure 4 Maximum-likelihood phylogenetic tree of partial 16S rRNA gene

sequences from the isolates, reference cyanobacterial strains and

the outgroup Chloroflexus aurantiacus J-10-fl. 32

Figure S1 Light micrographs of the cyanobacterial isolates. 40

Figure S2 Light micrographs of the cyanobacterial isolates (cont.). 41

xxvi FCUP


Figure S3

Neighbor-joining phylogenetic tree of the 16S rRNA gene

sequences from the isolates, reference cyanobacterial strains and

the outgroup Chloroflexus aurantiacus J-10-fl. 42

Chapter III

Figure 1 Micrographs of Geminocysis-like sp. LEGE 07459. 55

Figure 2 Micrographs of Hyella sp. LEGE 07179. 57

Figure 3 Micrographs of Leptolyngbya sp. LEGE 06188. 59

Figure 4 Micrographs of LEGE 07167, previously assigned to Leptolyngbya

fragilis. 60


fragilis. 60


mycoidea. 61

Figure 7 Micrographs of LEGE 06111, previously assigned to

Phormidium/Leptolyngbya. 62

Figure 8 Absorption spectrum obtained for LEGE 06111, previously

assigned to Phormidium/Leptolyngbya. 62

Figure 9 Minimum evolution tree of partial 16S-ITS-23S rRNA gene

sequences from the seven isolates and strains retrieved from the

BLAST analysis. 64

Figure S1 Minimum evolution (ME) phylogenetic tree of partial 16S-rRNA

gene sequences from Geminocysis-like sp. LEGE 07459. 75

Figure S2 Minimum evolution phylogenetic tree of partial 16S rRNA gene

sequence from the Hyella sp. LEGE 07179. 76

Figure S3 Minimum evolution phylogenetic tree of partial 16S- rRNA gene

sequence from the Leptolyngbya sp. LEGE 06188. 77

Figure S4 Minimum evolution phylogenetic tree of partial 16S rRNA gene

sequence from the LEGE 07176 (Leptolyngbya fragilis), LEGE

06111 (Phormidium sp.), LEGE 07167 (Leptolyngbya fragilis) and

LEGE 07157 (Leptolyngbya mycoidea). 78

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xxvii

Chapter IV

Figure 1 Frequency of the in silico predicted PKS and NRPS genes in 61

marine cyanobacterial strains from Portugal using the NaPDoS

software for PKS genes and the PKS/NRPS analysis tool for the

NRPS genes. 85

Figure 2 Transcription of genes encoding the KS and A domains of the PKS

and NRPS genes evaluated by RT-PCR. 85

Figure 3 MS/MS-based molecular network of the extracts from 57 cultured

cyanobacterial strains seeded with 105 reference compounds using

a cosine similarity score cut-off of 0.6. 86

Figure S1 Proposed structure dolastatin16 analogue ion of 760. 97

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xxix

List of Tables

Chapter I

Table 1 General characteristics of the cyanobacterial Orders/Subsections. 7

Table 2 Examples of cyanobacteria secondary metabolites and respective

biological activities. 12

Chapter II

Table 1 Localization and characteristics of the sampling sites. 26

Table 2 Target genes and oligonucleotide primers used in this study. 28

Table 3 Morphological identification and molecular analysis of the

cyanobacterial isolates. 29

Table S1 Morphological identification of the cyanobacteria isolated from

intertidal zones of the Portuguese coast. 37

Chapter III

Table 1 Cyanobacterial strains, culture collection code and sampling sites. 49

Table 2 Oligonucleotide primers used in this study. 51

Table 3 Secondary structure of 16S-23S rRNA internal transcribed (ITS) of

Geminocystis-like sp. LEGE 07459. 56

Chapter IV

Table 1 Oligonucleotide primers used for RT-PCR targeting the

cyanobacterial sequences encoding the KS (KS primers) and A

(AD primers) domains of the PKS and NRPS genes, respectively.

83

xxx FCUP


Table 2

PCR screening for sequences encoding the ketosynthase domain

(KS) of PKS genes and the adenylation-domain (A) of NRPS genes

in cyanobacterial isolates from intertidal zones of the Portuguese

coast. 84

Table S1 Cyanobacterial KS domain sequence based prediction of PKS

using the NaPDoS software. 93

Table S2 Cyanobacterial A domain sequence based prediction of NRPS

using the PKS/NRPS analysis software. 96

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xxxi

List of Abbreviations

A Adenylation

ACP Acyl carrier protein

Adp Anabaenopeptilide

antiSMASH Antibiotics and secondary metabolite analysis shell

ARA Acetylene reduction assay

AT Acyltransferase

ATP Adenosine triphosphate

Bleom Bleomycin

bp base pairs

C Condensation

C2H2 Acetylene

CCAP Culture Collection of Algae and Protozoa

Cch Coelichelin

cDNA complementary DNA

CH2Cl2 Dichloromethane

CH3CN Acetonitrile

Chl a Chlorophyll a

CNI Close-neighbor-interchange

Cur Curacin

Cy Cyclisation

CyanoGEBA Genomic Encyclopedia of Bacteria and Archaea

DH Dehydratase

Dhoya 2,2-dimethyl-3-hydroxy-7-octynoic acid

DNA Deoxyribonucleic acid

E Epimerization

EDTA Ethylenediaminetetraacetic acid

Epo Epothilone

EPS Extracellular polymeric substances

ER Enoyl reductase

F Formlyation

GC Gas chromatograph

xxxii FCUP


HMM Hidden Markov Model

Hmoya 3-hydroxy-2-methyl-7-octynoic acid

HSAF Heat stable antifungal factor

ITS Internal transcribed spacer

Jam Jamaicamide

Kirr Kirromycin

KR Ketoreductase

KS Ketosynthase

LC-MS Liquid chromatography - mass spectrometry

LEGE Laboratório de Ecotoxicologia Genómica e Evolução

Lnm Leinamycin

Lovas Lovastatin

m/z mass-to-charge ratio

Mcy Microcystin

ME Minimum Evolution

MeOH Methanol

ML Maximum likelihood

Mo Molybdenium

MS Mass spectrometry

MS/MS Tandem MS

Mt N-methyllation

Mta Myxothiazol

Mxa Myxalamide

Myc Mycosubtilin

N2 Dinitrogen

NaCl Sodium chloride

NaPDoS Natural product domain seeker

NCBI National center for biotechnology information

Nda Nodularin

NH3 Ammonia

NJ Neighbor - Joining

NMR Nuclear magnetic resonance

Nos Nostopeptolide

NRP Non-ribosomal peptide

NRPS Non-ribosomal peptide syntethase

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast xxxiii

ORF Open reading frame

Ox Oxidoreductase

PC Phycocyanin

PCP Peptidyl carrier protein

PCR Polymerase chain reaction

PE Phycoerythrin

Phs Phosphitricin

PK Polyketide

PKS Polyketide synthase

PP1 Protein phosphatase 1

PP2A Protein phosphatase 2A

PSP Paralytic shellfish poison

Pvd Pyoverdin

R Reduction

rDNA Ribosomal DNA

RNA Ribonucleic acid

RP Ribosomal peptide

RPS Released polysaccharides

RT-PCR Reverse transcription polymerase chain reaction

RuBisCO Ribulose-1,5-bisphosphate carboxylase/oxygenase

Saf Saframycin

SEM Scanning electron microscopy

Srf Surfactin

SST Sea surface temperature

Sti Stigmatellin

Sxt Saxitoxin

TDC Thermal conductivity detector

TE Thioesterase

TEM Transmission electron microscopy

Tyc Tyrocidine

UV Ultra violet

V Vanadium

Vir Virginiamycin

xxxiv FCUP


CHAPTER I

General Introduction

FCUP

Characterization of cyanobacteria isolated from the Portuguese coast 3

1. Introduction

1.1 Cyanobacteria

Cyanobacteria are ancient photosynthetic prokaryotes that are among the most

successful and oldest forms of life (Schopf, 2000). It is widely accepted that they were

the first group of organisms with the capacity to perform oxygenic photosynthesis, a

process that radically changed the atmosphere and the living conditions on Earth

(Whitton & Potts, 2000; Shi & Falkowski, 2008). Geological records indicate that the

biochemical capacity to use water as electron donor for photosynthesis arose in our

planet’s history at least 2450-2320 million years ago, in a common ancestor of

cyanobacteria (Tomitani et al., 2006; Knoll, 2008), and it is estimated that nowadays

cyanobacteria still contribute up to 30% of the annual oxygen production (Cavalier-

Smith, 2006; DeRuyter & Fromme, 2008). Cyanobacteria are also considered the

photosynthetic ancestors of chloroplasts, a process that occurred by an endosymbiotic

event (Bergman et al., 2008b). This hypothesis has been supported by solid evidences,

such as phylogenetic analysis that showed that the chloroplasts form a monophyletic

lineage closely related to cyanobacteria in particular to the nitrogen-fixing filamentous

strains (Deusch et al., 2008; Falcón et al., 2010; Ochoa de Alda et al., 2014).

Cyanobacteria play a key role in the global carbon cycle as important primary

producers and the diazotrophic strains are fundamental to the nitrogen cycle, mainly in

oceans (Díez et al., 2007; Knoll, 2008; Díez & Ininbergs, 2014). Moreover, they can

also form symbiotic associations with a range of eukaryotic hosts including plants,

fungi, sponges and protists, providing combined nitrogen and/or carbon to the host

(Adams & Duggan, 2008; Bergman et al., 2008a).

During evolution, a complexity and variety of morphologies as well as eco-

physiological properties emerged, placing cyanobacteria among the more diverse

group of prokaryotes (Shi & Falkowski, 2008), occupying a wide variety of

environments including freshwater, terrestrial, and marine habitats. They can also be

found in extreme habitats such as hot springs, deserts, hypersaline and polar lakes.

The ability to colonize these ecosystems can be attributed to their morphological and

metabolic versatility (Potts, 1999; Miller & Castenholz, 2000; Whitton & Potts, 2000).

Cyanobacteria display different morphologies, including unicellular, colonial and

filamentous forms. Some filamentous strains have the ability to produce hormogonia,

4 FCUP


chains of 5-15 cells that can be distinguished from the parent filament by differences in

size, shape or motility. Their primary function is short-distance dispersal (migrating

phase) but they are also essential for host infection in cyanobacteria symbioses

(Whitton & Potts, 2000; Castenholz, 2001; Maldener et al., 2014). Moreover, some

filamentous cyanobacteria can differentiate heterocysts, cells specialized in nitrogen

fixation, and produce akinetes - “resting cells” (Castenholz, 2001).

Cyanobacteria display a wide variety of colours, ranging from blue-green to

violet/brown, due to the different combinations of photosynthetic pigments: chlorophyll

a, carotenoids and phycobiliproteins (allophycocyanin, phycocyanin, and phycoerythrin)

(DeRuyter & Fromme, 2008). Chlorophyll a and phycobiliproteins are the predominant

pigments, however, some prochlorophytes (e. g. Prochloron didemni, Prochlorothix

hollandica, Prochlorococcus marinus), possess chlorophyll b, and Acaryochloris

marina, even contains chlorophyll d (Post, 2006; Swingley et al., 2008). Recently, the

presence of chlorophyll f was also reported in a filamentous cyanobacteria isolated

from stromatolites (Chen et al., 2010; Chen et al., 2012).

Cyanobacteria possess a unique cell wall that combines characteristics of Gram-

negative bacteria, like the presence of outer membrane and lipopolysaccharides, with a

thick and highly cross-linked peptidoglycan layer similar to Gram-positive bacteria

(Hoiczyk & Hansel, 2000). Furthermore, many strains are able to produce extracellular

polymeric substances (EPS) mainly constituted by heteropolysaccharides. These EPS

can remain associated with the cell surface as sheaths, capsules and slimes or be

released into the surrounding environment as released polysaccharides (RPS) (De

Philippis & Micheletti, 2009; Pereira S. et al., 2009). Several roles can be attributed to

the EPS such as the protection of the cells against desiccation and UV-radiation,

formation of biofilms, and concentration of nutrients and metal ions (Rossi & De

Philippis, 2015). In the costal intertidal zones where cyanobacteria are exposed to

extremely stressful conditions such as desiccation, temperature and salinity

fluctuations and often form cohesive mats, EPS are believe to play both protective and

structural roles (Stal, 2000; Díez et al., 2007).

Cyanobacteria are also known as a prolific source of secondary metabolites. Some

of these compounds are toxic to a wide array of organisms, including humans

(Wiegand & Pflugmacher, 2005; Dittmann et al., 2013). Cyanotoxins are mainly

produced by cyanobacteria that form water-blooms induced by increased levels of

nutrients (phosphorus, nitrogen) in close aquatic ecosystems due to anthropogenic

influence. In the aquatic ecosystem these toxins accumulate within cyanobacterial

cells, but can be released in high concentrations during cell lysis causing countless

deaths among wild and domestic animals all over the world (Sivonen & Jones, 1999).

FCUP


Swimming in water bodies, drinking water or consuming dietary supplements that have

accumulated the toxins are also risky to human and animals (Dittmann et al., 2013).

The most fatal case of human intoxication was reported from a hemodialysis unit in

Caruaru (Brazil), where several patients died with symptoms of neurotoxicity and

hepatotoxicity after exposure to contaminated water (Jochimsen et al., 1998).

Understandably, the majority of early cyanobacterial metabolites research focused on

toxins due to their obvious deleterious effects on ecosystem equilibrium and human

health. Nevertheless, cyanobacteria can also produce compounds with potential for

biotechnological applications, in particular in the pharmacological field. A wide range of

secondary metabolites exhibiting promising activities, such as anticancer, antibacterial,

antiviral and antifungal have been reported (Nunnery et al., 2010; Singh et al., 2011;

Shishido et al., 2015). Among marine cyanobacteria, the most prolific sources for the

discovery of secondary metabolites have been the tropical species. It has been

reported that a single genus can produce an array of secondary metabolites with

different structures classes and bioactivities, for e.g. the genus Moorea, formally known

as Lyngbya (Engene et al., 2012), produces jamaicamides (Edwards et al., 2004),

malyngamides (Cardellina et al., 1979) and apratoxins (Gutierrez et al., 2008).

From a biotechnological perspective cyanobacteria are receiving a growing attention

as promising “low cost” microbial cell factories (e.g. for the sustainable production of

secondary metabolites and biofuels) due to their simple nutritional requirements, their

metabolic plasticity and availability of tools for their genetic manipulation (Wase &

Wright, 2008; Lindblad et al., 2012).

1.2 Taxonomy of cyanobacteria

Traditionally, cyanobacteria were mainly classified based on their morphological

characteristics, according to the International Code of Botanical Nomenclature.

However, since their prokaryotic nature was revealed, a classification more consistent

with prokaryotic systematics was included by Rippka (1979) and later on by Castenholz

(2001). Currently, the taxonomy of these organisms is still a controversial subject, with

two prevailing approaches, the so-called botanical (Geitler, 1932; Komárek &

Anagnostidis, 1989; Komárek & Anagnostidis, 1998; Komárek & Anagnostidis, 2005)

and the bacterial (Rippka et al., 1979; Castenholz, 2001).

According to the botanical approach, cyanobacteria are classified into five Orders

(Geitler, 1932; Komárek & Anagnostidis, 1989; Komárek & Anagnostidis, 1998;

Komárek & Anagnostidis, 2005) that broadly coincide with the Subsections of the

6 FCUP


bacterial one (Rippka et al., 1979; Castenholz, 2001): Chroococcales (Subsection I),

Pleurocapsales (Subsection II), Oscillatoriales (Subsection III), Nostocales (Subsection

IV) and Stigonematales (Subsection V).

Briefly, members of Chroococcales (Subsection I) are unicellular cyanobacteria that

reproduce by equal binary fission or by budding. Cells are spherical, ellipsoidal or rod-

shaped (0.5-30 µm diameter), fission occurs in one or more successive planes and

cells are single or in aggregates. Colonial forms are held together by mucilage or

multilaminated sheath layers (Castenholz, 2001). Pleurocapsales (Subsection II)

comprises the unicellular cyanobacteria that can undergo multiple fission, leading to

the formation of small, easily dispersed cells called baeocytes, a feature that

distinguishes them from other groups of cyanobacteria (Castenholz, 2001).

Oscillatoriales (Subsection III), include filamentous cyanobacteria in which the

trichomes are composed only by vegetative cells and are uniseriate since undergo

binary fission in a single plane. The terminal cell of the trichome may be tapered, with

or without a cap or calyptra. The trichome diameter may range from 1 to >100 µm and

it can be surrounded by a sheath (Castenholz, 2001). Nostocales (Subsection IV)

comprise filamentous cyanobacteria that divide exclusively by binary fission in one

plane and some of the vegetative cells can differentiate into heterocysts or akinetes. In

the absence of combined nitrogen a small fraction of the vegetative cells differentiate

into heterocysts that can be terminal or intercalar (Castenholz, 2001). Stigonematales

(Subsection V) include filamentous cyanobacteria that are able to divide by binary

fission in multiple planes, displaying true branching. Longitudinal and oblique cells

divisions occur in addition to transverse cell divisions and this results in true branching

and in multiseriate trichomes (two or more rows of cells) (Castenholz, 2001) (for a

summary see Table 1).

Another important discriminatory taxonomic character is the thylakoids arrangement,

a feature that can only be visualized using transmission electron microscopy. This

arrangement can be parietal, radial or irregular (Liberton & Pakrasi, 2008). The only

known cyanobacterium that lacks thylakoids is the unicellular Gloeobacter violaceus

(Nakamura et al., 2003). In terms of ultrastructure, different sub-cellular compartments

and inclusion bodies can also be observed in cyanobacteria cells, e.g. the

carboxyssomes which are polyhedral bodies that contain enzymes involved in carbon

fixation (RuBisCO) and a variety of reserve materials can also be visible as

cytoplasmatic inclusions, such as glycogen, polyphosphate, and cyanophycin

(nitrogen) (Castenholz, 2001). The gas vesicles, occurring mainly in planktonic strains,

provide buoyancy allowing the organisms to perform vertical migrations.

http://en.wikipedia.org/wiki/Carbon_fixation

http://en.wikipedia.org/wiki/Carbon_fixation

FCUP


Table 1. General characteristics of the cyanobacterial Orders/Subsections.

In the last decades, the advances in molecular biology lead to the use of molecular

markers to provide insight into the identification and to infer phylogenetic relationships

between cyanobacteria. For cyanobacteria as for other bacteria, the small subunit

ribosomal RNA (16S rRNA) gene sequence is the most commonly used as molecular

marker (Castenholz, 2001). The 16S rRNA gene is reliable and a vast number of

sequences are available for comparisons in the databases. However, this marker have

limited resolution at the species level, due to its slow evolution (Konstantinidis & Tiedje,

2007). Consequently, several other less conserved markers providing complementary

information to the 16S rRNA gene are also used (Neilan et al., 1995; Rajaniemi et al.,

2005; Sherwood & Presting, 2007; Singh et al., 2013), for instance the genes of the

phycocyanin operon encoding the two phycobiliprotein subunits (cpcB and cpcA) and

the internal transcribed spacer region (ITS) separating the 16S and 23S ribosomal

genes (Neilan et al., 1995; Rocap et al., 2002; Haverkamp et al., 2008). The 16S-23S

ITS region is highly variable and have demonstrated to be useful for phylogenetic

studies and identification of new taxa (Rocap et al., 2002; Casamatta et al., 2006). The

16S-23S spacer region contains many conserved domains (D1-D5, Box A, Box B) and

variable regions (V1-V3). In addition, this region often contains one or two tRNA genes

(tRNAGlu, tRNAAla or both tRNAAla and tRNAIle) (Iteman et al., 2000). In addition, the

Order/

Subsection General characteristics Genera examples

Unicelullar

Single cells or colonial

aggregates

Chroococcales/ Subsection I

Reproduction by binary fission (one or more planes) or by budding.

Cyanothece

Gloeothece

Synechocystis

Pleurocapsales/ Subsection II

Reproduction by multiple fission with production of small daughter cells - baeocytes, or by multiple plus binary fission.

Chroococcidiopsis

Myxosarcina

Pleurocapsa

Filamentous

Trichome (filament) of

cells: uniseriate or multiseriate

Oscillatoriales/ Subsection III

Reproduction by binary fission in one plane; trichomes composed exclusively by vegetative cells.

Leptolyngbya

Lyngbya

Oscillatoria

Pseudanabaena

Nostocales/ Subsection IV

Reproduction by binary fission in one plane; trichomes composed of vegetative cells that can differentiate into heterocysts or akinetes.

Anabaena

Calothrix

Nostoc

Stigonematales/

Subsection V

Reproduction by binary fission in more than one plane originating multiseriate trichomes (true branching); trichomes composed of vegetative cells that can differentiate into heterocysts or akinetes.

Chlorogloeopsis

Fisherella

Stigonema

8 FCUP


nitrogenase structural genes (see below), in particular nifH (encoding the dinitrogenase

reductase) are also widely used for nitrogen fixing strains/communities (Singh et al.,

2013).

Overall, the classification of cyanobacteria is still a complex task, since there are

several species defined only by morphology, as well as many cyanobacterial gene

sequences without a corresponding morphological description (Castenholz, 2001).

There are numerous species morphologically indistinguishable but that do not share a

common evolutionary history (molecular phylogeny is more diverse than

morphological), therefore the biodiversity is underestimated using the criterion of

morphological variability only. This discrepancy resulted in “cryptic species” (distinct

species classified as a single species) (Casamatta et al., 2003). In addition, the

morphological characters can change in different environmental or growth conditions

and even be lost during cultivation (morphological plasticity) (Komárek, 2014).

Consequently, a polyphasic approach, combining morphological, ecological and

molecular data is considered the best one for the determination and description of

cyanobacterial taxa (Garcia-Pichel, 2008; Komárek, 2010).

1.3 Nitrogen fixation

In cyanobacteria, as in any diazotrophic bacteria, the reduction of dinitrogen (N2) to

ammonia (NH3) is carried out by an enzymatic complex - the nitrogenase - and the

process is coupled with the formation of molecular hydrogen (Berman-Frank et al.,

2003).

Nitrogenase is a highly conserved enzyme complex consisting of two components, a

dinitrogenase (iron-molybdenum protein) and a dinitrogenase reductase (iron protein)

(Figure 1). Dinitrogenase is a α2β2 heterotetramer, and the α and β subunits are

encoded by nifD and nifK genes, respectively. The dinitrogenase reductase, encoded

by nifH, is a homodimer (Orme-Johnson, 1992; Berman-Frank et al., 2003). In addition,

some diazotrophic organisms synthesize alternative dinitrogenases that contain

vanadium (V) or iron, instead of molybdenium (Mo) (Eady, 1996). In cyanobacteria the

alternative nitrogenases are rare but the presence of a vanadium containing enzyme

was demonstrated in some Anabaena strains (Thiel, 1993; Boison et al., 2006).

Similarly to others organisms, these cyanobacteria also possess the “conventional” Mo-

containing nitrogenase, but under Mo deficiency they express the V-nitrogenase. It is

known that alternative nitrogenases are less efficient than the Mo-enzyme in N2

reduction (Stal & Zehr, 2008).

FCUP


Fig.1 - Schematic representation of nitrogenase enzymatic complex. Adapted from Tamagnini et al. (2007).

Since the nitrogenase is irreversibly inactivated by molecular oxygen, nitrogen

fixation occurs in anoxic or microaerobic conditions. Since cyanobacteria are the only

diazotrophs that produce O2 as a by-product of the photosynthesis, they had to develop

strategies to protect their nitrogenases. The main strategies are usually the temporal

and spatial separation, but there are other mechanisms (for more details on this subject

see Berman-Frank et al., 2003). Temporal separation (day/night) between

photosynthetic oxygen evolution and nitrogen fixation seems to be the most common

strategy adopted by non-heterocystous cyanobacteria (Huang et al., 1999; Stal, 2000;

Berman-Frank et al., 2003), while spatial separation is achieved in filamentous strains

that are able to differentiate heterocysts (Böhme, 1998; Stal, 2000; Berman-Frank et

al., 2003). Basically, the heterocyst has a thick cell wall that limits the diffusion of

oxygen, and it lacks photosystem II activity (Stal, 2000). In the marine filamentous non-

heterocystous Trichodesmium spp., a spatial separation occurs without any obvious

cellular differentiation and only a small fraction of cells along the filament are capable

of nitrogen fixation (Bergman et al., 1997; Bergman et al., 2013).

There are diverse cyanobacteria in terrestrial and aquatic ecosystems that can fix N2

and generally they are regarded as the major N2-fixing organisms in the open ocean

(Zehr, 2011). In the intertidal zones, cyanobacteria have also an important role in N2

fixation in microbial mats which are often nutrient-poor and nitrogen depleted

environments (Severin et al., 2010). The identification of N2-fixing cyanobacteria strictly

from morphology is an impossible task, with the obvious exception of filamentous

heterocyst-forming cyanobacteria, and for this reason, a genetic approach that target

the genes encoding the nitrogenase has been used to detect putative N2-fixing strains

(Zehr et al., 1998; Zehr, 2011). As mentioned previously, the structural genes encoding

the nitrogenase, nifHDK, with particular focus on nifH are often used as molecular

markers (Singh et al., 2013) and its presence and expression is also been used to

estimate the contribution of certain microbial mats to N2 fixation (Short & Zehr, 2005).

10 FCUP


1.4 Cyanobacterial secondary metabolites

Cyanobacterial secondary metabolites display a wide range of chemical diversity

(peptides, polyketides and alkaloids) and may have different bioactivities, such as anti-

cancer, anti-fungal, neurotoxic and hepatotoxic. This diversity possibly reflects the

group long evolutionary history and their adaptation to different ecosystems (Méjean &

Ploux, 2013).

Marine environments remain a major source for an unmeasured biological and

chemical diversity and, during the last few decades, marine cyanobacteria have been

increasingly recognized as a prolific source of secondary metabolites (Jones et al.,

2009; Tan, 2010; Tan, 2013a). Nearly 800 compounds have been identified in marine

cyanobacteria, with the filamentous Oscillatoriales (Subsection III) contributing with

almost half of the reported compounds (49%), Nostocales (Subsection IV) with 26%,

and Stigonematales (Subsection V) with merely 4%. Among the unicellular, the

Chroococcales (Subsection I) contribute 15% and the Pleurocapsales (Subsection II)

with about 6% (Gerwick et al., 2008; Jones et al., 2010). In part, this can be explained

by the fact that the filamentous cyanobacteria grow at relatively high densities in

coastal ecosystems forming macroscopic clumps, whereas the unicellular

cyanobacteria usually need to be cultivated to produce enough biomass for chemical

and biological studies (Leão et al., 2013a).

Among the cyanobacterial secondary metabolites, non-ribosomal peptides (NRPs)

and polyketides (PKs), synthesized by the modular multienzymatic complexes non-

ribosomal peptide syntethases (NRPSs) and polyketide synthases (PKSs), respectively

(Barrios-Llerena et al., 2007; Wang et al., 2014), are particularly relevant. PKSs can be

divided in three major classes: type I PKSs that are multifunctional enzymes organized

in modules, each of one harbouring a set of distinct, non-iteratively acting activities

responsible for the catalysis of one cycle of polyketide chain elongation (e.g.

erythromycin A); type II PKSs that are multienzyme complexes that carry a single set

of iteratively acting activities (e.g. tetracenomycin); and type III PKSs, that are

homodimeric enzymes that essentially are iteratively-acting condensing enzymes (e.g.

flavolin). Type I and II PKSs use acyl carrier protein (ACP) to activate the acyl-CoA

substrates whereas type III PKSs act directly on the acyl-CoA substrates, independent

of ACP (Shen, 2003; Cheng et al., 2009). The majority of the bioactive metabolites

isolated so far from cyanobacteria are type I PKs, NRPs or a hybrid of these two. Type

I PKS and NRPS are characterized by a modular organization in which each module is

responsible for the incorporation of a structural carboxylic acid or amino acid unit,

FCUP


respectively. A minimal type I PKS module consists of a ketosynthase (KS),

acyltransferase (AT), and acyl carrier protein (ACP) domains. Additional domains such

as ketoreductase (KR), dehydratase (DH) and enoyl reductase (ER) can also be

present that will lead to different reduction states of the incorporated keto groups. A

minimal NRPS module is composed of an amino acid-activating adenylation (A)

domain, a peptidyl carrier protein (PCP) domain for thioesterification of the activated

amino acid and a condensation (C) domain for transpeptidation between the aligned

peptidyl and amino acyl thioesters. Along with these domains, additional domains

[epimerization (E), oxidoreductase (Ox), cyclisation (Cy), N-methyllation (Mt),

formlyation (F) and reduction (R)] can also be present, introducing structural diversity

by modifying the growing chain (Figure 2) (Barrios-Llerena et al., 2007; Wase & Wright,

2008; Kehr et al., 2011). Although, the majority of the compounds are synthesized from

NRPS (or mixed with PKS), cyanobacteria are also known to produce a group of

ribosomally synthesized and post translationally modified peptides (ribosomal peptides

- RPs), such as cyanobactins and microviridins (Sivonen et al., 2010; Ziemert et al.,

2010).

Fig.2 - Schematic representation of enzymatic domains in polyketide synthases (a) and nonribosomal peptide

synthetases (b). C: Condensation domain; A: Adenylation domain; PCP: Peptidyl carrier protein; MT: Methyltransferase;

E: Epimerase; AT: Acyltransferase; ACP: Acyl carrier protein; KS: Ketosynthase; KR: Ketoreductase; DH: Dehydratase;

ER: Enoyl reductase; TE: Thioesterase. Adapted from Kehr et al. (2011).

12 FCUP


Some examples of secondary metabolites produced by cyanobacteria and their

biological activities are given in Table 2. The diversity of their chemical structures is

depicted in Figure 3.

Table 2. Examples of cyanobacterial secondary metabolites and respective biological activities.

Compound Cyanobacteria Genus

Bioactivity Reference

Aeruginosins Planktothrix

Microcystis

Anticancer Ishida et al., 2009

Anatoxin-a/

Homoanatoxin-a

Anabaena

Oscillatoria

Neurotoxin Méjean et al., 2010

Rantala-Ylinen et al., 2011

Apratoxins Lyngbya Anticancer Grindberg et al., 2011

Aurilide B, C Lyngbya Anticancer Han et al., 2006

Bisebromoamide Lyngbya Anticancer Teruya et al., 2009

Carmaphycins A, B Symploca Anticancer Pereira et al., 2012

Coibamide A Leptolyngbya Anticancer Medina et al., 2008

Cryptophycin Nostoc Anticancer Magarvey et al., 2006

Curacin A Lyngbya Anticancer Chang et al., 2004

Cyanovirin-N Nostoc Anti-viral Boyd et al., 1997

Dolastatin 10 Symploca Anticancer Luesch et al., 2001

Gallinamide A Schizothrix Anti-malarial Linington et al., 2008

Hassalidins Anabaena

Aphanizomenon

Nostoc

Antifungal Vestola et al., 2014

Hectochlorin Lyngbya Anticancer Ramaswamy et al., 2007

Hoiamides Lyngbya

Symploca

Oscillatoria

Neurotoxin (neuromodelating

agent)

Pereira A. et al., 2009

Choi et al., 2010

Jamaicamide Lyngbya Neurotoxin (neuromodelating

agent)

Edwards et al., 2004

Largazole Symploca Anticancer Taori et al., 2008

Lyngbyatoxins Lyngbya Dermatotoxin Edwards & Gerwick, 2004

Malyngamide 2 Lyngbya sp. Anti-inflammatory Malloy et al., 2010

Microcystin Microcystis

Planktothrix

Anabaena

Hepatotoxin Tillett et al., 2000

Rouhiainen et al., 2004

Nodularin Nodularia Hepatotoxin Moffitt & Neilan, 2004

Saxitoxin Anabaena

Aphanizomenon Neurotoxin Kellmann et al., 2008b

Somoscystinamide A Lyngbya Anticancer Wrasidlo et al., 2008

Symplostatin 4 Symploca sp. Anti-malarial Conroy et al., 2010

Veraguamides Oscillatoria Anticancer Mevers et al., 2011

http://pubs.acs.org/doi/abs/10.1021/np8003529

FCUP


Fig.3 - Examples of chemical structures of cyanobacterial metabolites (for more information see also Table 2).

14 FCUP


As mentioned previously, cyanobacteria can produce a variety of toxins

(cyanotoxins) and the characterization of cyanotoxin synthetase gene clusters allowed

the development of molecular methods to detect and differentiate toxic cyanobacterial

strains (Jungblut & Neilan, 2006; Kellmann et al., 2008b). The majority of the

cyanotoxins are associated with cyanobacteria from fresh and brackish waters

(Sivonen & Börner, 2008) but some marine cyanobacteria are known to produce toxins,

namely dermatotoxins (Codd et al., 2005). Moreover, it was also demonstrated that

extracts of marine Synechocystis and Synechococcus strains collected from

Portuguese waters were toxic to marine invertebrates (Martins et al., 2007).

Cyanotoxins can be classified in four functional groups according to their primary

target organ or effects, being designated as hepatotoxins (e.g. microcystin, nodularin),

neurotoxins (e.g. anatoxin, saxitoxin), cytotoxins (e.g. cylindrospermopsin), and

dermatotoxins (e.g. aplysiatoxin, lyngbyatoxin A) (Wiegand & Pflugmacher, 2005;

Dittmann et al., 2013).

The hepatotoxins (microcystin and nodularin) and neurotoxins (saxitoxin) are

between the most problematic toxins (Codd et al., 2005). Microcystin and nodularin are

cyclic peptides (Figure 4a and 4b) and are both inhibitors of the eukaryotic protein

phosphatases 1 and 2A (PP1, PP2A) (Runnegar et al., 1995). While microcystin is

produced by different genera including Microcystis, Anabaena, Planktothrix and Nostoc

(Sivonen & Jones, 1999), nodularin seems to be restricted to the genus Nodularia

(Moffitt & Neilan, 2001).

Fig.4 - Chemical structures of microcystin-LR (a), nodularin (b) and saxitoxin (c). Adapted from Dittmann et al. (2013).

Microcystin is the most extensively studied cyanobacterial secondary metabolite,

due to its distribution and toxicity. About 80 structural variants have been identified,

mostly differing in the amino acids positions X and Z (Sivonen & Jones, 1999), being

microcystin-LR (L-leucine (X) and L-arginine (Z) as the variable amino acids) the most

FCUP


common isoform, whereas only a few variants of nodularins have been identified

(Beattie et al., 2000).

The gene cluster involved in the biosynthesis of microcystin was identified in

Microcystis aeruginosa (Nishizawa et al., 2000; Tillett et al., 2000) making it the first

cyanobacterial hybrid PKS/NRPS cluster to be characterized. This cluster has also

been identified in several strains such as Planktothrix spp. (Christiansen et al., 2003)

and Anabaena strain 90 (Rouhiainen et al., 2004). The 55 kilobase (kb) cluster of

Microcystis aeruginosa PCC 7806 comprises ten open reading frames (ORF) (mcyA to

J) (Figure 5). mcyA to mcyC encode five NRPS modules, mcyD encodes two modules

type I PKS, and mcyE and mcyG code for hybrid NRPS-PKS proteins (Tillett et al.,

2000). The mcy cluster in Planktothrix agardhii also contains the gene mcyT that

encodes a second thioesterase domain (Christiansen et al., 2003).

Fig.5 - Structural organization of the microcystin biosynthetic gene cluster in Microcystis aeruginosa. Genes coding for

polyketide synthase (hatched), nonribosomal peptide synthetase (black), and tailoring enzymes (w hite) are indicated.

Adapted from Moffitt & Neilan (2004).

Nodularin, other hybrid PKS/NRPS, is structurally closely related to microcystin

sharing a highly similar biosynthetic pathway. This cluster has been particularly studied

in Nodularia spumigena, 48 kb comprising nine ORFs (ndaA to ndaI) (Figure 6). ndaA

and ndaB encode three NRPS modules, ndaD encodes two PKS modules, ndaC and

ndaF encode hybrid NRPS-PKS proteins and ndaE and ndaGHI encode tailoring

enzymes (Moffitt & Neilan, 2004).

Fig.6 - Structural organization of the nodularin biosynthetic gene cluster in Nodularia spumigena. Genes encoding

polyketide synthase (hatched), nonribosomal peptide synthetase (black), and tailoring enzymes (w hite) are indicated.

Adapted from Moffitt & Neilan (2004).

16 FCUP


Saxitoxin is a neurotoxin alkaloid (Figure 4c), which are also known as paralytic

shellfish poisons (PSPs) due to their occurrence and association with seafood

(Dittmann et al., 2013). Saxitoxins are among the most potent toxins and are

considered a serious toxicological health risk to humans, animals and ecosystems

worldwide (Kaas & Henriksen, 2000). These toxins block the voltage-gated sodium and

calcium channels (Kao & Levinson, 1986; Wiegand & Pflugmacher, 2005), which

prevent the transduction of neuronal signals, and thus cause muscular paralysis

(Kellmann et al., 2008a). The saxitoxin production has been reported for several

cyanobacteria genera, notably Aphanizomenon, Anabaena, and Lyngbya (Dittmann et

al., 2013) and more than 30 analogues have been identified (Kellmann et al., 2008a).

The gene cluster involved in the saxitoxin biosynthesis was initially identified in

Cylindrospermopsis raciborskii (Kellmann et al., 2008b) and subsequently in several

other strains (Mihali et al., 2009). Unlike NRPS and PKS biosynthesis, the saxitoxin

cluster encodes a series of monofunctional enzymes responsible for building the

saxitoxin core and tailoring enzymes (D'Agostino et al., 2014). In Cylindrospermopsis

raciborskii, this cluster (35 Kb) encodes genes involved in the biosynthesis, regulation

and export of saxitoxin. This analysis also revealed that it contains a PKS-like

structure, sxtA, with four catalytic domains (Kellmann et al., 2008b) (Figure 7).

Fig.7 - Structural organization of the saxitoxin biosynthetic gene cluster in Cylindrospermopsis raciborskii. Adapted from

Kellmann et al. (2008b).

Regarding cyanobacterial metabolites with therapeutic potential, most of the

research has been focused on compounds with anticancer activity. The cytotoxic

effects on human tumour cell lines of a high number of compounds isolated from

different strains, in particular from marine cyanobacteria, has been demonstrated

(Gutierrez et al., 2008; Mevers et al., 2011; Costa et al., 2012; Costa et al., 2014).

Molecular mechanisms of anticancer marine cyanobacterial natural products include

the inhibition of microtubule dynamics (dolastatins 10), inhibition of actin filaments

(bisebromoamide), histone deacetylase inhibitors (largazole), proteasome inhibitors

(carmaphycins A and B) or induction of apoptosis in cancer cells (apratoxins) (Nunnery

et al., 2010; Tan, 2010; Tan, 2013a; Tan, 2013b). The recently approved agent for

anaplastic large cell lymphoma and Hodgkin’s lymphoma, Brentuximab vedotin, was

FCUP


inspired by the marine cyanobacterial metabolite dolastatin 10 (Minich, 2012).

Cyanobacteria have also been reported as a source of neuromodulating molecules

(hoiamides) (Tan, 2013b), antibacterial (carbamidocyclophanes) (Bui et al., 2007;

Singh et al., 2011), and antiviral compounds (cyanovirin-N) (Boyd et al., 1997; Dey et

al., 2000; Singh et al., 2011). One of the most potent marine cyanobacterial antimalaria

compounds reported is symplostatin 4 (Nunnery et al., 2010; Tan, 2013b) and a

number of secondary metabolites have shown anti-inflammatory activity, such as

malyngamides F acetate and 2 (Tan, 2013b).

In summary, several studies have revealed the metabolic capabilities of

cyanobacteria, in particular of the marine strains, for the production of new therapeutic

natural products. However, despite the large number of cyanobacterial secondary

metabolites identified, there is only a small fraction for which the biosynthetic genes

have been identified (e.g. microcystin, nodularin, saxitoxin, aeruginosin,

cylindrospermopsin, jamaicamide, curacin A, barbamide, lyngbyatoxin, apratoxin,

nostopeptolide, cryptophycin).

1.4.1 Prediction of natural product biosynthetic gene clusters

Synechocystis sp. PCC 6803 was the first cyanobacteria to have the genome fully

sequenced (Kaneko et al., 1996) but, subsequently, 265 cyanobacterial genomes

sequences become available (Joint Genome Institute (JGI), Integrated Microbial

Genomes (IMG) database, June 2015). A recent sequencing project known as

CyanoGEBA (Genomic Encyclopedia of Bacteria and Archaea) (Shih et al., 2013)

significantly increased the number and diversity of sequences covering the

cyanobacteria phylum. Prior to this study, there were no publically available genomes

from Pleurocapsales (Subsection II). Moreover, the authors also assessed the genetic

potential of these cyanobacteria to produce secondary metabolites (presence of PKS,

NRPS and PKS/NRPS hybrids) highlighting their distribution across the phylum (Shih

et al., 2013). Recently, a large-scale analysis of cyanobacterial natural product

pathways was also performed (Calteau et al., 2014). In this study, 452 biosynthetic

gene clusters (190 NRPS, 162 PKS and 100 hybrids gene clusters) from 89 (out of

126) cyanobacteria were identified (Calteau et al., 2014). In total, 286 distinct

NRPS/PKS gene cluster families were found, comprising biosynthetic pathways of

known cyanobacterial compounds but also revealing a large number of orphan

pathways with high drug discovery potential (Calteau et al., 2014). These studies

highlighted the exceptional capacity of cyanobacteria to produce a large diversity of

18 FCUP


secondary metabolites, representing a range of distinct and unrelated biosynthetic

pathways (Shih et al., 2013; Calteau et al., 2014).

The knowledge of the secondary metabolic potential of specific strains can facilitate

the process of natural products discovery (Ziemert et al., 2012). The progress in

sequencing technologies allowed the development of novel screening methods based

on genome sequences. The principle of these genome mining approaches is to identify

genes involved in the biosynthesis of such molecules and to predict the products of the

identified pathways. Therefore, the development of bioinformatic tools for data analysis

was essential, and currently an array of available bioinformatics tools for secondary

metabolite biosynthetic gene clusters analysis are available (Weber, 2014).

Some softwares identify all the biosynthetic gene clusters in a given genome (e.g.

antiSMASH 3, BAGEL3 and SMURF). The Antibiotics and Secondary Metabolite

Analysis SHell (antiSMASH) (http://antismash.secondarymetabolites.org/) (Medema et

al., 2011; Weber et al., 2015) is the most comprehensive tool for the identification and

analysis of secondary metabolite biosynthetic gene clusters in fungi and bacteria. Many

algorithms and tools for the specialized analysis of gene clusters are directly

implemented within antiSMASH. Other softwares are used to identify specific enzyme

classes (e.g. PKS and NRPS pathways). Several bioinformatics tools take advantage

of the typical domain architecture presented by modular PKS and NRPS to predict

domain organization and biosynthetic products (e.g. NaPDoS, NP.Searcher and

SBSPKS), as well as substrate specificities (e.g. PKS/NRPS Analysis and NRPS

predictor2) (Weber, 2014).

The Natural Product Domain Seeker (NaPDoS) (Ziemert et al., 2012)

(http://napdos.ucsd.edu/), employs a phylogeny-based classification system that

extracts and classifies KS (PKS) and C (NRPS) domains from a wide range of

sequence data. Firstly, the domains are identified in genomic or metagenomic data

using a HMM (Hidden Markov Model) and homology based search strategy, and then

the identified domains are integrated into a manually curated phylogenetic tree

providing conclusions on phylogenetically related enzymes and their products (Ziemert

et al., 2012; Weber, 2014). With the PKS/NRPS Analysis tool (Bachmann & Ravel,

2009) (http://nrps.igs.umaryland.edu/) it is possible to investigate the domain type

encoded within an NRPS/PKS protein query sequence. PKS/NRPS analysis also

implements a BLAST-based substrate prediction of NRPS A domains, based on an

extensive database of eight-amino acid specificity-conferring codes extracted from

biochemically characterized NRPS A domains (Bachmann & Ravel, 2009). The A

domain plays a crucial role in the selection of the building blocks and the active site of

the A-domains reveal a “non-ribosomal code” correlating eight amino acids involved in

http://antismash.secondarymetabolites.org/

http://antismash.secondarymetabolites.org/

FCUP


building-up the active sites with their cognate substrates. This allowed to understand

the specificity of these complex enzymes and the prediction of substrate specificities

(Stachelhaus et al., 1999; Challis et al., 2000).

The fast-growing availability of sequenced genomes has contributed to the

knowledge on the biosynthetic pathways of secondary metabolites and in silico

analysis has become an indispensable resource to quickly assess the potential of an

organism to produce a certain metabolite, being an important approach in secondary

metabolites research and drug discovery.

1.4.2 Mass spectrometry (MS) based metabolomics: molecular networking

MS-based metabolomics has become a powerful tool for the detection of

metabolites as well as for the identification of novel ones (Baran et al., 2013). Recently,

a powerful tool – molecular networking – has been applied in order to obtain a global

view of the secondary metabolome of a given organism (Watrous et al., 2012; Yang et

al., 2013; Winnikoff et al., 2014) (Figure 8). Molecular networks are visual

representations of mass-spectral data based on MS-fragmentation similarities. MS-

based molecular networking relies on the observation that compounds with similar

structures share similar MS/MS fragmentation patterns and that molecular families tend

to cluster together within the network. For molecular network generation, firstly the

MS/MS spectra from samples and known standards are collected and then a molecular

network is generated using “cosine scores” which measure relatedness in MS/MS

spectra that are visualized using Cytoscape (Shannon et al., 2003). Within the network,

one node represents one consensus MS/MS spectrum (merging of MS/MS spectra with

nearly identical precursor mass and peak patterns). The relatedness between each

node is defined by an edge (line) and the thickness of the edge defines the degree of

similarity of the MS/MS spectra. These networks allow a simultaneous view of identical

molecules, analogues or compound families. The development of molecular networks

from MS/MS data has proven to be a useful tool for screening organisms that produce

valuable natural products (Watrous et al., 2012; Yang et al., 2013; Winnikoff et al.,

2014).

20 FCUP


Fig.8 - Graphical representation of a traditional molecular netw ork generated w ith cyanobacteria extracts and reference

compounds, displaying different natural products grouped into clusters w ith chemical relatedness.

Green nodes - reference (seed) compounds; pink nodes - samples.

1.5 Cyanobacterial diversity in portuguese waters and

secondary metabolites production

The Portuguese Continental coast has a complex and highly dynamic set of

intertidal ecosystems, with varied topographies, beach morphologies and wave

regimens, exposed to climatic influences from the North Atlantic Ocean and the

Mediterranean Sea (Pontes et al., 2005; Lima et al., 2006; Seabra et al., 2011).

In the last years, several studies on the diversity of cyanobacteria from Portuguese

fresh and estuarine waters have been performed (De Figueiredo et al., 2006; Valério et

al., 2009; Galhano et al., 2011; Lopes et al., 2012), and the majority of these studies

focused on toxins-producing strains (Vasconcelos, 1999; Martins et al., 2009; Lopes et

al., 2010; Martins et al., 2011). In contrast, the diversity of marine cyanobacteria has

been unexplored with only a few reports published previous to this work (Araújo et al.,

2009; Ramos et al., 2010). However, the bioactive potential of some marine

cyanobacterial strains have been investigated, with a couple of studies revealing that

extracts of cyanobacteria could be toxic to marine invertebrates (Martins et al., 2007;

Frazão et al., 2010), whereas others showed that cyanobacterial extracts have high

FCUP


activity in ecologically-relevant bioassays and toxicity against human cancer cell lines

(Leão et al., 2013a; Leão et al., 2013b; Costa et al., 2014).

Due to the importance that cyanobacteria have in the ecosystems equilibrium as

well as to their potential for several biotechnological applications, more studies are

needed to better understand the diversity and metabolic potential of isolates from the

extensive Atlantic Portuguese coastline.

1.6 Objectives

The general goal of this study was to gather knowledge on the diversity of

cyanobacteria isolated from the Portuguese coast as well as to assess their potential to

fix atmospheric nitrogen and to produce secondary metabolites. For this purpose,

cyanobacterial isolates previously collected from the intertidal zones and deposited at

Laboratório de Ecotoxicologia, Genómica e Evolução (LEGE Culture Collection,

CIIMAR, Porto) were:

(i) Characterized at morphological, ultrastructural and molecular level (16S-ITS-

23S sequences and phylogenetic analysis);

(ii) Screened for the potential to fix nitrogen (presence of nif genes and nitrogenase

activity);

(iii) Screened for the presence of genes involved in the production of the

cyanotoxins microcystin (mcyA, mcyE), nodularin (ndaF) and saxitoxin (sxtI);

(iv) Screened for the presence of genes encoding non-ribossomal peptide

synthetase (NRPS) and polyketide synthase (PKS), targeting the A and the KS

domains, respectively;

(v) Assessed for their potential to produce secondary metabolites (sequences

encoding KS and A domains were used to in silico predict PKS and NRPS

systems, and crude extracts were analysed to search for novel or otherwise

interesting compounds).

CHAPTER II

Culture dependent characterization of

cyanobacterial diversity in the intertidal

zones of the Portuguese coast: a polyphasic

study

Brito Â., Ramos V., Seabra R., Santos A., Santos C.L., Lopo M., Ferreira S.,

Martins A., Mota R., Frazão B., Martins R., Vasconcelos V., Tamagnini P. 2012.

Systematic and Applied Microbiology 35: 110-119.

Systematic and Applied Microbiology 35 (2012) 110– 119

Contents lists available at SciVerse ScienceDirect

Systematic and Applied Microbiology

j ourna l ho mepage: www.elsev ier .de /syapm

Culture-dependent characterization of cyanobacterial diversity in the intertidalzones of the Portuguese coast: A polyphasic study

Ângela Britoa,b, Vitor Ramosa,b,c, Rui Seabrab,c, Arlete Santosa,b, Catarina L. Santosa,Miguel Lopoa, Sérgio Ferreiraa, António Martinsc, Rita Motaa,b, Bárbara Frazãob,c,Rosário Martinsc,d, Vitor Vasconcelosb,c, Paula Tamagninia,b,∗

a IBMC – Instituto de Biologia Molecular e Celular, Universidade do Porto, Rua do Campo Alegre 823, 4150-180 Porto, Portugalb Faculdade de Ciências, Departamento de Biologia, Universidade do Porto, Rua do Campo Alegre, Edifício FC4, 4169-007 Porto, Portugalc Interdisciplinary Centre of Marine and Environmental Research (CIIMAR/CIMAR), University of Porto, Rua dos Bragas 289, 4050-123 Porto, Portugald CISA – Centro de Investigac ão em Saúde e Ambiente, Escola Superior de Tecnologia da Saúde, Instituto Politécnico do Porto, Rua Valente Perfeito 322, 4400-330 Vila Nova de Gaia,Portugal

a r t i c l e i n f o

Article history:Received 15 April 2011Received in revised form 21 July 2011Accepted 28 July 2011

Keywords:CyanobacteriaMarine16S rRNA geneNitrogen fixationPhylogenyToxins

a b s t r a c t

Cyanobacteria are important primary producers, and many are able to fix atmospheric nitrogen playing akey role in the marine environment. However, not much is known about the diversity of cyanobacteria inPortuguese marine waters. This paper describes the diversity of 60 strains isolated from benthic habitatsin 9 sites (intertidal zones) on the Portuguese South and West coasts. The strains were characterized bya morphological study (light and electron microscopy) and by a molecular characterization (partial 16SrRNA, nifH, nifK, mcyA, mcyE/ndaF, sxtI genes). The morphological analyses revealed 35 morphotypes (15genera and 16 species) belonging to 4 cyanobacterial Orders/Subsections. The dominant groups amongthe isolates were the Oscillatoriales. There is a broad congruence between morphological and molecularassignments. The 16S rRNA gene sequences of 9 strains have less than 97% similarity compared to thesequences in the databases, revealing novel cyanobacterial diversity. Phylogenetic analysis, based onpartial 16S rRNA gene sequences showed at least 12 clusters. One-third of the isolates are potential N2-fixers, as they exhibit heterocysts or the presence of nif genes was demonstrated by PCR. Additionally,no conventional freshwater toxins genes were detected by PCR screening.

© 2012 Elsevier GmbH. All rights reserved.

Introduction

Continental Portugal has an extensive coastline, of about940 km, facing the North Atlantic Ocean. It is one of the warmestEuropean countries, and its climate is classified as Mediterraneantype Csa in the south (C – warm temperate; s – summer dry; a –hot summer) and Csb in the north (C – warm temperate; s – sum-mer dry; b – warm summer), according to the Köppen’s scheme[23]. The near-shore wave energy has a strong spatial and seasonalvariability [30]. Western and southern coasts are evidently underdifferent wave regimes, with the unsheltered west coast sites expe-riencing higher wave height and power than southern ones, and amoderate decreasing gradient from north to south can be observed.Wave height and power in the winter are also much higher than inthe summer [30], and along the coast it is possible to encounter

∗ Corresponding author at: IBMC – Instituto de Biologia Molecular e Celular,Universidade do Porto, Rua do Campo Alegre 823, 4150-180 Porto, Portugal.Tel.: +351 226074900; fax: +351 226099157.

E-mail address: [email protected] (P. Tamagnini).

different topographies and beach morphologies – rocky beaches,beaches with sand and rocks and sandy beaches with dispersedrocks, resulting in diverse levels of exposure to the prevailing waveregimen.

Cyanobacteria are photosynthetic prokaryotes with a wide geo-graphical distribution that are present in a broad spectrum ofenvironmental conditions [49]. Taxonomy of cyanobacteria is acontroversial subject, with two prevailing approaches, the “botan-ical” and “bacterial”. The reorganized taxonomic revision based onthe botanical code uses morphological, biochemical and molec-ular characters [19–21]. After the recognition of the prokaryoticfeatures of cyanobacteria, Rippka et al. [36] published a bacterio-logical taxonomy based on morphological, biochemical and geneticcharacters of axenic cultures, while Bergey’s Manual of SystematicBacteriology [5] uses a phylogenetic approach primarily based on16S rRNA sequence comparisons. In summary, cyanobacteria canbe classified into five Subsections [5,36] that broadly coincide withOrders of the botanical approach [19–21]. Cyanobacteria belongingto subsections I (Chroococcales) and II (Pleurocapsales) are unicel-lular, but while the organisms in subsection I divide exclusivelyby binary fission, the ones from subsection II can also undergo

0723-2020/$ – see front matter © 2012 Elsevier GmbH. All rights reserved.doi:10.1016/j.syapm.2011.07.003

dx.doi.org/10.1016/j.syapm.2011.07.003

http://www.sciencedirect.com/science/journal/07232020

http://www.elsevier.de/syapm

mailto:[email protected]

dx.doi.org/10.1016/j.syapm.2011.07.003

Â. Brito et al. / Systematic and Applied Microbiology 35 (2012) 110– 119 111

Table 1Localization and characteristics of the sampling sites.

Sampling site latitude/longitude Wave exposure Beach type

1. Moledo do Minho High RockyN 41◦50′58.68′′/W 8◦52′0.18′′ (Rivermouth)2. S. Bartolomeu do Mar High Sand and rocksN 41◦34′25.59′′/W 8◦47′54.81′′

3. Lavadores High Sand and rocksN 41◦07′45.07′′/W 8◦40′6.88′′

4. Aguda High Sand and rocksN 41◦02′58.35′′/W 8◦ 39′19.22′′

5. Foz do Arelho High SandyN 39◦25′59.79′′/W 9◦13′48.99′′ (Rivermouth)6. Martinhal Intermediate SandyN 37◦01′07.30′′/W 8◦55′36.17′′

7. Burgau Intermediate-low Sand and rocksN 37◦04′15.84′′/W 8◦46′34.97′′

8. Luz Intermediate-low RockyN 37◦05′10.38′′/W 8◦43′31.35′′

9. Olhos d’ÁguaN 37◦05′23.01′′/W 8◦11′27.66′′ Low Sandy

multiple fission producing small easily dispersible cells called baeo-cytes. The subsection III (Oscillatoriales) includes the filamentousstrains without cell differentiation. Subsections IV (Nostocales) andV (Stigonematales) comprise the filamentous strains that are ableto differentiate heterocysts and akinetes. In addition, filaments ofcyanobacteria from subsection V are able to divide in multipleplanes displaying true branching.

Benthic cyanobacteria grow along the shore on different sub-strata, mainly in the intertidal zones, as part of complex multi-taxacommunities, often forming cohesive mats and biofilms. In thesehabitats, they are exposed to a range of daily stresses such asnutrient limitation, high UV-radiation and desiccation [1,6]. Thesuccessful adaptation to these harsh environments is largely dueto their morphological and functional versatility [31]. Cyanobac-teria play a major role in the global carbon cycle as importantprimary producers performing oxygenic photosynthesis, and thediazotrophic taxa are fundamental to the nitrogen cycle, partic-ularly in oceans [18]. In addition to their ecological importance,cyanobacteria are also recognized as being a prolific source of bio-logically active natural products; some of these compounds aretoxic to a wide array of organisms [50]. Nevertheless, little is knownabout the diversity of these organisms along the Portuguese coastwith only a few reports published [e.g. 2,9]. Araújo et al. [2] providedan updated checklist of the benthic marine algae of the northernPortuguese coast, including 26 species of cyanobacteria. However,these authors based their work on new records, literature refer-ences and herbarium data but did not isolate or maintain culturesof the observed specimens.

The aim of this work was to identify the cyanobacteria presentin the intertidal zones of the Portuguese coast using a polyphasicapproach. The isolated specimens are kept at LEGE Culture Collec-tion, and available for further studies. In addition to the assessmentof cyanobacterial diversity, a PCR-based screen for putativediazotrophs and toxin-producers was performed to unveil theirrole(s) in the ecosystem.

Materials and methods

Sampling sites

The sites (Table 1) were selected in order to represent thediversity of the Portuguese coast. Along the coast it is possi-ble to discriminate between rocky beaches (sampling sites 1 and8), beaches with sand and rocks (sampling sites 2, 3, 4, 7) andsandy beaches with dispersed rocks (sampling sites 5, 6 and 9)(Table 1). Several relevant variables: wave power [30], sea surface

temperature [SST [25]], river runoff and other important climaticvariables such as air temperature and precipitation (Instituto deMeteorologia, IP, Portugal) were also taken into account. In brief,West coast sampling sites experience generally higher energeticwave regimes, lower overall mean SSTs and mean air tempera-tures [40], and higher fresh water inputs, both from river runoffand precipitation, than their South coast counterparts.

Cyanobacteria sampling, isolation, and culturing

Biological samples were collected in both summer and autumnof 2006, and spring of 2007. Sample collection was performed byscraping the surface of a wide range of substrates (e.g. seaweeds,rock surfaces, seashells, Sabellaria alveolata reefs) present on barerocks or shallow puddles tidal pools. For the isolate LEGE 06009 seealso [9]. In addition, seawater samples from the surf zone were alsocollected.

Raw biological samples were screened for cyanobacterial spec-imens using a light microscope (Leica DMLB), and subsequentlysubjected to liquid culture enrichment, agar plates streaking ormicromanipulation. Whenever possible single cells/filaments wereisolated and transferred to liquid or solid medium using a Pas-teur pipette [34]. When micromanipulation was found unsuitable,aliquots of the enriched liquid cultures were transferred to liquidmedium or streaked on agar plates. Sea water samples were filteredwith sterile glass fiber filters (GF/C – Ø 47 mm, Whatman), and sub-sequently placed on liquid medium until a visible colony appeared.Single colonies were picked and streaked on agar plates. Newcolonies were transferred into liquid medium. Cultures were main-tained using the following media: MN [34], BG110, BG11 [44], andZ8 [22] supplemented with 25 g L−1 NaCl, further named Z8 25‰.The media were supplemented with B12 vitamin, and when nec-essary with cycloheximide or amphothericin B [34]. The cultureswere kept under 14 h light (10–30 �mol photons m−2 s −1)/10 hdark cycles at 25 ◦C. Cyanobacterial isolates are deposited at LEGECulture Collection (Laboratório de Ecotoxicologia, Genómica eEvoluc ão; CIIMAR, Porto, Portugal). Additionally, the isolate LEGE06123 is also deposited at Culture Collection of Algae and Protozoa(CCAP 1425/1), for details on this organism see [33].

Light and transmission electron microscopy (TEM) andmorphological identification

Cells were observed using a Leica DMLB microscope (LeicaMicrosystems GmbH), images were captured with a Leica ICCA Ana-logue Camera System, and the cells were measured using Qwin

112 Â. Brito et al. / Systematic and Applied Microbiology 35 (2012) 110– 119

Leica software (Leica Microsystems GmbH). Each morphometricparameter was measured 20 times in different individuals.

For TEM studies, cells were collected, centrifuged and processedas described by Seabra et al. [39]. Sections were examined using aZeiss EM 902.

The morphological identification was carried out following thecriteria of Komárek and Anagnostidis [19–21], Waterbury andStanier [47] and by using Bergey’s Manual of Systematic Bacteriol-ogy [4], i.e. whenever the identification differs in the two systems(“botanical” and “bacteriological”), the corresponding form-genusin Bergey’s classification is also mentioned. For PleurocapsalesWaterbury and Stanier [47] classification was followed.

DNA extraction, purification, PCRs and sequencing

Cells were harvested by centrifugation and DNA was extractedusing the Maxwell® 16 System, and the Cell DNA Purifi-cation Kit for Gram-negative bacteria (Promega Corporation)according to the instructions of the manufacturer. DNA frag-ments within the following genes were amplified using theoligonucleotide primers listed on Table 2: 16S rRNA gene(small subunit ribosomal gene), nifK (dinitrogenase � subunit),nifH (dinitrogenase reductase), mcyA and mcyE (microcystinsynthetase), ndaF (nodularin synthetase), and sxtI (saxitoxinbiosynthesis). The 16S rRNA gene was amplified using twoprimer pairs, each including one universal primer and one spe-cific for cyanobacteria [8-27F(universal)/CYA781R(specific), andCYA361F(specific)/1492R(universal)], this allowed us to amplify alonger fragment, and to avoid unspecific amplifications since thecultures are not axenic. For nifK, the first group of primer pairs(see Table 2) was designed for filamentous cyanobacteria, whereasthe second group was designed for unicellular cyanobacteria. PCRreactions were performed using a thermal cycler MyCyclerTM (Bio-Rad laboratories Inc., Hercules, CA, USA) following [45]. The PCRprofiles, after a 2–5 min at 94 ◦C of denaturation step, were the fol-lowing: 16S rRNA gene – 35 cycles of 94 ◦C 1 min, 50 ◦C 1 min, and72 ◦C 1 min 30 s; nifK – 30 cycles of 94 ◦C 1 min, 50 ◦C 1 min, and72 ◦C for 1 min 15 s; mcyA, mcyE, and ndaF – 30 cycles of 95◦ C 1 min30 s, 52◦ C 30 s, and 72 ◦C 50 s; sxtI – 30 cycles of 94◦ C 10 s, 52 ◦C20 s, and 72 ◦C 1 min. In all cases a final extension of 7 min at 72 ◦Cwas performed. Amplification of the nifH fragment was performedaccording to the methods described previously [29]. The PCR prod-ucts were separated by agarose gel electrophoresis using standardprotocols [38]. DNA fragments were isolated from gels using theNZYGelpure Kit (NZYtech, Lda. INOVISA, Lisbon, Portugal), accord-ing to the manufacturer’s instructions. Purified PCR products werecloned into pGEM®-T Easy vector (Promega, Madison, WI, USA), andtransformed into Escherichia coli DH5� competent cells followingthe manufacturer’s instructions, and the methodology described in[33]. Some purified PCR products were directly sequenced at STABVida (Lisbon, Portugal).

Nucleotide sequence accession numbers

Novel sequences associated with this study are avail-able in GenBank under the accession numbers FJ589716,HQ832895–HQ832951, JF708120 and JF708121.

Phylogenetic analysis

In order to integrate the cyanobacterial diversity along the Por-tuguese coast into a broader phylogenetic context, the 16S rRNAgene sequences were analyzed, compared with the currently avail-able databases and used to construct phylogenetic trees. Eachsequence was independently used as the query in a BLAST searchagainst the non-redundant nucleotide database of the National

Centre for Biotechnology Information (NCBI, March 2011). In orderto include full-length sequences and to obtain a reliable overall pic-ture of the cyanobacterial diversity, a number of reference strainsfrom each subsections/orders represented in our samples wereretrieved from GenBank and were included in the following phylo-genetic analyses. The reference strains were selected according tothe Bergey’s Manual of Systematic Bacteriology (2001), and fromthose the ones closely related to our samples were used. A multiplealignment encompassing 16S rRNA gene sequences from the iso-lates, the reference strains and Chloroflexus aurantiacus J-10-fl asthe outgroup was performed using the ClustalW2 algorithm [24],with all the default parameters. Due to the size differences in theamplification products of each isolate, and to avoid the consequentbias effect on the tree construction, this alignment was prunedto an internal core of 655 nucleotides present in all sequences –corresponding to nucleotides 519 to 1172 in E. coli. The phylo-genetic tree was computed using the Maximum-likelihood (ML)methodology [8]. The alignment was imported to Geneious Pro[7], and the tree was build with the PhyML [12] plugin, usingthe HKY85 as the substitution model and 1000 pseudo-replicatesfor the bootstrap analysis, and allowing the software to estimatethe transition/transversion ratio, the proportion of invariable sitesand the gamma distribution parameter with 4 substitution ratecategories.

Results and discussion

This work presents a polyphasic study of cyanobacterial iso-lates from selected intertidal zones along the Portuguese coast,combining the isolation of strains, and their characterization bymicroscopic observations and molecular analysis.

Morphological characterization

To evaluate the diversity of cyanobacteria, nine beaches thatreflect the heterogeneity of habitats present along the conti-nental Portuguese coast were selected (Table 1). Approximately100 cyanobacterial isolates were retrieved, from which 60 werecharacterized and shown to belong to 35 different morphotypes.Fifteen genera and 16 species belonging to four cyanobacterialorders/subsections were identified based on morphological char-acters (Table S1, supplementary material). In terms of the isolate’sspatial distribution no clear pattern was observed, i.e. the dif-ferent cyanobacterial groups are distributed by all the differentbeaches types. However, one must take into account that the num-ber of isolates of certain genera is much higher than others, andthat the different number of isolates obtain for each genus canbe due to their ubiquity or to the fact that they are easier toisolate. The isolation media were selected due to their differentcomposition in order to expose the highest possible diversity. MNmedium revealed the highest diversity, Z8 25‰ was particularlysuccessful for coccoids and Leptolyngbya spp., while BG110 andBG11 did not contribute to a higher diversity than the Z8 25‰,with the exception of Nostoc sp. LEGE 06158. The most represen-tative group among the isolates comprises the nonheterocystousforms, notably the filamentous Oscillatoriales and the unicellularChroococcales. Representatives from the unicellular Pleurocapsaleswere also found, as well as heterocystous taxa: Nostocales (Fig. 1).No true-branching filamentous, Stigonematales, were found. Thenonheterocystous cyanobacteria are, indeed, reported as thepredominant forms in marine environments [3,6,43,46]. This seemsto be also the case for Portugal, the Checklist of benthic marinealgae and cyanobacteria of northern Portugal (“Checklist”) reported26 species of cyanobacteria, 15 of which are Oscillatoriales [2].Strains belonging to the genera Hyella, Myxosarcina, Lyngbya,


Table 2Target genes and oligonucleotide primers used in this study.

Target genes Primer pair a Sequence 5′ → 3′ PCR product expected size (bp) Reference

16S rRNA 8-27F AGAGTTTGATCCTGGCTCAG [48]CYA781R (A) b GACTACTGGGGTATCTAATCCCATT 773 [28]CYA781R (B) GACTACAGGGGTATCTAATCCCTTT [28]CYA361F GGAATTTTCCGCAATGGG 1131 [27]1492R GGTTACCTTGTTACGACTT [48]

nifK nifK0F (A) CAAGGTTCTCAAGGTTGCGTT 1139 [33]nifK1F (B) CAAGGTTCTCAAGGTTGTGTG [33]nifK4R TGTAAGTGGTGGCGATCAAAGA [33]nifK0F (A)/nifK1F (B) See above 407 [33]nifK3′R GGGATGAAGTTGATTTTGCCGTT This worknifk146F CCTGGGAATATCGGGAA 319 This worknifk465R GCCATACAAGTTGTACAGACA This worknifk310F CGTCACTTCAAAGAGCCTT 1084 This worknifk1394R GGCGATCAAAGATAGGATA This worknifH4 TTYTAYGGNAARGGNGG [29]

nifH nifH3 ATRTTRTTNGCNGCRTA 361 [29]nifH2 ADNGCCATCATYTCNCC [29]nifH1 TGYGAYCCNAARGCNGA [29]

mcyA mcyA-Cd1F AAAAGTGTTTTATTAGCGGCTCAT 297 [14]mcyA-Cd1R AAAATTAAAAGCCGTATCAAA [14]

mcyE/ndaF HEPF TTTGGGGTTAACTTTTTTGGGCATAGTC 472 [16]HEPR AATTCTTGAGGCTGTAAATCGGTTT [16]

sxtI sxtI-F2 GGATCTCAAAGAAGATGGCA 991 This worksxtI-R GGTTCGCCGCGGACATTAAA [17]

a F (forward) and R (reverse).b (A) and (B) – primers used together in a mixture with equimolar concentration.

Pseudophormidium and Calothrix were found in the same area(north of Portugal) as reported in the Checklist. Cyanobium, Lep-tolyngbya and Pseudanabaena are the most numerous among ourisolates (Table S1 and Table 3). Some genera described by [2] werenot found in our study (Chamaecalyx, Dermocarpella, Entophysalis,Hydrococcus, Trichocoleus, Porphyrosiphon, Sirocoleum, and Spiro-coleus), whereas Xenococcus, Microcoleus, Spirulina morphotypeswere present in our field samples but its isolation was not suc-cessful. On the other hand, genera that are not described by theseauthors were present in our samples (Aphanothece, Cyanobium,Synechocystis, Chroococcidiopsis, Chroococcopsis, Leptolyngbya, Pseu-danabaena, Romeria, Schizothrix, Nostoc, and Scytonema), someof them also confirmed by a recent studies on the Portuguesecoast: Synechocystis, Cyanobium, and Leptolyngbya [9,26]. However,Araújo et al. [2] based their work not only on new records but alsoon literature references and herbarium data, and did not isolate theobserved specimens. Representatives of the order Stigonemataleswere not found in this study nor reported by Araújo et al. [2]. Thiswas expected since it is known that these organisms are poorly rep-resented in marine environments [15], and mainly encountered inthe flowing waters of hot springs and soils [11,36,52].

Fig. 1. Distribution of the isolates per taxonomic group.

The diversity of our cyanobacterial isolates is depicted inFigs. 2 and 3 (as well as in Figs. S1 and S2), where it is possibleto observe genera belonging to the four orders. The ChroococcalesSynechocystis and Cyanobium dividing in one plane (Fig. 2a andb, Fig. 3a and b), the Pleurocapsales Chroococcopsis, Myxosarcinaand Hyella dividing in more than one plane (Fig. 2c–g, Fig. 3d) andproducing baeocytes (Fig. 2f), the Oscillatoriales Romeria, Phormid-ium, Leptolyngbya, Pseudophormidium, Lyngbya and Plectonema(Fig. 2h–q, Fig. 3e–f), with Leptolyngbya cf. halophila exhibitingnodule-like structures (Fig. 2m, Fig. 3f) and Plectonema cf. radio-sum geminate false branches (Fig. 2q), and the Nostocales generaNostoc, Calothrix and Scytonema (Fig. 2r–u, Fig. 3g), with intercalar(Fig. 2t) or terminal (Fig. 2s) heterocysts. In Calothrix sp. LEGE 06100hormogonia were found (Fig. 2u). Additionally, the ultrastructuralimages provided information on the size and shape of the sheath(Fig. 3e, f and i), and the different arrangement of the thylakoids(Fig. 3a–c, e, g and h). It was also possible to observe the disrup-tion of the mother sheath leading to disintegration of the coloniesafter cell division (Fig. 3c), and a nodule detail of Leptolyngbya cf.halophila LEGE 06102 (Fig. 3f). A brief description of each of the iso-lates and the morphological characteristics used to taxonomicallyaffiliate them is depicted in Table S1, as well as their distributionand habitat description.

DNA sequence and phylogenetic analysis

Partial 16S rRNA gene sequences were obtained for all the iso-lates. These sequences were compared with the ones available inthe NCBI database (March 2011) using BLASTn, and the results areshown in Table 3. There was a quite good correlation between thephenotypic and genotypic based identifications, for more than one-third of the isolates (Table 3, highlighted in grey). For the remainingisolates discrepancies were observed between the two identifica-tions, possibly resulting from limitations of the databases and/ortaxonomical constraints – it is well recognized that cyanobacterialtaxonomy faces several problems, and is currently under revision[46]. One should point out that 9 of our isolates have less than97% similarity to the 16S rRNA gene sequences in the database,


Table 3Morphological identification (for details see Table S1) and molecular analysis of the cyanobacterial isolates.


Fig. 2. Light micrographs illustrating the diversity of cyanobacterial morphotypes isolated from the intertidal zones of nine beaches on the Portuguese coast. (a) Synechocystissalina LEGE 06099, (b) Cyanobium sp. LEGE 06184, (c) Chroococcopsis sp. LEGE 07168, (d) Chroococcopsis sp. LEGE 07187, (e) Myxosarcina sp. LEGE 06146, (f) Chroococcopsissp. LEGE 07161, (g) Hyella sp. LEGE 07179, (h) Romeria sp. LEGE 06013, (i) Phormidium sp. 1 LEGE 06111, (j) Phormidium laetevirens LEGE 06103, (k) Leptolyngbya mycoideaLEGE 06126, (l) Leptolyngbya mycoidea LEGE 06118, (m) Leptolyngbya cf. halophila LEGE 06102, (n) Leptolyngbya sp. LEGE 06188, (o) Pseudophormidium sp. LEGE 06125, (p)Lyngbya cf. aestuarii LEGE 07165, (q) Plectonema cf. radiosum LEGE 06105, (r) Nostoc sp. 1 LEGE 06106, (s) Calothrix sp. LEGE 06122, (t) Scytonema sp. LEGE 07189, (u) Calothrixsp. LEGE 06100. b – baeocytes; fb – false branching; hg – hormogonia; h – heterocysts; n – nodule. Scale bars – 10 �m.


Fig. 3. Transmission electron micrographs of selected isolates from the sampling sites. (a) Synechocystis salina LEGE 06099, (b) Cyanobium sp. LEGE 06097, (c) Gloeocapsopsiscf. crepidinum LEGE 06123, (d) Chroococcopsis sp. LEGE 07187, (e) Leptolyngbya mycoidea LEGE 06118, (f) Leptolyngbya cf. halophila LEGE 06102, insert – nodule detail (g)Nostoc sp. 2 LEGE 06158, (h) Calothrix sp. 2 LEGE 06122, (i) Rivularia sp. LEGE 07159. het – heterocyst; sh – sheath; sp – septum; th – thylakoid; veg – vegetative cell. arrowhead – disruption of the mother sheath. Scale bars – 1 �m.

emphasizing the presence of novel cyanobacterial diversity in Por-tuguese shore waters (Table 3, dashed boxes).

Phylogenetic analyses were performed to assess the relativepositioning of the cyanobacteria isolated in this study. An MLalgorithm was applied to a multiple alignment of partial 16SrRNA gene sequences (655 bp) of all isolates, reference strains,and C. aurantiacus J-10-fl as the outgroup. The resulting treerevealed 12 consistent clusters (A–L), which were defined as mono-phyletic groups with bootstrap values equal or higher than 70%(Fig. 4). The heterocystous types form a coherent genetic cluster,whereas unicellular and filamentous nonheterocystous forms were

intermixed and dispersed throughout the tree, as previouslydescribed [see for e.g. 10,51]. In general, the phylogenetic distri-bution is congruent with the classification results based on themorphology. Moreover, the species or genus attributed to theisolates in the morphological analysis is in agreement with thespecies/genus of the reference strains they cluster with. ClusterA includes two isolates identified as Leptolyngbya mycoidea sup-ported by a bootstrap value of 100%. Cluster B comprises oneisolate identified as Chroococcopsis sp. and the reference strainSynechocystis sp. PCC 6308. Cluster C includes the Chroococcid-iopsis sp. PCC 6712 and an isolate identified as Hyella sp. Cluster


Fig. 4. Maximum-likelihood phylogenetic tree of partial 16S rRNA gene sequences from the isolates, reference cyanobacterial strains and the outgroup Chloroflexus aurantiacusJ-10-fl. Numbers along branches indicate the percentage of bootstrap support considering 1000 pseudo-replicates: only those equal or higher than 50% are indicated. Isolatesare referred by their culture collection code and morphological identification, whereas reference strains are indicated in bold. The 12 recognized clusters (A–L) are indicatedalong the tree (see text for details).

D includes 3 isolates belonging to Pleurocapsales, and the refer-ence strains Pleurocapsa sp. PCC 7516, Pleurocapsa sp. PCC 7319,and Myxosarcina sp. PCC 7325, corroborating the close relation-ship between these two genera. This is in agreement with previousfindings, where Myxosarcina sp. PCC 7325 clusters with othermembers of Pleurocapsa-group: Pleurocapsa sp. PCC 7516 and PCC

7321 [37]. Chroococcidiopsis PCC 7203, Chroococcidiopsis PCC 6712and Stanieria cyanosphaera PCC 7437, appear distant from all theother baeocyte-forming cyanobacteria, which has been previouslyreported for Chroococcidiopsis PCC 7203 [37]. Cluster E comprisesisolates identified as Leptolyngbya and Phormidium and, althoughno reference strain is included, it is supported by a bootstrap value


of 100%. Cluster F contains three Synechocystis: Synechocystis sp.PCC 6803, and two isolates identified as Synechocystis salina. Thereference strain Synechocystis PCC 6308, is unrelated to the aboveisolates. The scattered distribution of Synechocystis strains has beendescribed previously; Wilmotte and Herdman [51] showed thatSynechocystis PCC 6308 did not group with Synechocystis PCC 6803and Synechocystis PCC 6909. Cluster G includes the genus referencestrain Lyngbya sp. PCC 7419, and one isolate identified as Lyngbyacf. aestuarii (Lyngbya PCC 7419 was originally identified as Lyngbyaaestuarii) [5]. Cluster H contains the heterocystous cyanobacte-ria, and within it is possible to distinguish two sub-clusters: theNostoc and Calothrix/Rivularia. Scytonema hofmanni PCC 7101 isnot included in this cluster due to a low bootstrap value (48.5)but forms a monophyletic group with the heterocystous types,which is in agreement with previous reports [10,32,51]. This clus-ter also contains an isolate identified as Plectonema cf. radiosum.As it described earlier, members of this “botanical” genus mightbe Scytonema individuals that do not exhibit heterocysts even innatural samples [35]. Cluster I, constituted mainly of Oscillato-riales morphotypes, contains isolates identified as Leptolyngbya,Pseudophormidium, Pseudanabaena and the reference strain Lep-tolyngbya PCC 7104. Cluster J contains the Leptolyngbya sp. PCC7375 and the isolate identified as Pseudanabaena cf. frigida. Cluster Kcomprises isolates identified as Pseudanabaena and the marine uni-cellular Synechococcus PCC 7335. This has been previously observedby Wilmotte and Herdman [51] who reported a close relation-ship between filamentous cyanobacteria and Synechococcus PCC7335. Cluster L includes cyanobacteria belonging to Chroococcalesand Oscillatoriales. Within this cluster it is possible differentiate aCyanobium and a Synechococcus sub-cluster. Previously, Herdmanet al. [13] reported that Synechococcus WH 8103 was included in theCyanobium spp. clade, corroborating the data presented here. Thesame 16S rRNA gene alignment was analyzed by Neighbor-Joining(NJ) (Fig. S3), and the computed tree supported all the 12 clusters,therefore validating the ML approach and reinforcing the describedphylogenetic analysis.

It is important to notice that some isolates, identified as thesame taxon (S. salina, L. mycoidea, Leptolyngbya fragilis), appear indifferent branches along the tree, indicating that the phylogeneticanalyses of small coccoids and some filamentous cyanobacteriaoffered a higher resolution than the morphological identification.

Screening for putative N2 fixers and toxin producers

A survey to evaluate the presence of putative N2 fixers andtoxin producers was carried out. Isolates were screened for thepresence of the genes nifK, nifH, mcyA, mcyE, ndaF and sxtI (see“Materials and methods” section). With the exception of the fil-amentous heterocystous strains, it is not possible to identifyN2-fixing cyanobacteria based on morphology; therefore targetingthe structural genes encoding the nitrogenase enzymatic complexis a common approach [53,54]. In this work, the presence of nifH,nifK and/or heterocysts were taken into account to determine theorganism’s potential for nitrogen fixation. 33% of the isolates (20out of 60, see Table S1) qualify as potential diazotrophs, suggest-ing that cyanobacteria may play an important role in N2 fixationin these intertidal zones. Intertidal ecosystems are harsh and oftennutrient-limited environments, so the presence of nitrogen fixersis expected and this ability may confer cyanobacteria a competi-tive advantage [6,41]. Indeed, cyanobacteria are the most importantN2-fixing organisms in the majority of the marine microbial mats,and high rates of nitrogen fixation have been observed [42]. Con-cerning the genes encoding proteins involved in toxin production,no amplification was obtained indicating the absence, among theisolates, of cyanobacteria producing the conventional freshwatertoxins: microcystin, nodularin and saxitoxin. One should bear in

mind that a negative PCR result does not exclude the presence of thegene and, conversely, the presence of the gene does not translateinto expression and activity (also valid for the nif genes screen-ing), but the primer pairs used in this study have been proven towork with a wide range of cyanobacterial strains [14,16,17]. Mar-tins et al. [26] demonstrated that extracts of marine Synechocystisand Synechococcus strains isolated from the Portuguese coast weretoxic to marine invertebrates possibly implicating the presence ofother toxic compounds.

Conclusions

This study unveils the cultivated diversity of cyanobacteriapresent in the intertidal zones of the continental Portuguese coast.The isolated cyanobacteria belong to 35 different morphotypes andcomprise members of all the cyanobacterial orders/subsections,with the exception of Stigonematales/subsection V. The predom-inant forms among the isolates were nonheterocystous and wereparticularly represented by the filamentous Oscillatoriales. The16S rRNA gene sequences of 9 strains have less than 97% similar-ity compared to the sequences in GenBank, revealing unreportedcyanobacterial diversity. The phylogenetic distribution, based onthe 16S rRNA gene, is congruent with the results obtained with themorphology-based classification. As expected, the heterocystouscyanobacteria form a coherent genetic cluster, whereas the uni-cellular and filamentous nonheterocystous were intermixed. Mostof the isolates cluster with reference strains of the same speciesor genus assigned to the isolates in the morphological identifica-tion. One-third of the isolated organisms are putative diazotrophs,suggesting that cyanobacteria may play an important role in N2fixation along the continental Portuguese coast, as it is common inthe marine environment. Additionally, no conventional freshwa-ter toxins genes were detected by PCR screening, indicating a lowprobability for the occurrence of producers of these cyanotoxins inthe analyzed zones.

Acknowledgments

This work was financially supported by FCT (POCTI/CTA/46733/2002, PTDC/MAR/102258/2008, SFRH/BD/29106/2006,SFRH/BD/70284/2010), ESF (III Quadro Comunitário de Apoio),COMPETE – Programa Operacional Factores de Competitividadena sua componente FEDER. We are grateful to Rita Ferreira for thehelp with the 16S rRNA gene sequencing.

Appendix A. Supplementary data

Supplementary data associated with this article can be found, inthe online version, at doi:10.1016/j.syapm.2011.07.003.

References

[1] Abed, R.M.M., Kohls, K., de Beer, D. (2007) Effect of salinity changes on the bacte-rial diversity, photosynthesis and oxygen consumption of cyanobacterial matsfrom an intertidal flat of the Arabian Gulf. Environ. Microbiol. 9, 1384–1392.

[2] Araújo, R., Bárbara, I., Tibaldo, M., Berecibar, E., Tapia, P.D., Pereira, R., Santos,R., Sousa-Pinto, I. (2009) Checklist of benthic marine algae and cyanobacteriaof northern Portugal. Bot. Mar. 52, 24–46.

[3] Bauer, K., Díez, B., Lugomela, C., Seppala, S., Borg, A.J., Bergman, B. (2008)Variability in benthic diazotrophy and cyanobacterial diversity in a tropicalintertidal lagoon. FEMS Microbiol. Ecol. 63, 205–221.

[4] Boone, D.R., Castenholz, R.W., Garrity, G.M. (2001) , Bergey’s Manual of System-atic Bacteriology – The Archaea and the Deeply Branching and PhototrophicBacteria, 1, 2nd ed., Springer, New York, USA, pp. 473–599.

[5] Castenholz, R.W. (2001) Phylum BX. Cyanobacteria. Oxygenic photosyntheticbacteria, in: Boone, D.R., Castenholz, R.W., Garrity, G.M. (Eds.), Bergey’s Manualof Systematic Bacteriology, 1, 2nd ed., Springer, New York, USA, pp. 473–599.

[6] Díez, B., Bauer, K., Bergman, B. (2007) Epilithic cyanobacterial communities ofa marine tropical beach rock (Heron Island, Great Barrier Reef): diversity anddiazotrophy. Appl. Environ. Microbiol. 73, 3656–3668.

http://dx.doi.org/10.1016/j.syapm.2011.07.003


[7] Drummond, A.J., Ashton, B., Buxton, S., Cheung, M., Cooper, A., Duran, C., Field,M., Heled, J., Kearse, M., Markowitz, S., Moir, R., Stones-Havas, S., Sturrock,S., Thierer, T., Wilson, A. (2011) Geneious v5.4, Available from http://www.geneious.com/.

[8] Felsenstein, J. (1981) Evolutionary trees from DNA sequences: a maximumlikelihood approach. J. Mol. Evol. 17, 368–376.

[9] Frazão, B., Martins, R., Vasconcelos, V. (2010) Are known cyanotoxins involvedin the toxicity of Picoplanktonic and filamentous North Atlantic marinecyanobacteria? Mar. Drugs 8, 1908–1919.

[10] Giovannoni, S.F., Turner, S., Olsen, G., Barns, S., Lane, D.J., Pace, N.R. (1988) Evolu-tionary relationships among cyanobacteria and green chloroplasts. J. Bacteriol.170, 3584–3592.

[11] Gugger, M.F., Hoffmann, L. (2004) Polyphyly of true branching cyanobacteria(Stigonematales). Int. J. Syst. Evol. Microbiol. 54, 349–357.

[12] Guindon, S., Gascuel, O. (2003) A simple, fast, and accurate algorithm to esti-mate large phylogenies by maximum likelihood. Syst. Biol. 52, 696–704.

[13] Herdman, M., Castenholz, R.W., Iteman, I., Waterbury, J.B., Rippka, R. (2001)Subsection I (formerly Chroococcales Wettstein 1924, emend. Rippka, Deru-elles, Waterbury, Herdman and Stanier 1979), in: Boone, D.R., Castenholz, R.W.(Eds.), Bergey’s Manual of Systematic Bacteriology, 1, 2nd ed., Springer, NewYork, pp. 493–514.

[14] Hisbergues, M., Christiansen, G., Rouhiainen, L., Sivonen, K., Börner, T. (2003)PCR-based identification of mycrocystin-producing genotypes of differentcyanobacterial genera. Arch. Microbiol. 180, 402–410.

[15] Hoffman, L., Castenholz, R.W. (2001) Subsection V (formerly StigonematalesGeitler 1925), in: Boone, D.R., Castenholz, R.W. (Eds.), Bergey’s Manual of Sys-tematic Bacteriology, 1, 2nd ed., Springer, New York, pp. 589–599.

[16] Jungblut, A.-D., Neilan, B.A. (2006) Molecular identification and evolution ofthe cyclic peptide hepatotoxins, microcystin and nodularin, synthetase genesin three orders of cyanobacteria. Arch. Microbiol. 185, 107–114.

[17] Kellmann, R., Michali, T.K., Neilan, B.A. (2008) Identification of a saxitoxinbiosynthesis gene with a history of frequent horizontal gene transfer. J. Mol.Evol. 67, 526–538.

[18] Knoll, A.H. (2008) Cyanobacteria and earth history. In: Herrero, A., Flores, E.(Eds.), Cyanobacteria: Molecular Biology, Genomics and Evolution, Caister Aca-demic Press, Norfolk, UK, pp. 1–19.

[19] Komárek, J., Anagnostidis, K. (1989) Modern approach to the classification sys-tem of Cyanophytes. 4. Nostocales. Arch. Hydrobiol. Suppl. 82, 247–345.

[20] Komárek, J., Anagnostidis, K. (1998) Cyanoprokaryota. 1. Teil: Chroococcales,in: Ettl, H., Gartner, G., Heynig, H., Mollenhauer, D. (Eds.), Süsswasserflora vonMitteleuropa, vol. 19/1, Gustav Fischer, Stuttgart, p. 548.

[21] Komárek, J., Anagnostidis, K. (2005) Cyanoprokaryota 2 Teil/2nd part: Oscillato-riales, in: Büdel, B., Krienitz, L., Gärtner, G., Schagerl, M. (Eds.), Süsswasserfloravon Mitteleuropa, vol. 19/2, Elsevier/Spektrum, Heidelberg, p. 759.

[22] Kotai, J. 1972 Instructions for Preparation of Modified Nutrient Solution Z8 forAlgae, Publication B-11769, Norwegian Institute for Water Research, Blindern,Oslo.

[23] Kottek, M., Grieser, J., Beck, C., Rudolf, B., Rubel, F. (2006) World map of theKöppen-Geiger climate classification updated. Meteorol. Z. 15, 259–263.

[24] Larkin, M.A., Blackshields, G., Brown, N.P., Chenna, R., McGettigan, P.A.,McWilliam, H., Valentin, F., Wallace, I.M., Wilm, A., Lopez, R., Thompson, J.D.,Gibson, T.J., Higgins, D.G. (2007) Clustal W and Clustal X version 2.0. Bioinfor-matics 23, 2947–2948.

[25] Lima, F.P., Queiroz, N., Ribeiro, P.A., Hawkins, S.J., Santos, A.M. (2006) Recentchanges in the distribution of a marine gastropod, Patella rustica Linnaeus, 1758,and their relationship to unusual climatic events. J. Biogeogr. 33, 812–822.

[26] Martins, R., Fernandez, N., Beiras, R., Vasconcelos, V. (2007) Toxicity assessmentof crude and partially purified extracts of marine Synechocystis and Synechococ-cus cyanobacterial strains in marine invertebrates. Toxicon 50, 791–799.

[27] Mühling, M., Woolven-Allen, J., Murrell, J.C., Joint, I. (2008) Improved group-specific PCR primers for denaturing gradient gel electrophoresis analysis of thegenetic diversity of complex microbial communities. ISME J. 2, 379–392.

[28] Nübel, U., Garcia-Pichel, F., Muyzer, G. (1997) PCR primers to amplify 16S rRNAgenes from cyanobacteria. Appl. Environ. Microbiol. 63, 3327–3332.

[29] Omoregie, E.O., Crumbliss, L.L., Bebout, B.M., Zehr, J.P. (2004) Comparison ofdiazotroph community structure in Lyngbya sp. and Microcoleus chthonoplastesdominated microbial mats from Guerrero Negro, Baja, Mexico. FEMS Microbiol.Ecol. 47, 305–318.

[30] Pontes, M.T., Aguiar, R., Pires, H.O. (2005) A nearshore wave energy atlas forPortugal. J. Offshore Mech. Arct. Eng. 127, 249–255.

[31] Potts, M. (1999) Mechanisms of desiccation tolerance in cyanobacteria. Eur. J.Phycol. 34, 319–328.

[32] Rajaniemi, P., Hrouzek, P., Kastovska, K., Willame, R., Rantala, A., Hoffmann,L., Komárek, J., Sivonen, K. (2005) Phylogenetic and morphological evaluationof the genera Anabaena, Aphanizomenon, Trichormus and Nostoc (Nostocales,Cyanobacteria). Int. J. Syst. Evol. Microbiol. 55, 11–26.

[33] Ramos, V., Seabra, R., Brito, Â., Santos, A., Santos, C.L., Lopo, M., Moradas-Ferreira, P., Vasconcelos, V.M., Tamagnini, P. (2010) Characterization of anintertidal cyanobacterium that constitutes a separate clade together with ther-mophilic strains. Eur. J. Phycol. 45, 394–403.

[34] Rippka, R. (1988) Isolation and purification of cyanobacteria, in: Packer, L.,Glazer, A.N. (Eds.), Methods in Enzymology, vol. 167, Academic Press, San Diego,pp. 3–27.

[35] Rippka, R., Castenholz, R.W., Herdman, M. (2001) Subsection IV (formerlyNostocales Castenholz 1989b sensu Rippka, Deruelles, Waterbury, Herdmanand Stanier 1979), in: Boone, D.R., Castenholz, R.W. (Eds.), Bergey’s Man-ual of Systematic Bacteriology, vol. 1, 2nd ed., Springer, New York, pp.562–589.

[36] Rippka, R., Deruelles, J., Waterbury, J.B., Herdman, M., Stanier, R.Y. (1979)Generic assignments, strain histories and properties of pure cultures ofcyanobacteria. J. Gen. Microbiol. 111, 1–61.

[37] Rippka, R., Waterbury, J.B., Herdman, M., Castenholz, R.W. (2001) Subsection II(formerly Pleurocapsales Geitler 1925, emend. Waterbury and Stanier 1978),in: Boone, D.R., Castenholz, R.W. (Eds.), Bergey’s Manual of Systematic Bacteri-ology, vol. 1, 2nd ed., Springer, New York, pp. 514–539.

[38] Sambrook, J., Russell, D.W. 2001 Molecular Cloning: A Laboratory Manual, 3rded., Cold Spring Harbor Laboratory Press, New York, USA.

[39] Seabra, R., Santos, A., Pereira, S., Moradas-Ferreira, P., Tamagnini, P. (2009)Immunolocalization of the uptake hydrogenase in the marine cyanobacteriumLyngbya majuscula CCAP 1446/4 and two Nostoc strains. FEMS Microbiol. Lett.292, 57–62.

[40] Seabra, R., Wethey, D.S., Santos, A.M., Lima, F.P. (2011) Side matters: micro-habitat influence on intertidal heat stress over a large geographical scale. J.Exp. Mar. Biol. Ecol. 400, 200–208.

[41] Severin, I., Acinas, S.G., Stal, L.J. (2010) Diversity of nitrogen-fixing bacteria incyanobacterial mats. FEMS Microbiol. Ecol. 73, 514–525.

[42] Stal, L.J. (2000) Cyanobacterial mats and stromatolites. In: Whitton, B.A., Potts,M. (Eds.), The Ecology of Cyanobacteria: Their Diversity in Time and Space,Kluwer Academic, Dordrecht, The Netherlands, pp. 61–120.

[43] Stal, L.J., Krumbein, W.E. (1985) Isolation and characterization of cyanobateriafrom a marine microbial mat. Bot. Mar. 28, 351–365.

[44] Stanier, R.Y., Kunisawa, R., Mandel, M., Cohen-Bazire, G. (1971) Purification andproperties of unicellular blue-green algae (order Chroococcales). Bacteriol. Rev.35, 171–205.

[45] Tamagnini, P., Troshina, O., Oxelfelt, F., Salema, R., Lindblad, P. (1997) Hydro-genases in Nostoc sp. strain PCC 73102, a strain lacking a bidirectional enzyme.Appl. Environ. Microbiol. 63, 1801–1807.

[46] Taton, A., Grubisic, S., Ertz, D., Hodgson, D.A., Piccardi, R., Biondi, N., Tredici,M.R., Mainini, M., Losi, D., Marinelli, F., Wilmotte, A. (2006) Polyphasic study ofAntarctic cyanobacterial strains. J. Phycol. 42, 1257–1270.

[47] Waterbury, J.B., Stanier, R.Y. (1978) Patterns of growth and development inpleurocapsalean cyanobacteria. Microbiol. Mol. Biol. Rev. 42, 2–44.

[48] Weisburg, W.G., Barns, S.M., Pelletier, D.A., Lane, D.J. (1991) 16Sribossomal DNA amplification for phylogenetic study. J. Bacteriol. 173,697–703.

[49] Whitton, B.A., Potts, M. (2000) Introduction to the cyanobacteria. In: Whitton,B.A., Potts, M. (Eds.), The Ecology of Cyanobacteria: Their Diversity in Time andSpace, Kluwer Academic, Dordrecht, The Netherlands, pp. 1–11.

[50] Wiegand, C., Pflugmacher, S. (2005) Ecotoxicological effects of selectedcyanobacterial secondary metabolites a short review. Toxicol. Appl. Pharmacol.203, 201–218.

[51] Wilmotte, A., Herdman, M. (2001) Phylogenetic relationships among thecyanobacteria based on 16S rRNA sequences, in: Boone, D.R., Castenholz, R.W.(Eds.), Bergey’s Manual of Systematic Bacteriology, 1, 2nd ed., Springer, NewYork, pp. 487–493.

[52] Wilmotte, A., van der Auwera, A.G., de Watcher, R. (1993) Structure of the16S ribosomal RNA of the thermophilic cyanobacterium Chlorogloeopsis HTF(‘Mastigocladus laminosus HTF’) strain PCC 7518, and phylogenetic analysis.FEBS Lett. 317, 96–100.

[53] Zehr, J.P. (2011) Nitrogen fixation by marine cyanobacteria. Trends Microbiol.19, 162–173.

[54] Zehr, J.P., Mellon, M.T., Zani, S. (1998) New nitrogen fixing microorganismsdetected in oligotrophic oceans by the amplification of nitrogenase (nifH) genes.Appl. Environ. Microbiol. 64, 3444–3450.

http://www.geneious.com/

http://www.geneious.com/

Supplementary Material I

1

Table S1

Morphological identificationa of the cyanobacteria isolated from intertidal zones of the Portuguese coast

ORDER (Subsection)

Genus and species

Culture collection

- LEGE code

(sampling sitee)

Morphology Habitat/Ecology

Cell size

length width f

Description

CHROOCOCCALES (I)

Aphanothece cf. salina

(Synechococcus) b 06149(1) d 1.5-2.1 1.0 -1.4 µm

oval to cylindrical cells

mucilaginous colonies

puddle, submerged stone,

epilithic

Cyanobium sp. Fig. 2b & 3b

06109(4), 06012(5),

06127(7), 06130(9),

06137(3), 06140(4),

06143(7), 07153(6),

06184(7), 06097(6)

0.6-1.0 μm oval cells to rod-shaped

picocyanobacteria,

Sabellaria alveolata reef, wave-

exposed or sheltered pools;

epipsamic, epilithic, plankton

Gloeocapsopsis cf. crepidinum c

Fig. 3c 06123(8) d

3.5-5.0 3.0-4.5 μm

Ø colony 10-14 μm

sub-spherical or rounded-polyhedral cells

colonial green macroalgae, epiphytic

Synechococcus nidulans 07171(7) d 2.1-2.7 0.6-1.0 µm rod-shaped cells, straight to slightly curved

presence of involuted cells

air-exposed rock surface,

epilithic

Synechococcus sp. 07172(9) 0.6-0.8 µm

rod-shaped cells,

presence of involuted cells and

pseudofilaments (2-5 cells)


epilithic

Synechocystis salina Fig. 2a & 3a

06099(1), 06155(2),

07173(9) 1.7- 1.9 μm spherical cells, in pairs or small groups

tide pool, stone or rock surfaces,

epilithic

PLEUROCAPSALES (II)

Chroococcidiopsis sp. 06174(4) 4.7-6.5 μm

baeocytes 2 3 μm spherical to sub-spherical cells plankton

Chroococcopsis sp.

(Myxosarcina and Pleurocapsa-

group) b Fig. 2c, 2d, 2f & 3d

07161(6), 07187(1) 4.0-7.0 5.0-11.0 μm

sheaths up to 2 μm

spherical to oval cells

reddish-brown

Patella sp. and Gibbula sp.

shells, brown macroalgae,

euendolithic/epizoic and

epiphytic

Hyella sp. (Pleurocapsa-group) b

Fig. 2g 07179(1) 2.2-3.8 3.7-5.0 μm

'true borer' cyanobacterium

highly differentiated colonies,

pseudofilaments

Patella sp. shell,

euendolithic/epizoic

Myxosarcina sp.

(Chroococcidiopsis) b Fig. 2e

06146(1) 3.5 μm binary fission in three planes cubical

aggregates, < 8 baeocytes per mother cell

wave-exposed pool, rock surface,

epilithic

OSCILLATORIALES (III)

Leptolyngbya cf. halophila 06102(2), 06110(1), 0.9-1.3 1.1-1.6 μm wavy or coiled trichomes, with nodule-like puddle, submerged stone,

2

Fig. 2m & 3f 06152(3) structures epilithicm, plankton

Leptolyngbya fragilis 07167(3) d, 07176(4) 2.0-4.0 1.2-2.1 μm cylindrical shortly narrowed or rounded apical

cells wave-exposed rock, epilithic

Leptolyngbya minuta 07181(1) d 1.6-2.6 0.7-0.9 µm

very thin trichomes, forming spiral bundles

and mats

reddish


epilithic

Leptolyngbya mycoidea Fig. 2k, 2l & 3e

06009(5), 06108(8),

06118(8), 06126(7),

07157(3)

2.0-3.0 1.4-1.8 μm long trichomes, motile, forming curved to

spiral bundles

tide puddle, rock surface,

epilithic, plankton

Leptolyngbya saxicola 07132(8) d, 07170(9) d 2.5-3.8 0.8-1.0 µm very thin trichomes, forming bundles tide puddle, rock surface,

epilithic

Leptolyngbya sp. 1 06121(9) 1.3-1.7 1.1-1.3 μm straight to slightly curved, cylindrical or

conical rounded apical cell

wave-exposed rock surface,

epilithic

Leptolyngbya sp. 2 Fig. 2n

06188(3) d 1.5-2.3 1.5-1.9 µm long and more or less straight trichomes,

forming radial aggregates plankton

Lyngbya cf. aestuarii Fig. 2p

07165(2) 2.5-4.5 12.0-16.0 μm

sheath up to 4.5 μm

long, straight trichomes, with necridic cells

and hormogonia tide pool, rock surface, epilithic

Phormidium laetevirens

(Oscillatoria) b Fig. 2j

06103(8) 2.2 3.0 μm trichome end shortly hooked, narrowed or

rounded conical apical cell


epilithic

Phormidium sp. 1

(Leptolyngbya) b Fig. 2i

06111(6) 3.0-6.0 2.2-2.8 μm

long trichomes straight or curved, irregularly

clustered or in bundles, forming "leathery"

mats

emersed rock, epilithic

Phormidium sp. 2 (Oscillatoria) b 07162(1) 3.0-4.0 2.5 μm

shortly narrowed trichome ends and often

bent, with thickened outer cell walls

tide pool, submerged stone,

epilithic

Plectonema cf. radiosum

(Scytonema) b Fig. 2q

06105(8) d 3.0-5.0 7.0-9.0 μm sheath

up to 5 μm

slightly narrowing trichomes, with geminate

false branches, apical cells widely rounded,

often capitate

green macroalgae, epiphytic

Pseudanabaena aff. curta 07160(9), 07169(4) 0.7-1.6 0.8-1.4 µm short trichomes, forming Nostoc-like

gelatinous aggregates

wave-exposed rock and puddle,

submerged stone, epilithic

Pseudanabaena aff. persicina 07163(1) d 3.5-4.1 1.2-1.6 μm

long trichomes, straight to curved, cylindrical

cells

reddish

Mytilus sp. shell, epizoic

Pseudanabaena cf. frigida 06144(7) d 1.5-2.1 0.7-0.9 μm straight to curved trichomes

brown culture

wave-sheltered zone, sand,

epipsamic

Pseudanabaena sp. 2 06129(4) 1.0-1.4 1.2-1.8 μm irregularly wavy trichomes plankton

Pseudanabaena sp. 3 06116(6), 07190(3) d 0.8-1.4 0.7-1.5 µm short trichomes (up to 15 µm),

with or without sheaths tide pool, rock surface, epilithic

3

Pseudophormidium sp. c

Fig. 2o 06125(2)

1.0-1.6 2.0-2.4 μm

sheath up to 0,5 μm

binary fission, in many planes (may not be

perpendicular to the major axis)


epilithic

Romeria sp. (Pseudanabaena) b

Fig. 2h 06013(5) 2.8-1.8 1.2-1.6 µm

small trichomes, 2-7 cells (up to 20)

heterogeneous cells wave-exposed rock, epilithic

Schizothrix aff. septentrionalis c 07164(1) 1.3-1.7 1.1-1.5 µm

filaments composed by helically coiled (rope-

like) trichomes (2-3 per filament)

reddish-brown

brown macroalgae, epiphytic

NOSTOCALES (IV)

Calothrix sp. 1 (Rivularia) b

Fig. 2u 06100(3) d

1.5-2.3 6.5-9.5 μm

heterocysts

3.5-4.0 6.0-7.5 μm

heteropolar narrowing trichomes (more

evident in young), without hair formation

one basal hemispherical heterocyst

wave-exposed rock, epilithic

Calothrix sp. 2 (Rivularia) b

Fig. 2s & 3h 06122(2) d, 07177(6) d

basal vegetative cells

2.0-3.0 7.0-9.0 μm

Ø heterocysts 6.5-7.5 μm

heteropolar narrowing trichomes, with hairs

submerged stone and green

macroalgae, epilithic and

epiphytic

Nostoc sp. 1 Fig. 2r

06106(2) d, 06150(2) d

6.0-8 .0 4.0-5.5 μm

Ø akinetes up to 10 μm

Ø colony12.0-26.0 μm

densely, irregularly coiled filaments enclosed

in a common mucilage (up to 2.5 μm wide),

ovoid mature vegetative cells

tide pool, rock surface, epilithic

Nostoc sp. 2 Fig. 3g

06158(9) d

1.8-2.4 2.3-3.1 μm

Ø intercalar heterocysts 2.4-

3.0 μm

colony with narrow wavy filaments, enclosed

in a common mucilage, spherical intercalar

heterocysts, young trichomes with conical

terminal heterocysts

wave-exposed puddle, epilithic

Rivularia sp. Fig. 3i

07159(7) d 2.0-3.0 8.0-12.0 µm

Ø heterocysts 6.5-8.5 µm

long heteropolar filaments, low degree of

narrowing, false branching


epilithic

Scytonema sp. Fig. 2t

07189(1) d

4.5-6.2 4.6-5.5 µm

intercalar heterocysts

6.5-7.5 7-8.5 µm

sheath up to 2.5 µm

false-branching filaments (initially as a

hairpin), constricted at cross walls, trichomes

frequently forming two rows, enclosed in a

common sheath, almost exclusively intercalar

and terminal heterocysts


epilithic

aCyanobacterial taxa were classified according to Komárek & Anagnostidis [19,20,21], Waterbury & Stanier [47] and/or according to Bergey’s Manual of

Systematic Bacteriology [4]. bWhen the identification is different in Bergey’s classification the corresponding form-genus is shown within parenthesis.

c No correspondence in Bergey’s classification.

dPotential nitrogen fixer (presence of a nif gene and/or heterocysts).

e1- Moledo do Minho, 2- S. Bartolomeu do Mar, 3- Lavadores, 4- Aguda, 5- Foz do Arelho, 6- Martinhal, 7- Burgau, 8- Luz, 9- Olhos d’ Água; for details see

Table 1. fRange of values within one standard deviation of the mean (n=20).

4

Fig. S1. Light micrographs of the cyanobacterial isolates. CHROOCOCCALES (Subsection I): (a) Aphanothece cf.

salina LEGE 06149, (b) Cyanobium sp. LEGE 06137, (c) Cyanobium sp. LEGE 06109, (d) Cyanobium sp. LEGE

06140, (e) Cyanobium sp. LEGE 06012, (f) Cyanobium sp. LEGE 06097, (g) Cyanobium sp. LEGE 07153, (h)

Cyanobium sp. LEGE 06127, (i) Cyanobium sp. LEGE 06143, (j) Cyanobium sp. LEGE 06130, (k)

Gloeocapsopsis cf. crepidinum LEGE 06123, (l) Synechococcus nidulans LEGE 07171, (m) Synechococcus sp.

LEGE 07172, (n) Synechocystis salina LEGE 06155; PLEUROCAPSALES (Subsection II): (o) Chroococcidiopsis

sp. LEGE 06174; OSCILLATORIALES (Subsection III): (p) Leptolyngbya cf. halophila LEGE 06110, (q)

Leptolyngbya cf. halophila LEGE 06152, (r) Leptolyngbya fragilis LEGE 07167, (s) Leptolyngbya fragilis LEGE

07176, (t) Leptolyngbya minuta LEGE 07181.

5

Fig. S2. Light micrographs of the cyanobacterial isolates (cont.). OSCILLATORIALES (Subsection III): (a)

Leptolyngbya mycoidea LEGE 07157, (b) Leptolyngbya mycoidea LEGE 06009, (c) Leptolyngbya mycoidea

LEGE 06108, (d) Leptolyngbya saxicola LEGE 07132, (e) Leptolyngbya saxicola LEGE 07170, (f) Leptolyngbya

sp. 1 LEGE 06121, (g) Pseudanabaena aff. curta LEGE 07169, (h) Pseudanabaena aff. curta LEGE 07160, (i)

Pseudanabaena aff. persicina LEGE 07163, (j) Pseudanabaena cf. frigida LEGE 06144, (k) Pseudanabaena

sp.2 LEGE 06129, (l) Pseudanabaena sp. 3 LEGE 07190, (m) Pseudanabaena sp. 3 LEGE 06116, (n)

Schizothrix aff. septentrionalis LEGE 07164; NOSTOCALES (Subsection IV): (o) Calothrix sp. 2 LEGE 07177,

(p) Nostoc sp. 1 LEGE 06150, (q) Nostoc sp. 2 LEGE 06158, (r) Rivularia sp. LEGE 07159.

6

Fig. S3. Neighbor-joining phylogenetic tree of the 16S rRNA gene sequences from the isolates, reference

cyanobacterial strains and the outgroup Chloroflexus aurantiacus J-10-fl. This tree was computed using the

MEGA5 software [2], with the Tamura-Nei substitution model, a gamma distributed parameter of 0.4450 (as

computed by jModelTest [1]) and considering 10000 pseudo-replicates for the bootstrap analysis (indicated only

in those branches with a support equal or higher than 50%). Isolates are referred by their culture collection code,

whereas reference strains are indicated in bold.

[1] Posada, D. (2008). jModelTest: Phylogenetic Model Averaging. Molecular Biology and Evolution. 25(7), 1253-1256. [2] Tamura, K.,

Peterson, D., Peterson, N., Stecher, G., Nei, M., Kumar, S. (2011). MEGA5: Molecular Evolutionary Genetics Analysis using

Maximum Likelihood, Evolutionary Distance, and Maximum Parsimony Methods. Molecular Biology and Evolution.

CHAPTER III

Isolation of cyanobacterial strains from the

Portuguese Atlantic coast disclosures novel

diversity

Brito Â., Mota R., Ramos V., Lima S., Santos A., Vieira J., Vieira C.P.,

Vasconcelos V.M., Tamagnini P. 2015. (Manuscript).

FCUP


Isolation of cyanobacterial strains from the Portuguese Atlantic

coast disclosures novel diversity

Ângela Brito1,2,3, Rita Mota1,2,3, Vitor Ramos3,4, Steeve Lima1,2,3, Arlete Santos1,2,3,

Jorge Vieira1,2, Cristina P. Vieira1,2, Vitor M. Vasconcelos3,4 and Paula

Tamagnini1,2,3*

1i3S - Instituto de Investigação e Inovação em Saúde, Universidade do Porto, Porto,

Portugal

2IBMC - Instituto de Biologia Molecular e Celular, Universidade do Porto, Porto,

Portugal

3Faculdade de Ciências, Departamento de Biologia, Universidade do Porto, Porto,

Portugal

4CIIMAR/CIMAR - Interdisciplinary Centre of Marine and Environmental Research,

University of Porto, Porto, Portugal

Keywords: Geminocystis-like sp.; Hyella; Leptolyngbya; marine cyanobacteria;

Phormidium; Portugal.

*Correspondence: Paula Tamagnini, IBMC - Instituto de Biologia Molecular e Celular,

Universidade do Porto, Rua do Campo Alegre 823, 4150-180 Porto, Portugal, Tel.:

+351226074900; fax: +351226099157, e-mail: [email protected]

Running title: Novel cyanobacterial strains from the Portuguese coast.

46 FCUP


Abstract

Cyanobacteria, in particular tropical marine strains, have been recognized as a prolific

source of secondary metabolites. However, little is known about their counterparts from

temperate regions. Previously, we isolated and characterized sixty cyanobacteria from

the intertidal zones of the Portuguese coast and their potential to produce secondary

metabolites was also evaluated. The 16S rRNA gene sequences of two unicellular and

five filamentous strains showed less than 97% similarity to sequences in the

databases, revealing novel diversity. In the present study, these strains were further

characterized in terms of morphology, ultrastructure and at the molecular level. To

determine their relative position in the cyanobacteria phylum, additional phylogenetic

analyses were carried out using 16S-ITS-23S rRNA gene sequences. The

cyanobacterium belonging to Chroococcales/Subsection I, LEGE 07459 is related to

the genera Geminocystis and Cyanobacterium, but shows distinctive characteristics

indicating that could be the representative of a new genus. The isolate LEGE 07179

was assigned to Hyella sp. (Pleurocapsales/Subsection II) and the molecular data

presented here are the first for members of this genus. Concerning the five isolates

belonging to Oscillatoriales/Subsection III, the new data indicate that LEGE 06188 can

indeed be assigned to a Leptolyngbya morphospecies (a genus known to be

polyphyletic) while the other LEGE 07167, LEGE 07176, LEGE 07157, LEGE 0611

(previously assigned to L. fragilis, L. mycoidea, Phormidium/Leptolyngbya,

respectively) and LEGE 07162 (previously assigned to Phormidium/Oscillatoria)

constitute a distinct group of filamentous cyanobacteria that probably should be

assigned to a new genus.

FCUP


Introduction

Continental Portugal has an extensive coastline with climatic influences from the North

Atlantic ocean and the Mediterranean sea and varied topographies, beach

morphologies and wave regimens [for details on these subjects see (Pontes et al.,

2005; Kottek et al., 2006; Lima et al., 2006; Seabra et al., 2011; Brito et al., 2012)]. Due

to their long evolutionary history and metabolic plasticity, cyanobacteria are

photosynthetic prokaryotes that can be found in a wide range of habitats including the

marine ones (Whitton & Potts, 2000). Marine cyanobacteria play an important role in

the global carbon cycle as primary producers and the ability to fix nitrogen confers

them an advantage in the often nutrient-poor/nitrogen-depleted marine environments

(Knoll, 2008; Zehr, 2011). In the intertidal zones cyanobacteria are exposed to

extremely stressful conditions such as desiccation, temperature and salinity

fluctuations, and often form cohesive mats held together by extracellular polymeric

substances (EPS) that play key structural and protective roles (Stal, 2000; Díez et al.,

2007). In these harsh environments, EPS besides contributing to a structurally-stable

and hydrated microenvironment, are believed to confer protection against UV-radiation

and promote the concentration of nutrients and metal ions (Rossi & De Philippis, 2015).

In the last years, several reports on the diversity of cyanobacteria from Portuguese

fresh and estuarine waters have been published (De Figueiredo et al., 2006; Valério et

al., 2009; Galhano et al., 2011; Lopes et al., 2012), however the diversity of marine

cyanobacteria has been relatively unexplored (Araújo et al., 2009; Ramos et al., 2010;

Brito et al., 2012). Araújo et al., (2009) produced a checklist of benthic marine algae

and cyanobacteria from the north of Portugal that included 26 cyanobacterial species.

The majority of these species were filamentous belonging to the order Oscillatoriales

(Subsection III), but unicellular strains belonging to Chroococcales (Subsection I) and

Pleurocapsales (Subsection II) were also found. Concerning the filamentous

heterocystous morphotypes only two species belonging to Nostocales (Subsection IV)

were reported. Nevertheless, the work performed by these authors was based on new

records, literature references and herbarium data, but did not include the

isolation/cultivation of any of the observed specimens. Subsequently, Brito et al. (2012)

characterized 60 cyanobacterial strains from the intertidal zones along the west and

south coast. The morphological analysis revealed 35 morphotypes (15 genera and 16

species) belonging to the four cyanobacterial Orders/Subsections mentioned above

(again no member of Stigonematales/Subsection V was found). The molecular

characterization based on partial 16S rRNA gene sequences of these isolates showed

that 15% of the sequences obtained had less than 97% similarity compared to the

48 FCUP


sequences available in the databases emphasizing the presence of novel

cyanobacterial diversity. The potential of these strains to fix nitrogen and to produce

secondary metabolites was also evaluated. Our results showed that three out of seven

of the novel strains are putative diazotrophs (presence of nif genes), and that one likely

produces a compound related to naopopeptin or antanapeptin while two other might

produce compounds related to hepatotoxins (Brito et al., 2012; Brito et al., 2015).

Although 16S rRNA gene sequences are the most frequently used to infer

phylogenetic relationships within cyanobacteria, this molecular maker has limited

resolution at the cyanobacteria species level (Konstantinidis & Tiedje, 2007).

Therefore, other less conserved markers can be used to provide complementary

information to the 16S rRNA gene (Komárek, 2014). For instance, the internal

transcribed spacer region (ITS), between the 16S and 23S ribosomal genes, contains

many conserved domains (D1-D5, Box A, Box B) and variable regions (V1-V3) (Iteman

et al., 2000) which have demonstrated to be useful for phylogenetic studies and

identification of new taxa (Rocap et al., 2002; Casamatta et al., 2006). In addition, RNA

secondary structures of this region have been used as discriminatory characters

(Siegesmund et al., 2008; Perkerson III et al., 2011).

The main goal of this study was to further characterize these seven isolates (two

unicellular and five filamentous strains) in terms of morphology, ultrastructure and at

the molecular level. To determine their relative position in the cyanobacteria phylum,

additional phylogenetic analyses were carried out, using 16S-ITS-23S rRNA gene

sequences to obtain a better resolution. This polyphasic study disclosured novel

biodiversity, including new putative genera, and contributed to increase the knowledge

on cyanobacteria from temperate Atlantic areas.

FCUP


Materials and methods

Cyanobacterial strains and culture conditions

The seven cyanobacterial strains studied in this work (Table 1) were previously isolated

from the intertidal zones along the Portuguese Atlantic coast and are deposited at the

LEGE Culture Collection (Laboratório de Ecotoxicologia Genómica e Evolução;

CIIMAR, Porto, Portugal) (Brito et al., 2012). The uni-cyanobacterial Leptolyngbya sp.

LEGE 06188 was grown in Z8 medium (Kotai, 1972) supplemented with 25 g L-1 NaCl

and the remaining uni-cyanobacterial cultures were maintained in MN medium (Rippka,

1988). Both media were supplemented with 10 µg mL-1 B12 vitamin. The cultures were

kept at 25 ºC, under 16 h light (~ 10 mol photons m -2 s-1) / 8 h dark regimen.

Table 1. Cyanobacterial strains, culture collection code and sampling sites.

Order (Subsection) Genus /species LEGE

Code

Sampling sitea

Chroococcales (I) Geminocystis-like sp. 07459 Martinhal

Pleurocapsales (II) Hyella sp. (Pleurocapsa-group) 07179 Moledo do Minho

Leptolyngbya sp. 06188 Lavadores

(Leptolyngbya fragilis)b 07167 Lavadores

Oscillatoriales (III) (Leptolyngbya fragilis)b 07176 Aguda

(Leptolyngbya mycoidea)b 07157 Lavadores

(Phormidium sp./Leptolyngbya)b 06111 Martinhal

a for details see (Brito et al., 2012). b previously assigned to these genera, probably belonging to a new genus.

Light microscopy

Cells were stained with Alcian Blue (0.5% (wt/vol) in 50% ethanol) and observed using

an Olympus X31 light microscope (Olympus, Japan). Light micrographs were acquired

with an Olympus DP25 camera and the CellˆB image software (Olympus, Japan).

Transmission Electron Microscopy (TEM)

For negative staining, 10 µl of culture were placed on a Formvar carbon-coated nickel

grids (200 mesh) for 2 min, the liquid in excess was removed with a filter paper, stained

with 1% uranyl acetate for 5 sec with the excess of liquid removed.

50 FCUP


For TEM, cells were collected, centrifuged and processed as previously described

(Seabra et al., 2009) with the exception of the resin used that was EMBed-812 resin

(Electron Microscopy Sciences, Hatfield, PA, USA). For the isolate LEGE 07157, the

cells were fixed with 2.5% glutaraldehyde and 2% paraformaldehyde in 50 mM sodium

cacodylate buffer (pH 7.2) for 72h and postfixed overnight with 2% osmium tetroxide in

50 mM sodium cacodylate buffer (pH 7.2). Subsequently, the cells were washed three

times in sodium cacodylate buffer and stained en bloc overnight with 2% uranyl

acetate.

The samples or ultrathin sections were examined using a JEM-1400Plus (Jeol, MA,

USA) electron microscope operating at 80 kV.

Scanning Electron Microscopy (SEM)

For SEM analysis, cells were collected by centrifugation and fixed in 2% glutaraldehyde

in 50 mM sodium cacodylate buffer (pH 7.2) and washed one time in double-strength

buffer. The dehydration was performed in a graded series of ethanol (25–100%; v/v)

and dried by critical-point drying. Samples were coated with a gold/palladium thin film,

for 100 sec and a 15 mA current by sputtering, using the SPI Module Sputter Coater

equipment (Structure Probe, Inc., PA, USA). The SEM analysis were performed using

a High Resolution Scanning Electron Microscope (JSM 6301F, Jeol Ltd., Tokyo,

Japan), operating at 15 kV.

DNA extraction, PCR amplification, cloning and sequencing

Genomic DNA was extracted using the phenol/chloroform method previously described

(Tamagnini et al., 1997) and the regions encoding part of 16S rRNA, the internal

transcribed spacer (ITS) and most of the 23S rRNA gene were amplified using the

oligonucleotide primers listed on Table 2. PCR reactions were performed using a

thermal cycler (MyCyclerTM, Bio-Rad laboratories Inc., Hercules, CA, USA) following

procedures previously described (Tamagnini et al., 1997). The PCR profiles included

an initial denaturation at 94 ºC for 5 min, followed by 35 or 30 cycles (for ITS) at 94 ºC

for 1 min, 52 ºC or 60 ºC (for ITS) for 1 min, 72 ºC for 1 min and a final extension at 72

ºC for 7 min.

The amplification of nifH was performed following the methodology previously

described (Omoregie et al., 2004). The PCR products were separated by agarose gel

electrophoresis (Sambrook & Russell, 2001) and DNA fragments were isolated from

gels using the NZYGelpure Kit (NZYTech, Lisbon, Portugal), according to the

manufacturer’s instructions. Purified products were cloned into pGEM®-T Easy vector

FCUP


(Promega, Madison, WI, USA) following with the methodology earlier described

(Ramos et al., 2010).

Table 2. Oligonucleotide primers used in this study.

Target

gene/sequence Primer Sequence 5´ 3´ Reference

16S rRNA 16SF CGCTAGTAATCGCAGGTCA This work

ITSR CGGCTACCTTGTTACGACTT This work

rRNA ITS CSIF GYCACGCCCGAAGTCRTTAC Janse et al., 2004

ULR CCTCTGTGTGCCTAGGTATC Janse et al., 2004

23S rRNA ITSF GGAGAACGCCAATACCCA This work

ULF GATACCTAGGCACACAGAGG Ramos et al., 2010

23S800F GGTTAGGGGTGAAATGCCAA Ramos et al., 2010

23S867F GTTGGCGGATGAGGTGTGGTTA This work

23S893F GCGGTACCAAATCGAGACAAAC This work

23S1682F CTCTCTAAGGAACTCGGCAAA Ramos et al., 2010

23S819R TTGGCATTTCACCCCTAACC This work

23S1027R GCTGGTGGTCTGGGCTGTTT Ramos et al., 2010

23S1932R CCTTAGGACCGTTATAGTTA Ramos et al., 2010

p23SrV_r1 TCAGCCTGTTATCCCTAGAG Sherwood & Presting, 2007

nifH nifH4 TTYTAYGGNAARGGNGG Omoregie et al., 2004

nifH3 ATRTTRTTNGCNGCRTA Omoregie et al., 2004

nifH2 ADNGCCATCATYTCNCC Omoregie et al., 2004

nifH1 TGYGAYCCNAARGCNGA Omoregie et al., 2004

Nucleotide sequence accession numbers

Sequence data were deposited in the GenBank database under the accession

numbers KR676346 - KR676352 and KC256766, KC256770, KC256774 (nifH).

Phylogenetic analysis

The 16S-ITS-23S rRNA gene sequences obtained in this study were used as query in

a megaBLAST, at NCBI (database limited to cyanobacteria, April 2015) and sequences

up to 50% sequence coverage were retrieved. A multiple alignment encompassing

16S-ITS-23S rRNA gene sequences was performed using the ClustalW2 algorithm

(Larkin et al., 2007), with all the default parameters. The evolutionary history was

inferred using the Minimum Evolution method (Rzhetsky & Nei, 1992). The optimal tree

52 FCUP


with the sum of branch length = 2.69316579 is shown. The percentage of replicate

trees in which the associated taxa clustered together in the bootstrap test (500

replicates) are shown next to the branches (Felsenstein, 1985). The tree is drawn to

scale, with branch lengths in the same units as those of the evolutionary distances

used to infer the phylogenetic tree. The evolutionary distances were computed using

the Maximum Composite Likelihood method (Tamura et al., 2004) and are in the units

of the number of base substitutions per site. The ME tree was searched using the

Close-Neighbor-Interchange (CNI) algorithm (Nei & Kumar, 2000) at a search level of

1. The Neighbor-joining algorithm (Saitou & Nei, 1987) was used to generate the initial

tree. The analysis involved 74 nucleotide sequences. All ambiguous positions were

removed for each sequence pair. There were a total of 5526 positions in the final

dataset. Evolutionary analyses were conducted in MEGA6 (Tamura et al., 2013). From

this analysis 4 groups were defined. However, due to the limited information available

in databases (majority of the sequences obtained were from cyanobacterial genomes),

a phylogenetic tree using only the 16S rRNA gene sequences was also performed. For

this purpose, the 16S rRNA gene sequences were searched against the NCBI

(database limited to cyanobacteria, April 2015) and the closest sequences (ordered by

max. identity) that appeared until the delimiting sequence (defined in the 16S-ITS-23S

tree) were retrieved as well as all the sequences bellow with the same identity

percentage. The ME tree (16S rRNA gene) was constructed for each group using the

procedure described above.

Secondary structure of ITS

ITS sequence was folded using CLC Genomics Workbench software 8 (CLC Bio-

Qiagen, Aarhus, Denmark) and the different regions of the ITS were identified.

Pigment analysis

Phycobiliproteins were extracted by disruption of the cells in 1 ml of buffer (50 mM Tris-

HCL pH 8.0, 5 mM EDTA and 50 mM NaCl) containing 0.3 g of 0.2 mm-diameter acid

washed glass beads (Sigma-Aldrich, St. Louis, USA) using a FastPrep®-24 instrument

(MP Biomedicals, CA, USA). Afterwards, samples were centrifuged at 4 °C and 16 000

g for 5 min. The absorbance spectra were recorded using an UV-2401PC

spectrophotometer (Shimadzu Corporation, Japan) in the range 350 - 750 nm.

FCUP


Nitrogenase activity

For the measurement of the in vivo nitrogenase activity by the acetylene reduction

assay (ARA), the cultures were grown in ASNIII0 medium (ASNIII without nitrate)

(Rippka et al., 1979) under 12 h light (25 µmol photons m-2 s-1) / 12 h dark cycles at 30

ºC for 5 days. 10 mL of each culture where transferred to Erlenmeyer flasks with a total

volume of 130 ml and incubated with 13% acetylene (C2H2) for 12 h in sealed flasks.

Afterwards, 1 ml gas samples were withdrawn to determine the amount of acetylene

reduced to ethylene. The measurements were performed using the gas chromatograph

(GC) Clarus 480® (Perkin Elmer, MA, USA), equiped with a Thermal Conductivity

Detector (TDC) and a capillary Porapak N 80/100 column, under the control of the

TotalChrom® software (Perkin Elmer, MA, USA). Nitrogen was used as a carrier gas

and the following temperatures were set for the GC (injector – 120 ºC, detector – 120

ºC and column – 60 ºC). Calibration was performed using a range of different ethylene

gas concentrations. The nitrogenase activity was expressed per chlorophyll a and time.

Chlorophyll a content was determined as previously described (Meeks & Castenholz,

1971). For positive and negative controls, cells of Anabaena sp. PCC 7120 grown in

BG110 (Stanier et al., 1971) or BG110 supplemented with ammonium chloride,

respectively.

Results and discussion

In a previous study we isolated and characterized 60 cyanobacterial strains from the

intertidal zones of the Portuguese coast and it was possible to identify 35 morphotypes

with a clear preponderance of filamentous non-heterocystous strains (Brito et al.,

2012). Among these isolates there was a high degree of novel diversity since nine 16S

rRNA gene sequences exhibit less than 97% similarity compared to sequences

available in the databases. These sequences were from two unicellular strains (one

belonging to Chroococcales/Subsection I and the other to Pleurocapsales/Subsection

II) and seven filamentous strains (Oscillatoriales/Subsection III). However, from these

nine strains only seven could be maintained for an extended period of time (more than

one year) in the LEGE Culture Collection. One of cultures, LEGE 07161, contained in

fact two cyanobacterial strains (Chroococcopsis sp. and a Geminocystis-like sp.), being

the later initially mistaken for nanocytes (see Fig. 1a). The partial 16S rRNA gene

sequence obtained at the time was from the Geminocystis-like isolate to which a new

LEGE code was now attributed - LEGE 07459 (sequence details also corrected at

GenBank: KR676352). Unfortunately, the Chroococcopsis sp. morphologically

54 FCUP


characterized by Brito et al. 2012 did not survived, so from these two co-cultured

strains only the Geminocystis-like sp. LEGE 07459 was characterized in this study. The

comprehensive characterization presented here contemplates the following strains:

Geminocystis-like sp. LEGE 07459 (Chroococcales/Subsection I), Hyella sp. LEGE

07179 (Pleurocapsales/Subsection II), Leptolyngbya sp. LEGE 06188, L. fragilis LEGE

07167, L. fragilis LEGE 07176, L. mycoidea LEGE 07157 and Phormidium sp. LEGE

06111 (Oscillatoriales/Subsection III).

Isolate belonging to Chroococcales (Subsection I)

Chroococcales comprise the unicellular cyanobacteria that reproduce by equal binary

fission or by budding. Cells are spherical, ellipsoidal or rod-shaped (0.5-30 µm), and

occurring single or in aggregates (Castenholz, 2001).

The recently described genus Geminocystis is characterized by solitary, spherical or

oval cells (3-10 µm in diameter) that divide by binary fission into two planes. The

organization of thylakoids is irregular over the whole cell protoplast (Korelusová et al.,

2009).

Geminocystis-like sp. LEGE 07459 (aff. Geminocystis sp.) (Fig. 1), isolated from the

Martinhal beach (N 37º01´07.30´´/ W 8º55´36.17´´) south of Portugal, displays

spherical cells with 2.0 to 3.0 µm in diameter that form aggregates (Fig. 1e) surrounded

by slime (Fig. 1c and e). The cell size is smaller than most of Geminocystis members

and similar to members of the genus Synechocystis. However, the arrangement of the

thylakoids is irregular over almost the whole cell (Fig. 1d) as in Geminocystis and

Cyanobacterium, and contrast with the genus Synechocystis in which the arrangement

is generally parietal (Korelusová et al., 2009). Using negative staining, long and

abundant pilus-like could be observed (Fig. 1f), resembling Synechocystis sp. PCC

6803 type IV pili (Bhaya et al., 2000).

FCUP


Fig.1 - (a) Micrograph of the initial culture containing tw o cyanobacterial strains, one morphologically identif ied as

Chroococcopsis sp. LEGE 07161 (larger cells) and the other as Geminocystis-like sp. LEGE 07459 (cells indicated by

arrow s); (b) Light micrograph of the uni-cyanobacterial culture of Geminocystis-like sp. LEGE 07459 and (c) Culture

stained w ith Alcian blue, slime (sl); (d) TEM ultrathin section of Geminocystis-like sp. LEGE 07459 show ing the

thylakoids (th) arrangement and a forming septum (sp; arrow s); (e) SEM image w here it is possible to observe slime

surrounding cell aggregates; (f) Negative staining show ing the presence of pili (pl). Scale bars: (a) = 10 µm; (b, c) = 20

µm; (d) = 0.2 µm; (e) =10 µm; (e – insert) = 3µm; (f) = 0.5 µm.

In the phylogenetic analysis based on 16S-ITS-23S rRNA gene sequences, LEGE

07459 is in a cluster comprising members of the genera Geminocystis and

Cyanobacterium (closer to Geminocystis) with a strong bootstrap support, and differs

distinctly from members of Synechocystis that are in a different position in phylogenetic

tree (Fig. 9, highlighted in grey). In the 16S rRNA gene sequence tree, the isolate

LEGE 07459 is closely related to several marine uncultured cyanobacteria strains from

very distinct locations (without bootstrap support): marine reef, Oahu, Hawaii (Zheng et

al., 2011), Cíes Island, Galicia, Spain (Acosta‐González et al., 2013) and South Atlantic

Bight (SAB) off the coast of Savannah (Hunter et al., 2006), and the members of the

genera Geminocystis and Cyanobacterium are in a distinct cluster (Fig. S1). It was

previously described that Geminocystis differs from Synechocystis by position in the

phylogenetic tree, arrangement of thylakoids and ITS secondary structure, sharing

these characteristics with Cyanobacterium. Geminocystis differs from Cyanobacterium

in cell shape and number of planes of fission (Korelusová et al., 2009). As stated

previously, the LEGE 07459 spherical cell shape is similar to Geminocystis but

apparently divides in one plane as Cyanobacterium. The 367 nucleotides long 16S-23S

rRNA ITS secondary structure of our isolate was also analysed (Table 3) taking into

account the conserved and variable regions previously described by Iteman (Iteman et

al., 2000). The secondary structure of the region D1-D1´ of LEGE 07459 is short -18

56 FCUP


nucleotides - and similar to those reported for Geminocystis and Cyanobacterium

(Korelusová et al., 2009), the boxB is similar to Geminocystis, Cyanobacterium and

Synechocystis, and the V3 region could not be determined (as for Geminocystis

papuanica). The ITS region of LEGE 07459 contains also the tRNAAla and tRNAIle as

Geminocystis and Cyanobacterium contrasting with Synechocystis that only contains

tRNAIle (Korelusová et al., 2009).

In summary, LEGE 07459 is more related to cyanobacteria of the genera

Geminocystis and Cyanobacterium than to genus Synechocystis, but possess some

distinctive characteristics that might indicate that this isolate could be the

representative of a new genus.

Table 3. Secondary structure of 16S-23S rRNA internal transcribed spacer (ITS) of

Geminocystis-like sp. LEGE 07459.

D1-D1´ Box B V3 tRNA

Not

determined

Ala Ile

Isolate belonging to Pleurocapsales (Subsection II)

This Order/Subsection comprises unicellular cyanobacteria that can reproduce by

multiple fission producing small and dispersible cells - baeocytes.

Some members of the cyanobacterial genus Hyella (Bornet & Flahault, 1888) have

been described as endoliths penetrating calcareous substrates (e.g. shells) (Al‐Thukair

& Golubic, 1991; Castenholz, 2001). Pseudofilaments (uniseriate to multiseriate) grow

into the stony substrate enveloped by thin or thickened, firm or mucilaginous,

sometimes layered sheaths (Guiry & Guiry, 2015). Cells display different sizes (2.5-5.5

µm in diameter) and are more or less spherical, polygonal-rounded and oval or club-

shaped at the ends of pseudofilaments and oldest cells (basal) may divide into

baeocytes (reproductive cells) (Komárek, 2003). Most of the described species are

FCUP


from marine coastal environments (Lukas & Golubic, 1983; Al‐Thukair & Golubic,

1991).

The isolate Hyella sp. LEGE 07179 was collected from Moledo do Minho beach (N

41º50´58.68´´/ W 8º52´0.18´´) north of Portugal, from shells of the marine gastropod

mollusc Patella sp.. The isolate was previously described as a “true borer”

cyanobacterium with highly differentiated colonies and pseudofilaments (Brito et al.,

2012). Cells are more or less spherical with 2.5-5 µm in diameter and form colonies

held together by extracellular polymeric substances (EPS) (Fig. 2a) from which the

pseudofilaments emerge (Fig. 2c to e). The cells are surrounded by a stratified multi-

layered sheath that can be easily observed in the TEM micrographs (Fig. 2b and c). In

the TEM images is also evident the parietal arrangement of the thylakoids and the very

particular shape of the pseudofilament cells (Fig. 2c).

Fig.2 - Micrographs of Hyella sp. LEGE 07179. (a) Light microscopy showing the presence of slime (sl) surrounding the

cells stained w ith Alcian blue; (b) and (c) TEM ultrathin sections of a cell and a pseudofilament, respectively. In these

images the arrangement of the thylakoids (th) and the thick stratif ied sheath (sh) are highlighted; (d) SEM image w here

it is possible to observe the colonial morphology and the emerging pseudofilament (pf); (e) Detail of the pseudofilament

(pf). Scale bars: (a) = 50 µm; (b, c) = 1 µm; (d) = 60 µm; (e) = 10 µm.

In the phylogenetic analysis based on 16S-ITS-23S rRNA gene sequences, Hyella

sp. LEGE 07179 clearly clusters with Stanieria cyanosphaera PCC 7437 a member of

Pleurocapsales (Subsection II), but it is a very divergent sequence (Fig. 9, highlighted

in green). In the 16S rRNA phylogenetic tree this isolate is within a weakly supported

cluster that includes an uncultured cyanobacteria strain (microbial mat from

58 FCUP


stromatolite, Lagoa Salgada, Brazil) and Chroococcidiopsis sp. PCC 6712

(Pleurocapsales/Subsection II), although forming a distinct branch (Fig. S2).

In summary, this preliminary analysis clearly showed that this isolate cluster with

members of Pleurocapsales but without close relatives. This isolate was assigned as

Hyella, a genus belonging to the Pleurocapsales/Subsection II. This is a group of

cyanobacteria for which there is a lack of molecular data. At the moment, our 16S

rRNA gene sequence is the only one available on the databases for the genus Hyella

(Brito et al., 2012; Komárek et al., 2014) and therefore it was not possible to infer the

phylogenetic relationship between our isolate and other members of Hyella.

Isolates belonging to Oscillatoriales (Subsection III)

Oscillatoriales/Subsection III comprises filamentous cyanobacteria that undergo binary

fission in a single plane and with filaments composed exclusively by vegetative cells.

The genus Leptolyngbya (Anagnostidis & Komárek, 1988) includes filamentous

cyanobacteria that exhibit very thin trichomes (<3 µm), consisting of cylindrical or

isodiametric cells that are enclosed in a well-defined sheath. Apical cells are rounded

(or exceptionally conical) and the trichome breakage may be trans or inter-cellular. In

some members of this genus trichomes can be surrounding by a “mucus -like” matrix

(Castenholz, 2001).

Four of our strains, isolated from two beaches on the north of Portugal, Lavadores

and Aguda (N 41º07´45.07´´/ W 8º39´19.22´´ and N 41º02´58.35´´/ W 8º39´19.22´´,

respectively), were assigned to the genus Leptolyngbya (Brito et al., 2012).

Leptolyngbya sp. LEGE 06188 has thin trichomes (width less than 3 µm)

surrounded by a well-defined sheath and slime (Fig. 3a) and the cells are barrel

shaped with sharp constrictions at the cells junctions (Fig. 3b to e). TEM micrographs

showed the stratified sheath, the parietal arrangement of the thylakoids, and the

presence of carboxysomes and polyphosphate bodies in the cytoplasm (Fig. 3b and c).

In SEM micrographs the barrel shape of cells and the breakage of trichome can also be

observed (Fig. 3d and e).

FCUP


Fig.3 - Micrographs of Leptolyngbya sp. LEGE 06188. (a) Light micrograph of the f ilaments stained w ith Alcian blue

highlighting the sheath (sh); (b) and (c) TEM ultrathin sections show ing the parietal arrangement of the thylakoids (th)

and the stratif ied sheath (sh); (d) and (e) SEM images w here it is possible to observe the barrel shaped cells and the

breakage of a trichome w ith the tw o parts still connected by the sheath (sh). Scale bars: (a) = 50 µm; (b, c) = 0.5 µm; (d)

= 6 µm; (e) = 10 µm.

In the phylogenetic analysis based on 16S-ITS-23S rRNA gene sequences,

Leptolyngbya sp. LEGE 06188 clusters with a unicellular cyanobacterium isolated from

the south Portuguese coast - Gloeocapsopsis crepidinum (Ramos et al., 2010; Brito et

al., 2012) (Fig. 9, highlighted in pink). The phylogenetic tree using only the 16S rRNA

gene sequences revealed a different topology (Fig. S3). The isolate LEGE 06188

clusters with several marine uncultured cyanobacteria strains from north-eastern

Mexico (Sahl et al., 2011). This sub-cluster is within a clade composed by

Leptolyngbya members (Fig. S3).

In summary, the morphological and ultrastructural data together with the preliminary

phylogenetic studies supported the assignment of this isolate to the genus

Leptolyngbya, a genus known to be polyphyletic, in contrast with the other four

filamentous isolates investigated in this study (see below).

The isolates LEGE 07167 and LEGE 07176 were previously assigned to

Leptolyngbya fragilis (Brito et al., 2012). Both display thin trichomes (width less than 3

µm) composed by cylindrical cells that are much more elongated (Fig. 4 and Fig. 5)

than the ones from the strain described above. In both isolates, the terminal cells are

more or less conical (see for e.g. Fig. 4c) and large amount of slime can be observed

surrounding the filaments stained with Alcian blue (Fig. 4a and 5a). The thylakoids

arrangement is clearly parietal in the isolate LEGE 07167 whereas in LEGE 07176 the

thylakoids are more or less parallel over almost the whole cell (Fig. 4b and 5b).

60 FCUP


Fig.4 - Micrographs of LEGE 07167, previously assigned to Leptolyngbya fragilis. (a) Beams of f ilaments surround by

slime (sl) stained w ith Alcian blue; (b) Longitudinal section of a f ilament in w hich is evident the parietal arrangement of

thylakoids (th) w ithin each cell; (c) SEM image w here it is possible to observe the cylindrical shaped cells and the

conical terminal cell. Scale bars: (a) = 50 µm; (b) = 1 µm; (c) = 4 µm.

In negatively stained LEGE 07176 cells, small spherical structures emerging from

the outer membrane could be observed (Fig. 5c and d). It is known that many bacteria

can release vesicles that play a role in different processes such as quorum signalling

(Mashburn & Whiteley, 2005) and horizontal gene transfer (Kolling & Matthews, 1999;

Rumbo et al., 2011). Recently, the formation of extracellular vesicles was reported in

the unicellular marine cyanobacteria Prochlorococcus (Biller et al., 2014) and in the

filamentous freshwater Anabaena sp. PCC 7120 strain (Oliveira et al., 2015). Overall,

these data indicate that a wide range of morphological distinct cyanobacteria have the

ability to produce and release vesicles that, most probably, play an important role in the

interactions between the microorganisms and between microorganisms and the

environment (Biller et al., 2014).

Fig.5 - Micrographs of LEGE 07176, previously assigned to Leptolyngbya fragilis. (a) Aggregated f ilaments stained w ith

Alcian blue highlighting the presence of slime (sl); (b) TEM ultrathin section show ing the arrangement of the thylakoids

(th); (c) and (d) Negative staining show ing the presence of vesicles (vs) emerging from the outer membrane and the

presence of thin pili (pl). Scale bars: (a) = 50 µm; (b) = 0.2 µm; (c) = 0.1 µm; (d) = 0.2 µm.

FCUP


LEGE 07157 was previously assigned to Leptolyngbya mycoidea and also has thin

trichomes with less than 3 µm (Fig. 6a) but, in contrast with the previously described

strains, the cells are more or less isodiametric (Fig. 6b). Once again, a large amount of

polysaccharidic material can be discerned around the filaments stained with Alcian blue

(Fig. 6a) and in between the parallel arranged filaments in the SEM image (Fig. 6d).

The terminal cells are rounded with a calyptra-like structure (Fig. 6c), a thick ended

cap on apical cells of some filamentous cyanobacteria (Palinska & Marquardt, 2008). In

the TEM micrograph the parietal arrangement of the thylakoids can be observed as

well as the cytoplasm and cytoplasmatic inclusions (Fig. 6b).

Fig.6 - Micrographs of LEGE 07157, previously assigned to Leptolyngbya mycoidea. (a) Light microscopy of the thin

f ilaments surrounded by large amounts of slime (sl) stained w ith Alcian blue; (b) Ultrastructure of the more or less

isodiametric cells w here the parietal arrangement of the thylakoids (th) and several cell inclusions can be observed; (c)

and (d) SEM images of the f ilaments surface w here it possible to observe the calyptra on a terminal cell (arrow head)

and slime in betw een the f ilaments (sl). Scale bars: (a) = 50 µm; (b) = 0.5 µm; (c) = 6 µm; (d) = 3 µm.

The isolate LEGE 06111 was collected from the Martinhal beach (N 37º01´07.30´´/

W 8º55´36.17´´) south of Portugal (Brito et al., 2012). As stated by Brito et al. (2012)

this isolate could either be assigned to the genus Phormidium according to Komárek

and Anagnostidis (Komárek & Anagnostidis, 2005) or to the genus Leptolyngbya

following the Bergey’s Manual of Systematic Bacteriology (Castenholz, 2001). LEGE

06111 displays thin trichomes (2.5 µm wide) composed by elongated cells (Fig. 7a and

b), the apical cells are pointed with a calyptra-like structure (Fig. 7d) and some

polysaccharidic material (slime) can be discerned around the filaments stained with

Alcian blue (Fig. 7a). TEM ultrathin sections showed the cylindrical shape of the cells,

the parietal arrangement of thylakoids and the presence of subcellular structures like

carboxysomes in the centre of the cell (Fig. 7b). Although the culture is uni-

62 FCUP


cyanobacterial, it is not axenic and it was possible to observe bacteria tightly bound to

the thin sheath that surrounds the filament (Fig. 7c).

Fig.7 - Micrographs of LEGE 06111, previously assigned to Phormidium/Leptolyngbya. (a) Light micrograph of the

f ilaments stained w ith Alcian blue dye show ing the presence of slime (sl); (b) TEM ultrathin section highlighting the cells

shapes and the parietal arrangement of the thylakoids (th); (c) Negative staining show ing the presence of bacteria (bac)

tightly bound to the thin sheath; (d) SEM image exposing the calyptra (arrow head). Scale bars: (a) = 50 µm; (b, c) = 0.5

µm; (d) = 5 µm.

Recently, twenty-nine Phormidium-like strains were analysed concerning their

phycobilin pigment composition (Palinska et al., 2011) and the authors showed that

they group in three well defined types. Most of the strains (23) contained phycocyanin

(PC) but not phycoerythrin (PE), while the other groups contained both PC and PE with

a low or a high PE:PC ratio. The absorption spectrum observed for the strain LEGE

06111 (Fig. 8) fits the second type describe by Palinska et al. (Palinska et al., 2011)

that contains both phycobiliproteins and a low PE:PC ratio.

Fig.8 - Absorption spectrum obtained for LEGE 06111, previously assigned to Phormidium/Leptolyngbya. Chl a -

chlorophyll a; Carot - carotenoids; PE - phycoerythrin; PC - phycocyanin.

In the phylogenetic analysis based on 16S-ITS-23S rRNA gene sequences, the four

isolates mentioned above LEGE 07167, LEGE 07176, LEGE 07157 and LEGE 06111

are in a distinct sub-cluster, supported by strong bootstrap value. LEGE 07176 forms a

distinct branch and appeared as a “loner sequence”. This sub-cluster is within a clade

FCUP


composed by filamentous strains belonging to the genera Microcoleus, Arthospira and

Spirulina (Fig. 9, highlighted in blue). In the tree based on the 16S rRNA gene

sequences, LEGE 07176 is in a different position compared to the other 3 isolates (Fig.

S4), a weakly supported cluster that includes members of Spirulina as well as a

filamentous cyanobacteria isolate and several uncultured marine cyanobacteria

collected from microbial mats, Elkhorn Slough estuary, CA, USA (Woebken et al.,

2012), but constituting a separate branch. The other three isolates, together with

another LEGE 07162 previously assigned to the genus Phormidium and characterized

in Brito et al. 2012, form a well-supported distinct sub-cluster.

In summary, the phylogenetic data presented here indicate that LEGE 06188 can

indeed be assigned to the genus Leptolyngbya, a genus known to be polyphyletic,

while the isolates LEGE 07167, LEGE 07176, LEGE 07157, LEGE 06111 (previously

assigned to L. fragilis, L. mycoidea, Phormidium/Leptolyngbya, respectively) and LEGE

07162 (previously assigned to Phormidium/Oscillatoria) constitute a distinct group of

filamentous cyanobacteria that probably could be assigned to a new genus firstly

isolated from the Portuguese coast.

64 FCUP


Fig.9 - Minimum evolution (ME) tree of partial 16S-ITS-23S rRNA gene sequences from the seven isolates and strains

retrieved from the BLAST analysis. Numbers along branches indicate bootstrap support considering 500 pseudo-

replicates: only those equal or above 0.5 are indicated. Each isolate is indicated in bold and by the respective GenBank

accession number (brackets). The 4 groups defined are highlighted in the tree.

FCUP


Capacity to fix atmospheric nitrogen and to produce secondary metabolites

It was previously demonstrated that three of the strains described above, Hyella sp.

LEGE 07179, Leptolyngbya sp. LEGE 06188 and L. fragilis LEGE 07167 harboured the

genes encoding the nitrogenase complex, notably nifK (nitrogenase beta subunit)

and/or nifH (dinitrogenase reductase) (Brito et al., 2012). In this work partial sequences

of nifH were obtained (GenBank accession numbers: KC256774, KC256766,

KC256770), and the cyanobacteria ability to fix nitrogen was further evaluated by

growing the strains in ASNIII medium with decreasing concentrations of combined

nitrogen until depletion. Moreover, the nitrogenase activity was evaluated by the

acetylene reduction assay (ARA). Although the strains could grow (Leptolyngbya sp.

LEGE 06188) or survive (Hyella sp. LEGE 07179, Leptolyngbya fragillis LEGE 07167)

in medium without combined nitrogen the nmoles of ethylene produced/chl a/h were

residual compared to our control strain Anabaena/Nostoc sp. PCC 7120 (data not

show), indicating they may have the ability to fix nitrogen in the environment but that

this capacity might have been suppressed during the prolonged period of cultivation in

media with combined nitrogen. An in situ assay will help to clarify if these

cyanobacterial strains indeed play a role as N2 fixers in the marine environment.

Additionally, crude extracts of these seven cyanobacterial strains previously analysed

by LC-MS/MS coupled with molecular networking (Brito et al., 2015) revealed that two

of the filamentous strains - LEGE 07167 and LEGE 07176 - may produce compounds

related to the hepatotoxins generally produced by their freshwater counterparts,

microcystin and nodularin. Preliminary LC-MS/MS analysis directed to detect

microcystin showed that LEGE 07167 extract had a MS fragmentation pattern

coinciding with microcystin-LR (data not shown). Further studies are required to

unequivocally demonstrate that these marine cyanobacteria are indeed capable to

produce these cyanotoxins.

Conclusions

This polyphasic study allowed a better characterization of the seven isolates from the

intertidal zones of the Portuguese coast and unveiled their position within the

cyanobacteria phylum. We could demonstrate that although the unicellular isolate

LEGE 07459 (Chroococcales/Subsection I) is related to the genera Geminocystis and

Cyanobacterium, it possess distinctive characteristics indicative of a different genus.

The molecular data reported here for the isolate LEGE 07179, Hyella sp.

(Pleurocapsales/Subsection II), were the first for members of this genus. Concerning

66 FCUP


the five filamentous isolates belonging to Oscillatoriales/Subsection III, the new data

indicate that LEGE 06188 is a Leptolyngbya morphospecies, while the other four -

LEGE 07167, LEGE 07176, LEGE 07157, LEGE 06111 - together with LEGE 07162

constitute a distinct group of filamentous cyanobacteria that probably should be

assigned to a new genus firstly isolated from the Portuguese coast. Moreover, our

results suggest that the strains Hyella sp. LEGE 07179, Leptolyngbya sp. LEGE 06188

and LEGE 07167 might contribute to N2 fixation in their environment. Furthermore, two

of the filamentous strains - LEGE 07167 and LEGE 07176 - may produce cyanotoxins

(Brito et al., 2015). If further studies confirm our preliminary work on this subject, this

constitutes the first report of microcystin-producing filamentous cyanobacterial strain

from the temperate Atlantic region.

Acknowledgments

This work was funded by National Funds through FCT - Fundação para a Ciência e a

Tecnologia, grants SFRH/BD/70284/2010 (to AB), SFRH/BD/84914/2012 (to RM) and

SFRH/BD/80153/2011 (to VR). To this work also contribute the project MARBIOTECH

(reference NORTE-07-0124-FEDER-000047) within the SR&TD Integrated Program

MARVALOR - Building research and innovation capacity for improved management

and valorisation of marine resources, supported by ON.2 Program and by the

European Regional Development Fund (ERDF) through the COMPETE - Operational

Competitiveness Programme, through the project NOVOMAR and national funds

through FCT – Foundation for Science and Technology, through the project “PEst-

C/MAR/LA0015/2013”. We are also grateful to Daniela Silva from Centro de Materiais

da Universidade do Porto (CEMUP), Porto, Portugal, for technical assistance with SEM

and to Paulo Oliveira, IBMC, for the help with the GC measurements.

FCUP


References

Acosta‐González A, Rosselló‐Móra R & Marqués S (2013) Characterization of the

anaerobic microbial community in oil‐polluted subtidal sediments: aromatic

biodegradation potential after the Prestige oil spill. Environ Microbiol 15: 77-92.

Al‐Thukair AA & Golubic S (1991) New endolithic cyanobacteria from the arabian gulf.

I. Hyella immanis SP. NOV. 1. J Phycol 27: 766-780.

Anagnostidis K & Komárek J (1988) Modern approach to the classification system of

cyanophytes. 3. Oscillatoriales. Arch Hydrobiol Algolog Studies 50-53: 327-472.

Araújo R, Bárbara I, Tibaldo M, Berecibar E, Tapia PD, Pereira R, Santos R & Pinto IS

(2009) Checklist of benthic marine algae and cyanobacteria of northern Portugal.

Bot Mar 52: 24-46.

Bhaya D, Bianco NR, Bryant D & Grossman A (2000) Type IV pilus biogenesis and

motility in the cyanobacterium Synechocystis sp. PCC 6803. Mol Microbiol 37: 941-

951.

Biller SJ, Schubotz F, Roggensack SE, Thompson AW, Summons RE & Chisholm SW

(2014) Bacterial vesicles in marine ecosystems. Science 343: 183-186.

Bornet E & Flahault C (1888) Note sur deux nouveaux genres d'algues perforantes. J

Bot Morot 2: 161-165.

Brito Â, Gaifem J, Ramos V, Glukhov E, Dorrestein PC, Gerwick WH, Vasconcelos VM,

Mendes MV & Tamagnini P (2015) Bioprospecting Portuguese Atlantic coast

cyanobacteria for bioactive secondary metabolites reveals untapped chemodiversity.

Algal Res 9: 218-226.

Brito Â, Ramos V, Seabra R, et al. (2012) Culture-dependent characterization of

cyanobacterial diversity in the intertidal zones of the Portuguese coast: a polyphasic

study. Syst Appl Microbiol 35: 110-119.

Casamatta DA, Gomez SR & Johansen JR (2006) Rexia erecta gen. et sp. nov. and

Capsosira lowei sp. nov., two newly described cyanobacterial taxa from the Great

Smoky Mountains National Park (USA). Hydrobiologia 561: 13-26.

Castenholz RW (2001) Phylum BX. Cyanobacteria. Oxygenic Photosynthetic Bacteria.

Bergey’s Manual of Systematic Bacteriology, Vol. 1 (Boone DR, Castenholz RW &

Garrity G, eds.), pp. 473-599. Springer, New York.

De Figueiredo DR, Reboleira AS, Antunes SC, Abrantes N, Azeiteiro U, Goncalves F &

Pereira MJ (2006) The effect of environmental parameters and cyanobacterial

blooms on phytoplankton dynamics of a Portuguese temperate lake. Hydrobiologia

568: 145-157.

68 FCUP


Díez B, Bauer K & Bergman B (2007) Epilithic cyanobacterial communities of a marine

tropical beach rock (Heron Island, Great Barrier Reef): diversity and diazotrophy.

Appl Environ Microb 73: 3656-3668.

Felsenstein J (1985) Confidence limits on phylogenies: an approach using the

bootstrap. Evolution 39: 783-791.

Galhano V, De Figueiredo DR, Alves A, Correia A, Pereira MJ, Gomes-Laranjo J &

Peixoto F (2011) Morphological, biochemical and molecular characterization of

Anabaena, Aphanizomenon and Nostoc strains (Cyanobacteria, Nostocales)

isolated from Portuguese freshwater habitats. Hydrobiologia 663: 187-203.

Guiry M & Guiry G (2015) AlgaeBase. World-wide electronic publication, National

University of Ireland, Galway. http://www.algaebase.org; searched on 08 April 2015.

Hunter EM, Mills HJ & Kostka JE (2006) Microbial community diversity associated with

carbon and nitrogen cycling in permeable shelf sediments. Appl Environ Microb 72:

5689-5701.

Iteman I, Rippka R, de Marsac NT & Herdman M (2000) Comparison of conserved

structural and regulatory domains within divergent 16S rRNA–23S rRNA spacer

sequences of cyanobacteria. Microbiology 146: 1275-1286.

Janse I, Kardinaal WEA, Meima M, Fastner J, Visser PM & Zwart G (2004) Toxic and

nontoxic Microcystis colonies in natural populations can be differentiated on the

basis of rRNA gene internal transcribed spacer diversity. Appl Environ Microb 70:

3979-3987.

Knoll AH (2008) Cyanobacteria and earth history. The Cyanobacteria: Molecular

Biology, Genomics, and Evolution,(Herrero A & Flores E, eds.), pp. 1-20. Caister

Academic Press, Nortfolk, UK.

Kolling GL & Matthews KR (1999) Export of virulence genes and Shiga toxin by

membrane vesicles of Escherichia coli O157: H7. Appl Environ Microb 65: 1843-

1848.

Komárek J (2003) Coccoid and colonial cyanobacteria. Freshwater Algae of North

America: Ecology and Classification, Vol. 3 (Wehr JD & Sheath RG, eds.), pp. 59-

116. Academic Press, New York.

Komárek J (2014) Modern classification of cyanobacteria. Cyanobacteria: An Economic

Perspective,(Sharma NK, Rai aK & Stal LJ, eds.), pp. 21-39. Willey Blackwell, UK.

Komárek J & Anagnostidis K (2005) Cyanoprokaryota-2. Teil/2nd part: Oscillatoriales.

Süsswasserflora von Mitteleuropa 19/2,(Büdel B, Krienitz L, Gärtner G & Schagerl

M, eds.), pp. 759. Elsevier/Spektrum, Heidelberg.

FCUP


Komárek J, Kaštovský J, Mareš J & Johansen JR (2014) Taxonomic classification of

cyanoprokaryotes (cyanobacterial genera) 2014, using a polyphasic approach.

Preslia 86: 295-335.

Konstantinidis KT & Tiedje JM (2007) Prokaryotic taxonomy and phylogeny in the

genomic era: advancements and challenges ahead. Curr Opin Microbiol 10: 504-

509.

Korelusová J, Kaštovský J & Komárek J (2009) Heterogeneity of the cyanobacterial

genus Synechocystis and description of a new genus, Geminocystis. J Phycol 45:

928-937.

Kotai J (1972) Instructions for preparation of modified nutrient solution Z8 for algae.

Publication B-11/69. Norwegian Institute for Water Research, Blindern, Oslo.

Kottek M, Grieser J, Beck C, Rudolf B & Rubel F (2006) World map of the Köppen-

Geiger climate classification updated. Meteorol Z 15: 259-263.

Larkin MA, Blackshields G, Brown NP, et al. (2007) Clustal W and Clustal X version

2.0. Bioinformatics 23: 2947-2948.

Lima FP, Queiroz N, Ribeiro PA, Hawkins SJ & Santos AM (2006) Recent changes in

the distribution of a marine gastropod, Patella rustica Linnaeus, 1758, and their

relationship to unusual climatic events. J Biogeogr 33: 812-822.

Lopes VR, Ramos V, Martins A, Sousa M, Welker M, Antunes A & Vasconcelos VM

(2012) Phylogenetic, chemical and morphological diversity of cyanobacteria from

Portuguese temperate estuaries. Mar Environ Res 73: 7-16.

Lukas KJ & Golubic S (1983) New endolithic cyanophytes from the north atlantic

ocean. II. Hyella gigas lukas & golubic sp. nov. from the Florida continental margin.

J Phycol 19: 129-136.

Mashburn LM & Whiteley M (2005) Membrane vesicles traffic signals and facilitate

group activities in a prokaryote. Nature 437: 422-425.

Meeks JC & Castenholz RW (1971) Growth and photosynthesis in an extreme

thermophile, Synechococcus lividus (Cyanophyta). Arch Mikrobiol 78: 25-41.

Nei M & Kumar S (2000) Molecular evolution and phylogenetics. Oxford University

Press, New York.

Oliveira P, Martins NM, Santos M, Couto NA, Wright PC & Tamagnini P (2015) The

Anabaena sp. PCC 7120 Exoproteome: Taking a peek outside the box. Life 5: 130-

163.

Omoregie EO, Crumbliss LL, Bebout BM & Zehr JP (2004) Comparison of diazotroph

community structure in Lyngbya sp. and Microcoleus chthonoplastes dominated

microbial mats from Guerrero Negro, Baja, Mexico. FEMS Microbiol Ecol 47: 305-

318.

70 FCUP


Palinska KA & Marquardt J (2008) Genotypic and phenotypic analysis of strains

assigned to the widespread cyanobacterial morphospecies Phormidium autumnale

(Oscillatoriales). Arch Microbiol 189: 325-335.

Palinska KA, Deventer B, Hariri K & Lotocka M (2011) A taxonomic study on

Phormidium-group (cyanobacteria) based on morphology, pigments, RAPD

molecular markers and RFLP analysis of the 16S rRNA gene fragment. Fottea 11:

41-55.

Perkerson III RB, Johansen JR, Kovácik L, Brand J, Kaštovský J & Casamatta DA

(2011) A unique pseudanabaenalean (cyanobacteria) genus Nodosilinea gen. nov.

based on morphological and molecular data. J Phycol 47: 1397-1412.

Pontes MT, Aguiar R & Pires HO (2005) A nearshore wave energy atlas for Portugal. J

Offshore Mech Arct 127: 249-255.

Ramos V, Seabra R, Brito Â, Santos A, Santos CL, Lopo M, Moradas-Ferreira P,

Vasconcelos VM & Tamagnini P (2010) Characterization of an intertidal

cyanobacterium that constitutes a separate clade together with thermophilic strains.

Eur J Phycol 45: 394-403.

Rippka R (1988) Isolation and purification of cyanobacteria. Method Enzymol, Vol. 167

(Packer L & Glazer AN, eds.), pp. 3-27. Academic Press, San Diego.

Rippka R, Deruelles J, Waterbury JB, Herdman M & Stanier RY (1979) Generic

assignments, strain histories and properties of pure cultures of cyanobacteria. J Gen

Microbiol 111: 1-61.

Rocap G, Distel DL, Waterbury JB & Chisholm SW (2002) Resolution of

Prochlorococcus and Synechococcus ecotypes by using 16S-23S ribosomal DNA

internal transcribed spacer sequences. Appl Environ Microb 68: 1180-1191.

Rossi F & De Philippis R (2015) Role of cyanobacterial exopolysaccharides in

phototrophic biofilms and in complex microbial mats. Life 5: 1218-1238.

Rumbo C, Fernández-Moreira E, Merino M, Poza M, Mendez JA, Soares NC,

Mosquera A, Chaves F & Bou G (2011) Horizontal transfer of the OXA-24

carbapenemase gene via outer membrane vesicles: a new mechanism of

dissemination of carbapenem resistance genes in Acinetobacter baumannii.

Antimicrob Agents Chemoter 55: 3084-3090.

Rzhetsky A & Nei M (1992) A simple method for estimating and testing minimum-

evolution trees. Mol Biol Evol 9: 945-967.

Sahl JW, Gary MO, Harris JK & Spear JR (2011) A comparative molecular analysis of

water‐filled limestone sinkholes in north‐eastern Mexico. Environ Microbiol 13: 226-

240.

FCUP


Saitou N & Nei M (1987) The neighbor-joining method: a new method for

reconstructing phylogenetic trees. Mol Biol Evol 4: 406-425.

Sambrook J & Russell DW (2001) Molecular cloning: a laboratory manual. Cold Spring

Harbor Laboratory Press New York.

Seabra R, Wethey DS, Santos AM & Lima FP (2011) Side matters: microhabitat

influence on intertidal heat stress over a large geographical scale. J Exp Mar Biol

Ecol 400: 200-208.

Seabra R, Santos A, Pereira S, Moradas‐Ferreira P & Tamagnini P (2009)

Immunolocalization of the uptake hydrogenase in the marine cyanobacterium

Lyngbya majuscula CCAP 1446/4 and two Nostoc strains. FEMS Microbiol Lett 292:

57-62.

Sherwood AR & Presting GG (2007) Universal primers amplify a 23S rDNA plastid

marker in eukaryotic algae and cyanobacteria. J Phycol 43: 605-608.

Siegesmund MA, Johansen JR, Karsten U & Friedl T (2008) Coleofasciculus gen. nov.

(cyanobacteria): morphological and molecular criteria for revision of the genus

Microcoleus Gomont. J Phycol 44: 1572-1585.

Stal LJ (2000) Cyanobacterial mats and stromatolites. The ecology of cyanobacteria:

Their diversity in time and space,(Whitton BA & Potts M, eds.), pp. 61-120. Springer,

The Netherlands.

Stanier R, Kunisawa R, Mandel M & Cohen-Bazire G (1971) Purification and properties

of unicellular blue-green algae (order Chroococcales). Bacteriol Rev 35: 171.

Tamagnini P, Troshina O, Oxelfelt F, Salema R & Lindblad P (1997) Hydrogenases in

Nostoc sp. strain PCC 73102, a strain lacking a bidirectional enzyme. Appl Environ

Microbiol 63: 1801-1807.

Tamura K, Nei M & Kumar S (2004) Prospects for inferring very large phylogenies by

using the neighbor-joining method. P Natl Acad Sci USA 101: 11030-11035.

Tamura K, Stecher G, Peterson D, Filipski A & Kumar S (2013) MEGA6: molecular

evolutionary genetics analysis version 6.0. Mol Biol Evol 30: 2725-2729.

Valério E, Chambel L, Paulino S, Faria N, Pereira P & Tenreiro R (2009) Molecular

identification, typing and traceability of cyanobacteria from freshwater reservoirs.

Microbiology 155: 642-656.

Whitton BA & Potts M (2000) Introduction to the Cyanobacteria. The ecology of

cyanobacteria: their diversity in time and space,(Whitton BA & Potts M, eds.), pp. 1-

11. Springer, The Netherlands.

Woebken D, Burow LC, Prufert-Bebout L, Bebout BM, Hoehler TM, Pett-Ridge J,

Spormann AM, Weber PK & Singer SW (2012) Identification of a novel

72 FCUP


cyanobacterial group as active diazotrophs in a coastal microbial mat using

NanoSIMS analysis. ISME J 6: 1427-1439.

Zehr JP (2011) Nitrogen fixation by marine cyanobacteria. Trends Microbiol 19: 162-

173.

Zheng G, Xin W, Hannides AK & Sansone FJ (2011) Impact of redox-stratification on

the diversity and distribution of bacterial communities in sandy reef sediments in a

microcosm. Chin J Oceanol Limnol 29: 1209-1223.

Supplementary Material II

FCUP


Fig.S1 - Minimum evolution (ME) phylogenetic tree of partial 16S- rRNA gene sequences from Geminocysis-like sp.

LEGE 07459. Numbers along branches indicate bootstrap support considering 500 pseudo-replicates: only those equal

or above 0.50 are indicated. Geminocysis-like sp. LEGE 07459 is indicated in bold.

76 FCUP


Fig.S2 - Minimum evolution phylogenetic tree of partial 16S rRNA gene sequence from the Hyella sp. LEGE 07179.

Numbers along branches indicate bootstrap support considering 500 pseudo-replicates: only those equal or above 0.5

are indicated. Hyella sp. LEGE 07179 is indicated in bold.

FCUP


Fig.S3 - Minimum evolution phylogenetic tree of partial 16S- rRNA gene sequence from the Leptolyngbya sp. LEGE

06188. Numbers along branches indicate bootstrap support considering 500 pseudo-replicates: only those equal or

above 0.5 are indicated. Leptolyngbya sp. LEGE 06188 is indicated in bold.

78 FCUP


Fig.S4 - Minimum evolution phylogenetic tree of partial 16S rRNA gene sequence from the LEGE 07176 (Leptolyngbya

fragilis), LEGE 06111 (Phormidium sp.), LEGE 07167 (Leptolyngbya fragilis) and LEGE 07157 (Leptolyngbya mycoidea)

(highlighted in blue). Numbers along branches indicate bootstrap support considering 500 pseudo-replicates: only those

equal or above 0.5 are indicated in bold.

CHAPTER IV

Bioprospecting Atlantic cyanobacteria for

bioactive secondary metabolites reveals

untapped chemodiversity

Brito Â., Gaifem J., Ramos V., Glukhov E., Dorrestein P.C., Gerwick W.H.,

Vasconcelos V.M., Mendes M.V., Tamagnini P. 2015. Algal Research 9: 218-

226.

Bioprospecting Portuguese Atlantic coast cyanobacteria for bioactivesecondary metabolites reveals untapped chemodiversity

Ângela Brito a,b,c, Joana Gaifem a,b, Vitor Ramos d, Evgenia Glukhov e, Pieter C. Dorrestein f,g,William H. Gerwick e,g, Vitor M. Vasconcelos c,d, Marta V. Mendes a,b, Paula Tamagnini a,b,c,⁎a i3S — Instituto de Investigação e Inovação em Saúde, Universidade do Porto, Porto, Portugalb IBMC— Instituto de Biologia Molecular e Celular, Universidade do Porto, Porto, Portugalc Faculdade de Ciências, Departamento de Biologia, Universidade do Porto, Porto, Portugald CIIMAR/CIMAR — Interdisciplinary Centre of Marine and Environmental Research, University of Porto, Porto, Portugale Center for Marine Biotechnology and Biomedicine, Scripps Institution of Oceanography, University of California San Diego, La Jolla, CA 92093, USAf Department of Chemistry and Biochemistry, University of California San Diego, La Jolla, CA 92093, USAg Skaggs School of Pharmacy and Pharmaceutical Sciences, University of California San Diego, La Jolla, CA 92093, USA

a b s t r a c ta r t i c l e i n f o

Article history:Received 5 January 2015Received in revised form 10 March 2015Accepted 23 March 2015Available online xxxx

Keywords:Atlantic OceanBioactive compoundsMarine cyanobacteriaMolecular networkingNRPSPKS

Continental Portugal has an extensive Atlantic Ocean coastline; however little is known about the diversity of itsmarine cyanobacteria, with only a few reports published. Cyanobacteria are a prolific source of bioactive com-pounds with promising therapeutic applications. Previously, several cyanobacterial strains (Subsections I–IV)were isolated from the Portuguese Atlantic coast and characterized using a polyphasic approach. Preliminaryscreening indicated that these cyanobacteria lacked the genes for proteins involved in the production of conven-tional cyanotoxins. However, it was previously shown that extracts of marine Synechocystis and Synechococcuswere toxic to invertebrates, with crude extracts causing stronger effects than partially purified ones. To furtherevaluate the potential of our temperate region Portuguese isolates to produce bioactive compounds, a PCRscreening for the presence of genes encoding non-ribosomal peptide synthetases (NRPSs) and polyketidesynthases (PKSs), targeting the adenylation (A) and ketosynthase (KS) domains respectively, was performed.DNA fragments were obtained for more than 80% of the strains tested, and the results revealed that PKS genesare more ubiquitous than NRPS genes. The sequences obtained were used in an in silico prediction of the PKSand NRPS systems. RT-PCR analyses revealed that these genes are transcribed under routine laboratoryconditions in several selected strains. Furthermore, LC–MS analysis coupled with molecular networking, amass spectrometric tool that clustersmetaboliteswith similarMS/MS fragmentation patterns, was used to searchfor novel or otherwise interesting metabolites revealing an untapped chemodiversity.

© 2015 Elsevier B.V. All rights reserved.

1. Introduction

Cyanobacteria are ancient prokaryotes with the capacity to performoxygenic photosynthesis, and can be found in a great diversity of habitats,including extreme environments that are challenging to most of the lifeforms in Earth. Given their metabolic plasticity, cyanobacteria are a valu-able andprolific source of biologically active secondarymetabolites [1–6],and some of these products can be toxic to higher animals includinghumans. These cyanotoxins can be classified into five functional groupsaccording to their primary target organ or effects being designated ashepatotoxins, neurotoxins, cytotoxins, dermatotoxins and irritant toxins[7,8]. Understandably, amajority of early cyanobacterial natural productsresearch focused mostly on toxins, particularly on the hepatotoxin

microcystin which is widespread in freshwater ecosystems worldwide.However, cyanobacteria can also produce secondary metabolites withpromising therapeutic properties such as anticancer, antibiotic andanti-inflammatory [4,9,10].

Marine cyanobacteria, in particular, are increasingly being recog-nized as important source of structurally diverse secondarymetabolites[2,4,11,12]. Nearly 800 compounds have been identified in marinecyanobacteria, mainly in filamentous strains. Oscillatoriales (SubsectionIII) contribute to almost half of the reported compounds (49%),Nostocales (Subsection IV) with 26%, and Stigonematales (SubsectionV) with merely 4%. Among the unicellular strains, the Chroococcales(Subsection I) contribute 15% and the Pleurocapsales (Subsection II)about 6% [3,13]. However, the above percentages may be skewed dueto selection and/or coverage bias.

The majority of the bioactive metabolites isolated from cyanobacteriaare polyketides (PKs), non-ribosomal peptides (NRPs) or a hybridof these two, and they are synthesized by modular multienzymatic

Algal Research 9 (2015) 218–226

⁎ Corresponding author at: IBMC — Instituto de Biologia Molecular e Celular,Universidade do Porto, Rua do Campo Alegre 823, 4150-180 Porto, Portugal.

E-mail address: [email protected] (P. Tamagnini).

http://dx.doi.org/10.1016/j.algal.2015.03.0162211-9264/© 2015 Elsevier B.V. All rights reserved.

Contents lists available at ScienceDirect

Algal Research

j ourna l homepage: www.e lsev ie r .com/ locate /a lga l

http://crossmark.crossref.org/dialog/?doi=10.1016/j.algal.2015.03.016&domain=pdf

http://dx.doi.org/10.1016/j.algal.2015.03.016

mailto:[email protected]


http://www.sciencedirect.com/science/journal/

www.elsevier.com/locate/algal

complexes, namely type I polyketide synthases (PKSs) and non-ribosomal peptide synthetases (NRPSs). Type I PKS and NRPSpathways are characterized by a modular organization in which eachmodule is responsible for the incorporation of a structural carboxylicacid or amino acid unit respectively [5,14,15].

In a previous study, several cyanobacterial strainswere isolated fromnine sites in the intertidal zones along the Portuguese Atlantic coast andextensively characterized using a polyphasic approach [16]. Thesestrainswere shown to bemorphologically and phylogenetically very di-verse belonging to orders Chroococcales, Pleurocapsales, Oscillatorialesand Nostocales (Subsections I through IV). In this previous work, thegenes encoding proteins involved in the production of the conventionalcyanotoxins were not detected by PCR screening. However, it wasshown that extracts of Synechocystis and Synechococcus, also isolatedfrom the Portuguese coast, were toxic to invertebrates [17]. Thisprevious study showed that the crude extracts from these twopicocyanobacteria had stronger effects than partially purified ones,suggesting the presence of different compounds and synergisticeffect(s). Other recent studies, also using isolates from the Portuguesecoast, revealed that extracts from these strains can have high activityin ecologically-relevant bioassays and toxicity against human cancercell lines [18–20].

In the current study, a PCR screen for the presence of genes encodingtype I PKS and NRPS pathways, targeting the ketosynthase (KS) andadenylation (A) domains, respectively was performed using thecyanobacterial strains previously characterized [16]. Based on thesequences obtained, an in silico prediction of PKS andNRPS typemetab-oliteswas also undertaken. RT-PCR analyseswere carried out for severalselected strains to understand if the genes were transcribed underroutine laboratory conditions. Furthermore, LC–MS analyses of thecyanobacterial extracts coupled with molecular networking, visualrepresentations of mass-spectral data based on similarities in MS-fragmentations [21], were used to search for specific metabolites ofinterest and their analogues, as well as novel compounds.

2. Materials and methods

2.1. Cyanobacterial isolates and culture conditions

The cyanobacterial strains used in this study were previously isolatedfrom the Portuguese coast and are deposited at the LEGE CultureCollection (Laboratório de Ecotoxicologia Genómica e Evolução;CIIMAR, Porto, Portugal) [16]. Additionally, the isolate LEGE 06123is also deposited at Culture Collection of Algae and Protozoa (CCAP1425/1). The uni-cyanobacterial cultures were maintained in thefollowing media: MN [22], BG110, BG11 [23], Z8 [24] supplementedwith 25 g l−1 NaCl, further named Z8 25‰ and ASNIII [25]. The mediawere supplemented with B12 vitamin. The cultures were kept at 25 °C,under a 16 h light (~10 μmol photons m−2 s−1)/8 h dark regimen.

2.2. DNA extraction, PCR amplification and sequencing

Cells were harvested by centrifugation and DNAwas extracted usingthe Maxwell® 16 System according to the instructions of the manufac-turer or by the phenol/chloroform method previously described [26].PCR amplifications of the regions encoding the ketosynthase (KS) andadenylation (A) domains from the polyketide synthase (PKS) andnon-ribosomal peptide synthetase (NRPS) gene clusters, respectivelywere performed using the degenerate oligonucleotide primer pairsDKF/DKR [27] and MTF2/MTR [28], respectively. PCR reactions wereperformed using a thermal cycler MyCycler™ (Bio-Rad laboratoriesInc., Hercules, CA, USA) following procedures previously described[26]. The PCR profiles included an initial denaturation at 94 °C for4 min, followed by 30 cycles of denaturation at 94 °C for 30 s, annealingfor 30 s at 58 °C (DKF/DKR) and 48 °C (MTF2/MTR), followed by exten-sion at 72 °C for 1 min, and a final extension at 72 °C for 7 min. The

PCR products were separated by agarose gel electrophoresis using stan-dard protocols [29]. DNA fragments were isolated from gels usingthe NZYGelpure Kit (NZYtech, Lisbon, Portugal), according to themanufacturer's instructions. Purified PCR products were cloned intopGEM®-T Easy vector (Promega, Madison, WI, USA), and transformedinto Escherichia coli DH5α competent cells following the manufacturer'sinstructions, and the methodology earlier described [30].

2.3. Sequence analysis and in silico prediction of PKS and NRPS systems

Nucleotide sequences were analysed using the CLUSTAL W algo-rithm for multiple sequence alignments [31]. Sequence comparisonswere carried out using BLAST databases at the National Center forBiotechnology Information (NCBI). The obtained sequences encodingKS and A domains were submitted to the web-based tools NaPDoS[32], and the PKS/NRPS analysis [33] to in silico predict PKS and NRPSsystems, respectively. Default parameter settings were used in theseanalyses.

2.4. Nucleotide sequence accession numbers

Novel sequences associatedwith this study are available in GenBankunder the accession numbers KC842284–KC842372 (PKS), andKC813168–KC813194 (NRPS).

2.5. RNA extraction and RT-PCR

Cellswere collected from48h and 2month old cultures, resuspendedwith 1 ml RNAprotect® Bacterial Reagent (Qiagen, GmbH, Duesseldorf,Germany), collected again by centrifugation, and frozen at −80 °C. Thecells were disrupted in TRIzol® Reagent (Ambion, Carlsbad, CA, USA)containing 0.2 g of 0.2mm-diameter acidwashed glass beads (Sigma-Al-drich) using a FastPrep®-24 instrument (MP Biomedicals, CA, USA), andRNAwas extracted with the Purelink™ RNAMini Kit (Ambion, Carlsbad,CA, USA). For cDNA synthesis 500 ng of total RNA was transcribed withthe iScript™ Reverse Transcription Supermix for RT-qPCR (Bio-Rad,Hercules, CA, USA), following the manufacturer's instructions. The PCRswere performed using the primer pairs listed in Table 1, and followingthe methodology previously described [26]. The PCR profile was: 4 minat 94 °C followed by 40 cycles of 1 min at 94 °C, 1 min at 52 °C and45 s at 72 °C, and a final extension at 72 °C for 7 min. Negative controls(no template cDNA) and positive controls (genomicDNA)were includedin all RT-PCRs.

2.6. Extraction, LC–MS analysis and molecular networking generation

To produce crude extracts, the freeze-dried biomass of each of the 57cyanobacterial strains (~30 mg, Table 2) was repeatedly extracted with2:1 CH2Cl2:MeOH (with manual fragmentation of the cyanobacterialclumps), filtered through a Whatman No. 1 filter paper, and dried byrotary evaporation. The resulting crude extract was dissolved in 2:1CH2Cl2:MeOH, transferred to a vial and dried again. Crude extract wasresuspended at ~30 mg ml−1 in 50% acetonitrile (CH3CN) in waterand passed through a Bond Elut-C18 OH cartridge (Agilent Technolo-gies, USA) that was prewashed with 3 ml of CH3CN. Afterwards, theBond Elut-C18 OH cartridge was washed with 3 ml of CH3CN, thesolvent obtained was dried under N2 and the residue was dissolved at20 mg ml−1 in 50% CH3CN in water. A 0.020 ml aliquot of each samplewas injected and analysed via LC–MS/MS on a Thermo Finnigan Survey-or Autosampler-Plus/LC–MS/MS/PDA-Plus system coupled to a ThermoFinnigan LCQ Advantage Max mass spectrometer with a gradient of30–100% CH3CN in water with 0.1% formic acid.

TheMS/MS spectra of these 57 cyanobacteria alongwith 105MS/MSspectra of reference cyanobacterial compounds (natural productsdiscovered in various cyanobacterial collections [34]) were used to gen-erate amolecular network following previously describedmethodology

219Â. Brito et al. / Algal Research 9 (2015) 218–226

[21] and visualized using Cytoscape (www.cytoscape.org). Algorithmsassumed a cosine threshold set at 0.6 and nodes were colour-codedaccording to their origin.

3. Results and discussion

Continental Portugal has an extensive coastline facing the AtlanticOcean; nevertheless, little is known about the biodiversity of marinecyanobacteria from this area with only two recently published reportson this topic [16,35]. However, there have been a few investigationsaimed at assessing their bioactive potential [18–20,36]. In thiscurrent study, 61 isolates belonging to different genera of the ordersChroococcales, Pleurocapsales, Oscillatoriales and Nostocales (corre-sponding to Subsections I through IV) were tested. These isolatescomprise the majority of those previously characterized strains [16] aswell as six others deposited at the LEGE culture collection [18,20].

3.1. PCR screen for PKS and NRPS encoding genes

To evaluate the potential of these cyanobacterial isolates to producesecondary metabolites, a PCR screen was performed for type I polyke-tide synthase (PKS) and non-ribosomal peptide synthetase (NRPS)genes, targeting the ketosynthase (KS) and adenylation (A) domains.Under the conditions tested, the presence of KS domain sequences

was detected in 85% of the strains, rising to 97% among filamentouscyanobacteria (Oscillatoriales and Nostocales) (Table 2). A-domain se-quences were detected in only 56% of the strains tested, and thispercentage fell to 33% in unicellular strains (Chroococcales) (Table 2).Similar to the KS sequences, A-domain sequences were more frequentin filamentous strains (66%). Overall, these data revealed that PKSand/or NRPS encoding genes are present in the majority of thesecyanobacterial isolates, with PKS genes being more ubiquitous thanNRPS. Our results are in agreement with previous reports that detectedthe presence of PKS and NRPS encoding genes in cyanobacteria anddemonstrated the higher frequency of these genes in filamentousstrains [1,14,39–41]. In this current work and accord with previousfindings [39,42], the only genus for which neither KS- nor A-domainsequences could be detectedwas Synechococcus (Table 2). However, re-cent studies found that PKS geneswere present in several Synechococcusstrains, including Synechococcus sp. PCC 7002 [40,41]. Christiansen et al.(2001) could not detect NRPS encoding genes in strains of the genusSynechocystis, while recent studies [40,41] observed the occurrence ofNRPS gene clusters in Synechocystis PCC 7509 (cluster 3, [43]) but notin another phylogenetically distinct strain Synechocystis PCC 6803(cluster 2.1, [43]). Interestingly, here we demonstrated the presenceof NRPS and PKS encoding genes in two strains of this genus,Synechocystis salina LEGE 06099 and LEGE 07173. One should bear inmind that a previous study reported that extracts of Synechocystis

Table 1Oligonucleotide primers used for RT-PCR targeting the cyanobacterial sequences encoding the KS (KS primers) and A (AD primers) domains of the PKS and NRPS genes, respectively.

Organism and target sequencea Primer pair Sequence 5′ → 3′ PCR product expected size (bp)

LEGE 06100 KS4FKS4R

ACAGTGGTATTGATGCTTACAGTAGACAACAATTCCGCAACCTTCC

325

LEGE 06109 (2) KS14-2FKS14-2R

CGGATCAGCTACTTCTACGATAGGCAGTGCGAATGAG

414

LEGE 06109 (4) KS14-4FKS14-4R

CTTCGTCGCTGGACTATCCTCTTTGTTCCCTTTGGC

406

LEGE 06111 (1) KS16-1FKS16-4FKS16-1.4R

CACCAATATCGGCATCGCAACGCCTACACCAGTATCCCATCATTGTTGCAGTCG

386394

LEGE 06111 (3) KS16-3FKS16-3R

TGCTCCTCTTCCATCGTCTCGGGTTGATTTCAAGGC

382

LEGE 06146 (1) KS59-1FKS59-1R

GACCGACTACGCACTATACCGAATGACCGCATAGACC

414

LEGE 06146 (2) KS59-2FKS59-2R

GCAACTTGTGTGGGAAGCCATAACCGTCGCCATTGG

419

LEGE 06146 (3) KS59-3FKS59-3R

ATTATCACCATCGCTTCCTCGAACGCCTGACACAATCC

302

LEGE 06158 (1) KS72-1FKS72-1R

CTGTCGAACCGTATTTCCTCAATGAGGCTCTGTTGG

413

LEGE 06158 (2) KS72-2FKS72-2R

ATCCTCAGCAGCGACTTCTCAGCCTGATCGAAACTC

385

LEGE 07160 (1) KS74-1FKS74-1R

GGATTGGCACGCACGACTATCTACAGCCGTCTTGATTGAC

451

LEGE 07160 (2) KS74-2FKS74-2R

CAAGCCACTGTCGCCAACTTGACCGCCATCATTGCC

396

LEGE 07160 (3) KS74-3FKS74-3R

TGCTGTTAGAGGTCTGCTGTCTGCTGCCATCATCTGC

382

LEGE 07179 (1) KS93-1FKS93-1R

GCTATTACTACGATTTCCACGATGACGGCGATGATGTC

317

LEGE 07179 (1) KS93-2FKS93-2R

AACCGCATCAGCTATTTCGAATGACGGCAAGGATCG

328

LEGE 06013 (2) KS118-2FKS118-2R

GCCTACAGTGTTCAGACCATATCCGCAGCAGTTAGAC

413

LEGE 06111 (2)LEGE 06194 (1)

KSFKSR

TATCTGGCGTTCAACCTGACCGCATGAATATTGTCG

419

LEGE 06109 AD14FAD14R

CTTACCTGATTCGGAAGATACAAGTTCGCCTGTTACTCC

403

LEGE 06158 (1) AD72-1FAD72-1R

CGTATCTTCTCTGAGCAACTTCGGCAAATAAACCTGTGTGTTAGCG

310

LEGE 07179 AD93FAD93R

GTGTGGCCTACGTCATCTATACCGTCTTGCTGTTTGCGATG

406

a When multiple sequences were obtained for a single cyanobacterial strain they are indicated by numbers in between brackets.

220 Â. Brito et al. / Algal Research 9 (2015) 218–226

http://www.cytoscape.org

strains isolated from the Portuguese coast were toxic towards marineinvertebrates suggesting the presence of bioactive compounds withinthis genus [17].

3.2. In silico prediction of PKS and NRPS systems

The DNA fragments obtained by PCRwere cloned and sequenced. Intotal, we obtained 89 sequences from 47 cyanobacteria strains thatencoded for KS-domains. For 18 strains a single KS-domain was identi-fied, whereas for the remaining cyanobacteria two or more KS se-quences were revealed (Supplementary Table S1). BLASTp analysisshowed identities ranging between 50 and 99% to known KS sequences.Concerning the A-domains, 27 sequences were obtained from 19isolates. A single A-domain was identified in 11 strains, while twoA-domainswere identified for the other 8 cyanobacteria (SupplementaryTable S2). BLASTp analysis showed identities ranging between 43 and87% to known NRPS sequences.

Furthermore, these sequences were used for in silico analyses oftheir PKS and NRPS systems using the following web-based softwarepackages: the NaPDoS [32] for PKS and PKS/NRPS analysis [33] forNRPS analysis. NaPDoS is a bioinformatic tool for the rapid detectionand analysis of secondarymetabolite encoding genes. This tool employsa phylogeny-based classification system of KS (PKS) and C (NRPS)domain sequences which extracts and classifies KS and C domainsfrom a wide range of sequence data; the results can be used to assessthe potential for PKS and NRPS secondary metabolite biosynthesis in agiven organism [32]. A preliminary candidate screening was performedfor all 89 KS sequences in NaPDoS and KS domains were confirmed andidentified for all queried sequences. Afterwards, these sequences werecompared against the reference database to identify matches toknown PKS and NRPS pathways. This preliminary analysis includedthe query identification, best database match, sequence identity andclassification of the biosynthetic pathway associated with the bestmatch. From the 89 sequences, biosynthetic gene clusters correspond-ing to 13 different compounds were identified (Fig. 1a, SupplementaryTable S1). BLAST hits for KS domains with more than 85% identity atthe amino acid level indicated that the query domains might be associ-ated with the production of the same or a similar compound as thoseproduced by the reference pathway [32]. In this analysis, all of the iden-tities obtained with the characterized domains in the NaPDoS databasewere low (less than 80%) indicating that the corresponding biosyntheticgene clusters are probably not yet characterized and that the encodedcompound may be new.

For NRPS analysis the PKS/NRPS analysis, one of the specificityprediction tools that have been developed, was used. The 27 A domainsequences were analysed and the information obtained is shown inSupplementary Table S2. This preliminary analysis comprises the bestdatabase match, the signature sequence and the predicted amino acid.From the 27 sequences in our dataset, biosynthetic gene clusterscorresponding to 10 different compounds were identified. Five NRPSA-domains from 5 different cyanobacterial strains had no correspon-dence in the databases, suggesting that the biosynthetic gene clustershave not yet been characterized (Fig. 1b and Supplementary Table S2).It is important to stress that these are predictions, and therefore,interpretation of these findings should be conservative; however,these results unquestionably highlight the potential of these strains toproduce a diverse range of secondary metabolites.

3.3. Transcriptional analysis

RT-PCR analyses of several of these isolates were performed to de-termine if the PKS/NRPS geneswere transcribedunder routine laborato-ry growth conditions. For this purpose we selected the faster growingcyanobacterial strains in order to be able to evaluate transcription inat least two different time points of the growth phase (strains andprimer pairs listed in Table 1). The strains and primer pairs for which

Table 2PCR screening for sequences encoding the ketosynthase domain (KS) of PKS genes and theadenylation-domain (A) of NRPS genes in cyanobacterial isolates from intertidal zones ofthe Portuguese coast.

Order (Subsection)Genus and species (LEGE code)

PKS NRPS

Chroococcales (Subsection I)Cyanobium sp. (06012) + −Cyanobium sp. (06097) + −Cyanobium sp. (06098) + +Cyanobium sp. (06109) + +Cyanobium sp. (06113) + −Cyanobium sp. (06127) − +Cyanobium sp. (06130) + −Cyanobium sp. (06134) + −Cyanobium sp. (06137) − −Cyanobium sp. (06140) − +Cyanobium sp. (06143) + −Cyanobium sp. (06184) + −Cyanobium sp. (07153) + −Cyanobium sp. (07175) − −Cyanobium sp. (07186) + −Gloeocapsopsis cf. crepidinum (06123) − +Synechococcus nidulans (07171) − −Synechococcus sp. (07172) − −Synechocystis salina (06099) + +Synechocystis salina (06155) − −Synechocystis salina (07173) + +

Pleurocapsales (Subsection II)Chroococcidiopsis sp. (06174) + +Chroococcopsis sp. (07161) + −Chroococcopsis sp. (07187) + +Hyella sp. (07179) + +Myxosarcina sp. (06146) + +

Oscillatoriales (Subsection III)Nodosilinea nodulosaa (06102) + −Nodosilinea nodulosaa (06110) + −Nodosilinea nodulosaa (06152) + +Leptolyngbya fragilis (07167) + −Leptolyngbya fragilis (07176) + −Leptolyngbya mycoidea (06009) + −Leptolyngbya mycoidea (06108) + −Leptolyngbya mycoidea (06118) + +Leptolyngbya mycoidea (06126) + +Leptolyngbya mycoidea (07157) − −Leptolyngbya saxicola (07132) + +Leptolyngbya saxicola (07170) + +Leptolyngbya sp. 1 (06121) + −Leptolyngbya sp. 2 (06188) + −Phormidium laetevirens (06103) + +Phormidium sp. 1 (06111) + +Plectonema cf. radiosum (06105) + +Pseudanabaena aff. curta (07160) + +Pseudanabaena aff. curta (07169) + −Pseudanabaena aff. persicina (07163) + +Pseudanabaena cf. frigida (06144) + +Pseudanabaena sp. (06194) + +Pseudanabaena sp. 2 (06129) + +Pseudanabaena sp. 3 (06116) + −Pseudanabaena sp. 3 (07190) + −Romeria sp. (06013) + +Schizothrix aff. septentrionalis (07164) + +

Nostocales (Subsection IV)Calothrix sp. 1 (06100) + +Calothrix sp. 2 (06122) + +Calothrix sp. 2 (07177) + +Nostoc sp. 1 (06106) + +Nostoc sp. 1 (06150) + +Nostoc sp. 2 (06158) + +Rivularia sp. (07159) + +Scytonema sp. (07189) + +

+, PCR amplification;−, no PCR amplification. For more details on the sampling sites/or-ganisms see [16].

a Formerly assigned to Leptolyngbya cf. halophila [37,38].


we obtained positive RT-PCR results/transcription (Fig. 2) are highlight-ed in grey in Table 1. From these results we found that under routinelaboratory growth conditions, the PKS encoding genes were transcribedfor three out of the nine strains tested, whereas the NRPS encodinggenes were transcribed in two out of three strains.

3.4. LC–MS and molecular networking studies

LC–MS analysis coupled withmolecular networking was carried outin order to investigate the production by our cyanobacterial strains ofknown metabolites, their analogues or novel compounds. MS/MSnetworking analysis is a mass spectrometric technique that clustersmetabolites on the basis of MS/MS fragmentation patterns [21]. Thistool has been previously applied in order to construct a global view of

the secondary metabolome of a given organism [21,34,44]. Cytoscapesoftware provides a visual representation of the molecular networkwhere each node (circle) represents a single consensus MS/MSspectrum for a given parent mass, and the relatedness between eachnode is defined by an edge (lines). The thickness of the edge definesthe degree of similarity of the MS/MS spectra. Molecular networks aregenerated using cosine scores which measure relatedness in the MS/MS spectra; a cosine value of 1.0 indicates identical spectra [21,44].The mass-spectral data set used in this work was obtained via LC–MS/MS analysis and the network was generated with the MS/MS spectraof 57 of the 61 cyanobacterial strains and also the MS/MS spectra of105 reference cyanobacterial metabolites (seed compounds).

In the resulting network (Fig. 3) it is possible to discern clusters ofnodes representing families of structurally related molecules. These

Fig. 1. Frequency of the in silico predicted PKS and NRPS genes in 61marine cyanobacterial strains from Portugal using (a) the NaPDoS software for PKS genes and (b) the PKS/NRPS anal-ysis tool for the NRPS genes. Abbreviations for the proposed compound produced by gene clusters: Bleom— bleomycin; Cur— curacin; Epo— epothilone; HSAF— heat stable antifungalfactor; Jam — jamaicamide; Kirr — kirromycin; Lnm — leinamycin; Lovas — lovastatin; Mta — myxothiazol; Mxa — myxalamide; Nos — nostopeptolide; Sti — stigmatellin; Vir —virginiamycin; Adp — anabaenopeptilide; Cch — coelichelin; Mcy — microcystin; Myc — mycosubtilin; Phs — phosphinothricin; Pvd — pyoverdin; Saf — saframycin; Srf — surfactin;and Tyc — tyrocidine.

Fig. 2. Transcription of genes encoding the KS and A domains of the PKS and NRPS genes evaluated by RT-PCR. The cells were collected from 2-month old cultures and then cDNAs wereproducedwith randomprimers andused in PCR amplificationswith specific primers (Table 1). 1 and 1′—Myxosarcina sp. (LEGE 06146) sequences KC842286 and KC842288, respectively;2 and 2′ — Pseudanabaena aff. curta (LEGE 07160) KC842368 and KC842370; 3 — Calothrix sp. 1 (LEGE 06100) KC842315; 4 — Cyanobium sp. (LEGE 06109) KC813182; 5 — Nostoc sp. 2(LEGE 06158) KC813190; M — marker, GeneRuler™ DNA Ladder Mix, (Fermentas Waltham, MA, USA); c− — negative controls: no template cDNA; c+ positive controls: genomic DNA.


results, togetherwith the genetic analysis, confirm themetabolic poten-tial of our strains. In the network, nodes are represented by differentcolours depending on their origin: green nodes — seed compounds,pink nodes — samples, and red nodes — consensus of samples andseed spectra. Inclusion of the MS/MS data of 105 reference compoundsin this analysis was highly useful as 41 formed consensus orwere highlycorrelated with compounds retrieved from our samples. Some exam-ples are highlighted below:

i) A red node with mass m/z 759.421 (Hyella sp. LEGE 07179)exhibited a consensus with naopopeptin [M + Na]+ (m/z =759.4) (Fig. 3a); however, despite the fact that they have thesame mass, this result was not confirmed since the retentiontimes and the MS/MS fragmentation patterns were different.Moreover, the same node is connected with antanapeptin B[M + Na]+ (m/z 761.39) and antanapeptin C [M + Na]+ (m/z763.37) forming a three-node cluster (Fig. 3a). Both naopopeptinand antanapeptins B and C are cyclic depsipeptides [45,46]

containing the Hmoya (3-hydroxy-2-methyl-7-octynoic acid)unit or its derivatives (Hmoea or Hmoaa). Since the isolation ofkulolide, the first compound reported to contain the 2,2-dimeth-yl-3-hydroxy-7-octynoic acid (Dhoya) [47], a variety of structur-ally similar compounds have been isolated from diversecyanobacteria [45,48,49]. The mass we recorded for this node(759.421)was queried against theMarinLit database (Universityof Canterbury, New Zealand) and there were several possibilitiesidentified, one of which was antanapeptin A. Altogether, theseresults suggests that Hyella sp. LEGE 07179 likely produces acompound related to naopopeptin or antanapeptin.

ii) The nodewithmassm/z 929.7 (Chroococcopsis sp. LEGE 07187) isconnected in the network with dolastatin 16 [M+ Na]+ (m/z=901.4). This node represents a compound 28 Da larger thandolastatin 16, and theMS/MS fragmentation patterns of our com-pound anddolastatin are similar (Fig. 3b). This family of bioactivecompounds has been identified in several cyanobacterial taxa[45,50,51], and therefore, this node probably represents an

Fig. 3.MS/MS-basedmolecular network of the extracts from 57 cultured cyanobacterial strains seededwith 105 reference compounds using a cosine similarity score cut-off of 0.6. Greennodes— seeded compounds; pinknodes— samples; red nodes—match between seed and samples. Clusters highlighted in network: (a) three-node cluster composed bynaopopeptin andantanapeptins B and C; (b) seed dolastatin-16, potential analogue of dolastatin-16 and MS/MS spectra of both compounds; (c) malyngamide family cluster composed by the seedsmalyngamide C, malyngamide C acetate, malyngamide H, malyngamide I, malyngamide K, malyngamide L/T and malyngamide U/V/W and nodes from different Portuguese marinecyanobacterial samples; (d) cluster of microcystin, nodularin and nodes from the Portuguese cyanobacterial samples; and (e) cluster composed only of nodes from different extracts ofour cyanobacterial strains. Chemical structures retrieved from: naopopeptin [46]; antanapeptins A–C and dolastatin 16 [45]; malyngamides H, K [54]; and microcystin and nodularin[8]. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)


analogue of dolastatin 16. A structure of dolastatin 16 analogueion of 760 is also proposed (Supplementary Fig. S1).

iii) The network also contained amalyngamide cluster comprised ofthe seed compounds malyngamide C [M+H]+ (m/z=455.92),malyngamide C acetate [M+H]+ (m/z=497.95) malyngamideH [M+ H]+ (m/z = 432.01), malyngamide I [M + H]+ (m/z =484.04), malyngamide K [M + H]+ (m/z = 423.97),

malyngamide L/T [M + H]+ (m/z = 467.99) and malyngamideU/V/W [M+H]+ (m/z=409.95) as well as related nodes deriv-ing frommetabolites in our cyanobacterial strains (Fig. 3c). All ofthe new masses appearing in this cluster were also queriedagainst the MarinLit database; in the m/z 483–484 range (node:m/z 484.05) one known metabolite was malyngamide S. Takentogether, these data strongly suggest that malyngamide-type

Fig. 3 (continued).


molecules are present in our cyanobacterial samples. A large arrayof malyngamides has been reported in various cyanobacteria andthey are characterized by a fatty acid chain, commonly lyngbicacid, attached to a cyclic head group with nitrogen and oxygenfunctional groups [52–54].

iv) In another casewe could observe amolecular network connectionbetween nodularin, microcystin and nodes from the samplesm/z516.982 (Leptolyngbya fragilis LEGE 07167), m/z 623.33 (L. fragilisLEGE 07176) andm/z 350.02 (Cyanobium sp. LEGE 07175, Romeriasp. LEGE 06013) (Fig. 3d), revealing that these cyanobacterialstrains may produce compounds related to these hepatotoxins.

The results shown in Fig. 3 also strongly suggest that ourcyanobacterial strains produce novel compounds (Fig. 3e). However,the few molecular network connections obtained with the seedcompoundsmay result because our isolates are froma temperate regionand the standards were isolated from tropical cyanobacteria. Indeed, itis well known that environmental factors can strongly influence theproduction of secondary metabolites from cyanobacteria [55–57].Furthermore, for each strain the MS spectra were checked for majorpeaks and the masses were queried against the MarinLit database. Forsome masses, m/z 1785.41 (Cyanobium sp. LEGE 06109), m/z 1957.64(Calothrix sp. LEGE 07177) and m/z 1746.46 (Pseudanabaena persicinaLEGE 07163) no compound with a corresponding mass was found inthe MarinLit database, suggesting that these strains produce com-pounds not yet identified and thus deserving of further investigation.

Overall, this work reinforces the hypothesis that prospecting newcyanobacterial diversity is a valid and promising approach for the dis-covery of novel bioactive compounds. Here, we demonstrated by genet-ic and analytical approaches that our set of isolates possess an untappedsource of secondary metabolites and highlight the potential of marinecyanobacterial strains from temperate regions to produce novel bioac-tive compounds.

4. Conclusions

The results from this work demonstrated that both PKS and NRPSgenes are present in the majority of the 61 cyanobacteria isolated fromthe Portuguese Atlantic coast, with the former biosynthetic class beingmore ubiquitous than the latter. As expected, a higher frequency ofthese genes was detected in the filamentous strains. Furthermore, itwas possible to demonstrate that, under routine laboratory growth con-ditions, some of these genes are transcribed in a set of selected strains.Moreover, LC–MS analysis together withmolecular networking is a pow-erful tool that revealed the presence of analogues in known compoundfamilies as well as novel compounds. Although these latter findingsshould be regarded as indicative rather than definitive, they unquestion-ably highlight the metabolic potential of these cyanobacterial isolates toproduce many and diverse natural products. Moreover, these resultswill allow the selection of the most promising strains for further studies(e.g. isolation and characterization of bioactivemolecules) and constitutea valuable starting point for the use of marine cyanobacteria from tem-perate regions for the discovery of natural compounds with biomedicalvalue.

Acknowledgements

This work was funded by FEDER funds through the OperationalCompetitiveness Programme — COMPETE and by National Fundsthrough FCT— Fundação para a Ciência e a Tecnologia under the projectFCOMP-01-0124-FEDER-010600 (PTDC/MAR/102258/2008) and grantsSFRH/BD/70284/2010 (to AB) and SFRH/BPD/95683/2013 (toMVM), aswell as NIH grants NS053398 and GM107550 (to PCD and WHG). Weare grateful to Rosário Martins for supplying six of the isolates andPedro Leão for the valuable discussions and advice.

Appendix A. Supplementary data

Supplementary data to this article can be found online at http://dx.doi.org/10.1016/j.algal.2015.03.016.

References

[1] I.M. Ehrenreich, J.B. Waterbury, E.A. Webb, Distribution and diversity of naturalproduct genes in marine and freshwater cyanobacterial cultures and genomes,Appl. Environ. Microbiol. 71 (2005) 7401–7413.

[2] A.C. Jones, L. Gu, C.M. Sorrels, D.H. Sherman, W.H. Gerwick, New tricks from ancientalgae: natural products biosynthesis inmarine cyanobacteria, Curr. Opin. Chem. Biol.13 (2009) 216–223.

[3] A.C. Jones, E.A. Monroe, E.B. Eisman, L. Gerwick, D.H. Sherman, W.H. Gerwick, Theuniquemechanistic transformations involved in the biosynthesis of modular naturalproducts from marine cyanobacteria, Nat. Prod. Rep. 27 (2010) 1048–1065.

[4] J.K. Nunnery, E. Mevers, W.H. Gerwick, Biologically active secondary metabolitesfrom marine cyanobacteria, Curr. Opin. Biotechnol. 21 (2010) 787–793.

[5] J.C. Kehr, D. Gatte Picchi, E. Dittmann, Natural product biosyntheses in cyanobacteria: atreasure trove of unique enzymes, Beilstein J. Org. Chem. 7 (2011) 1622–1635.

[6] A. Méjean, O. Ploux, A genomic view of secondary metabolite production incyanobacteria, in: F. Chauvat, C. Cassier-Chauvat (Eds.), Genomics of Cyanobacteria,Elsevier Academic Press, San Diego, 2013, pp. 189–234.

[7] C. Wiegand, S. Pflugmacher, Ecotoxicological effects of selected cyanobacterial sec-ondary metabolites: a short review, Toxicol. Appl. Pharmacol. 203 (2005) 201–218.

[8] E. Dittmann, D.P. Fewer, B.A. Neilan, Cyanobacterial toxins: biosynthetic routes andevolutionary roots, FEMS Microbiol. Rev. 37 (2013) 23–43.

[9] A.M. Burja, B. Banaigs, E. Abou-Mansour, J.G. Burgess, P.C. Wright, Marinecyanobacteria — a prolific source of natural products, Tetrahedron 57 (2001)9347–9377.

[10] R.K. Singh, S.P. Tiwari, A.K. Rai, T.M. Mohapatra, Cyanobacteria: an emerging sourcefor drug discovery, J. Antibiot. (Tokyo) 64 (2011) 401–412.

[11] L.T. Tan, Filamentous tropical marine cyanobacteria: a rich source of naturalproducts for anticancer drug discovery, J. Appl. Phycol. 22 (2010) 659–676.

[12] L.T. Tan, Pharmaceutical agents from filamentous marine cyanobacteria, DrugDiscov. Today 18 (2013) 863–871.

[13] W.H. Gerwick, R.C. Coates, N. Engene, L. Gerwick, R.V. Grindberg, A.C. Jones, C.M.Sorrels, Giant marine cyanobacteria produce exciting potential pharmaceuticals,Microbe 6 (2008) 277–284.

[14] M.E. Barrios-Llerena, A.M. Burja, P.C.Wright, Genetic analysis of polyketide synthaseand peptide synthetase genes in cyanobacteria as a mining tool for secondarymetabolites, J. Ind. Microbiol. Biotechnol. 34 (2007) 443–456.

[15] N.V. Wase, P.C. Wright, Systems biology of cyanobacterial secondary metaboliteproduction and its role in drug discovery, Expert Opin. Drug Discov. 3 (2008)903–929.

[16] A. Brito, V. Ramos, R. Seabra, A. Santos, C.L. Santos, M. Lopo, S. Ferreira, A. Martins, R.Mota, B. Frazao, R. Martins, V. Vasconcelos, P. Tamagnini, Culture-dependentcharacterization of cyanobacterial diversity in the intertidal zones of the Portuguesecoast: a polyphasic study, Syst. Appl. Microbiol. 35 (2012) 110–119.

[17] R. Martins, N. Fernandez, R. Beiras, V. Vasconcelos, Toxicity assessment of crude andpartially purified extracts of marine Synechocystis and Synechococcus cyanobacterialstrains in marine invertebrates, Toxicon 50 (2007) 791–799.

[18] M. Costa, M. Garcia, J. Costa-Rodrigues, M.S. Costa, M.J. Ribeiro, M.H. Fernandes, P.Barros, A. Barreiro, V. Vasconcelos, R. Martins, Exploring bioactive properties ofmarine cyanobacteria isolated from the Portuguese coast: high potential as a sourceof anticancer compounds, Mar. Drugs 12 (2014) 98–114.

[19] P.N. Leão, V. Ramos, P.B. Goncalves, F. Viana, O.M. Lage, W.H. Gerwick, V.M.Vasconcelos, Chemoecological screening reveals high bioactivity in diverseculturable Portuguese marine cyanobacteria, Mar. Drugs 11 (2013) 1316–1335.

[20] P.N. Leão, M. Costa, V. Ramos, A.R. Pereira, V.C. Fernandes, V.F. Domingues, W.H.Gerwick, V.M. Vasconcelos, R. Martins, Antitumor activity of hierridin B, acyanobacterial secondary metabolite found in both filamentous and unicellularmarine strains, PLoS One 8 (2013) e69562.

[21] J. Watrous, P. Roach, T. Alexandrov, B.S. Heath, J.Y. Yang, R.D. Kersten, M. van derVoort, K. Pogliano, H. Gross, J.M. Raaijmakers, Mass spectral molecular networkingof living microbial colonies, Proc. Natl. Acad. Sci. U. S. A. 109 (2012) E1743–E1752.

[22] R. Rippka, Isolation and purification of cyanobacteria, in: L. Packer, A.N. Glazer(Eds.), Method. Enzymol, Academic Press, 1988, pp. 3–27.

[23] R. Stanier, R. Kunisawa, M. Mandel, G. Cohen-Bazire, Purification and properties ofunicellular blue-green algae (order Chroococcales), Bacteriol. Rev. 35 (1971) 171.

[24] J. Kotai, Instructions for Preparation of Modified Nutrient Solution Z8 for Algae.Publication B-11/69, Norwegian Institute for Water Research, Blindern, Oslo, 1972.

[25] R. Rippka, J. Deruelles, J.B. Waterbury, M. Herdman, R.Y. Stanier, Genericassignments, strain histories and properties of pure cultures of cyanobacteria, J.Gen. Microbiol. 111 (1979) 1–61.

[26] P. Tamagnini, O. Troshina, F. Oxelfelt, R. Salema, P. Lindblad, Hydrogenases in Nostocsp. strain PCC 73102, a strain lacking a bidirectional enzyme, Appl. Environ.Microbiol. 63 (1997) 1801–1807.

[27] M.C. Moffitt, B.A. Neilan, On the presence of peptide synthetase and polyketidesynthase genes in the cyanobacterial genus Nodularia, FEMS Microbiol. Lett. 196(2001) 207–214.

[28] B.A. Neilan, E. Dittmann, L. Rouhiainen, R.A. Bass, V. Schaub, K. Sivonen, T. Börner,Nonribosomal peptide synthesis and toxigenicity of cyanobacteria, J. Bacteriol. 181(1999) 4089–4097.




http://refhub.elsevier.com/S2211-9264(15)00079-X/rf0005













































































[29] J. Sambrook, D.W. Russell, Molecular Cloning: A Laboratory Manual, 3rd ed. ColdSpring Harbor Laboratory Press, New York, 2001.

[30] V. Ramos, R. Seabra, A. Brito, A. Santos, C.L. Santos, M. Lopo, P. Moradas-Ferreira,V.M. Vasconcelos, P. Tamagnini, Characterization of an intertidal cyanobacteriumthat constitutes a separate clade together with thermophilic strains, Eur. J. Phycol.45 (2010) 394–403.

[31] M.A. Larkin, G. Blackshields, N. Brown, R. Chenna, P.A. McGettigan, H. McWilliam, F.Valentin, I.M. Wallace, A. Wilm, R. Lopez, Clustal W and Clustal X version 2.0,Bioinformatics 23 (2007) 2947–2948.

[32] N. Ziemert, S. Podell, K. Penn, J.H. Badger, E. Allen, P.R. Jensen, The natural productdomain seeker NaPDoS: a phylogeny based bioinformatic tool to classify secondarymetabolite gene diversity, PLoS One 7 (2012) e34064.

[33] B.O. Bachmann, J. Ravel, Methods for in silico prediction of microbial polyketide andnonribosomal peptide biosynthetic pathways from DNA sequence data, ComplexEnzymes in Microbial Natural Product Biosynthesis, Part A: Overview Articles andPeptides, Elsevier Academic Press Inc., San Diego, 2009. 181–217.

[34] J.R. Winnikoff, E. Glukhov, J. Watrous, P.C. Dorrestein, W.H. Gerwick, Quantitativemolecular networking to profile marine cyanobacterial metabolomes, J. Antibiot.67 (2014) 105–112.

[35] R. Araújo, I. Bárbara, M. Tibaldo, E. Berecibar, P.D. Tapia, R. Pereira, R. Santos, I. Sousa-Pinto, Checklist of benthic marine algae and cyanobacteria of northern Portugal, Bot.Mar. 8 (2009) 1908–1919.

[36] B. Frazao, R. Martins, V. Vasconcelos, Are known cyanotoxins involved in the toxicityof picoplanktonic and filamentous north Atlantic marine cyanobacteria? Mar. Drugs8 (2010) 1908–1919.

[37] Z.K. Li, J. Brand, Leptolyngbya nodulosa sp. nov. (Oscillatoriaceae), a subtropical ma-rine cyanobacterium that produces a unique multicellular structure, Phycologia 46(2007) 396–401.

[38] R.B. Perkerson, J.R. Johansen, L. Kovacik, J. Brand, J. Kastovsky, D.A. Casamatta, Aunique Pseudanabaenalean (Cyanobacteria) genus Nodosilinea gen. nov. based onmorphological and molecular data, J. Phycol. 47 (2011) 1397–1412.

[39] G. Christiansen, E. Dittmann, L.V. Ordorika, R. Rippka, M. Herdman, T. Borner,Nonribosomal peptide synthetase genes occur in most cyanobacterial genera as ev-idenced by their distribution in axenic strains of the PCC, Arch. Microbiol. 176(2001) 452–458.

[40] P.M. Shih, D.Wu, A. Latifi, S.D. Axen, D.P. Fewer, E. Talla, A. Calteau, F. Cai, N. TandeaudeMarsac, R. Rippka, M. Herdman, K. Sivonen, T. Coursin, T. Laurent, L. Goodwin, M.Nolan, K.W. Davenport, C.S. Han, E.M. Rubin, J.A. Eisen, T. Woyke, M. Gugger, C.A.Kerfeld, Improving the coverage of the cyanobacterial phylum using diversity-driven genome sequencing, Proc. Natl. Acad. Sci. U. S. A. 110 (2013) 1053–1058.

[41] A. Calteau, D.P. Fewer, A. Latifi, T. Coursin, T. Laurent, J. Jokela, C.A. Kerfeld, K.Sivonen, J. Piel, M. Gugger, Phylum-wide comparative genomics unravel thediversity of secondary metabolism in Cyanobacteria, BMC Genomics 15 (2014) 977.

[42] E. Dittmann, B.A. Neilan, T. Borner, Molecular biology of peptide and polyketide bio-synthesis in cyanobacteria, Appl. Microbiol. Biotechnol. 57 (2001) 467–473.

[43] R.W. Castenholz, Phylum BX. Cyanobacteria. Oxygenic photosynthetic bacteria, in:D.R. Boone, R.W. Castenholz, G.M. Garrity (Eds.), Bergey's Manual® of SystematicBacteriology, Springer, New York, 2001, pp. 473–599.

[44] J.Y. Yang, L.M. Sanchez, C.M. Rath, X. Liu, P.D. Boudreau, N. Bruns, E. Glukhov, A.Wodtke, R. de Felicio, A. Fenner, Molecular networking as a dereplication strategy,J. Nat. Prod. 76 (2013) 1686–1699.

[45] L.M. Nogle, W.H. Gerwick, Isolation of four new cyclic depsipeptides, antanapeptinsA–D, and dolastatin 16 from a Madagascan collection of Lyngbya majuscula, J. Nat.Prod. 65 (2002) 21–24.

[46] K.L. Malloy, Structure Elucidation of Biomedically Relevant Marine CyanobacterialNatural Products, UC San Diego, San Diego, 2011.

[47] M.T. Reese, N.K. Gulavita, Y. Nakao, M.T. Hamann, W.Y. Yoshida, S.J. Coval, P.J.Scheuer, Kulolide: a cytotoxic depsipeptide from a cephalaspidean mollusk,Philinopsis speciosa 1, J. Am. Chem. Soc. 118 (1996) 11081–11084.

[48] P.D. Boudreau, T. Byrum, W.T. Liu, P.C. Dorrestein, W.H. Gerwick, Viequeamide A, acytotoxic member of the kulolide superfamily of cyclic depsipeptides from a marinebutton cyanobacterium, J. Nat. Prod. 75 (2012) 1560–1570.

[49] R. Montaser, V.J. Paul, H. Luesch, Pitipeptolides C–F, antimycobacterialcyclodepsipeptides from the marine cyanobacterium Lyngbya majuscula fromGuam, Phytochemistry 72 (2011) 2068–2074.

[50] H. Luesch, R.E. Moore, V.J. Paul, S.L. Mooberry, T.H. Corbett, Isolation of dolastatin 10from the marine cyanobacterium Symploca species VP642 and total stereochemistryand biological evaluation of its analogue symplostatin 1, J. Nat. Prod. 64 (2001)907–910.

[51] G.G. Harrigan, W.Y. Yoshida, R.E. Moore, D.G. Nagle, P.U. Park, J. Biggs, V.J. Paul, S.L.Mooberry, T.H. Corbett, F.A. Valeriote, Isolation, structure determination, and biolog-ical activity of dolastatin 12 and lyngbyastatin 1 from Lyngbya majuscula/Schizothrixcalcicola cyanobacterial assemblages, J. Nat. Prod. 61 (1998) 1221–1225.

[52] R.D. Ainslie, J.J. Barchi Jr., M. Kuniyoshi, R.E. Moore, J.S. Mynderse, Structure ofmalyngamide C, J. Org. Chem. 50 (1985) 2859–2862.

[53] J. Orjala, D. Nagle, W.H. Gerwick, Malyngamide H, an ichthyotoxic amide possessinga new carbon skeleton from the Caribbean cyanobacterium Lyngbya majuscula, J.Nat. Prod. 58 (1995) 764–768.

[54] H. Gross, K.L. McPhail, D.E. Goeger, F.A. Valeriote, W.H. Gerwick, Two cytotoxic ste-reoisomers of malyngamide C, 8-epi-malyngamide C and 8-O-acetyl-8-epi-malyngamide C, from the marine cyanobacterium Lyngbya majuscula, Phytochemis-try 71 (2010) 1729–1735.

[55] M. Kaebernick, B.A. Neilan, T. Börner, E. Dittmann, Light and the transcriptional re-sponse of the microcystin biosynthesis gene cluster, Appl. Environ. Microbiol. 66(2000) 3387–3392.

[56] G. Shalev‐Malul, J. Lieman‐Hurwitz, Y. Viner‐Mozzini, A. Sukenik, A. Gaathon, M.Lebendiker, A. Kaplan, An AbrB‐like protein might be involved in the regulation ofcylindrospermopsin production by Aphanizomenon ovalisporum, Environ. Microbiol.10 (2008) 988–999.

[57] C.M. Sorrels, P.J. Proteau, W.H. Gerwick, Organization, evolution, and expressionanalysis of the biosynthetic gene cluster for scytonemin, a cyanobacterial UV-absorbing pigment, Appl. Environ. Microbiol. 75 (2009) 4861–4869.






























































































Supplementary Material III

Table S1 Cyanobacterial KS domain sequence based prediction of PKS using the NaPDoS software.

Organism (LEGE code) Accession

Number Database match ID % Identity E value

Pathway

Product

Domain

Class Cluster type

Cyanobium sp. (06012) KC842333 CurA_AAT70096_mod 53 3e-56 Curacin KS PKS

Cyanobium sp. (06097) KC842336 VirF_BAF50722_5T 47 8e-54 Virginiamycin trans PKS

Cyanobium sp. (06109) KC842322 KirAIV_CAN89634_9T 55 1e-69 Kirromycin trans PKS

KC842323 LnmJ_AF484556_4T 54 8e-65 Leinamycin trans PKS

KC842324 VirF_BAF50722_5T 55 1e-66 Virginiamycin trans PKS

KC842325 KirAII_CAN89632_5T 47 1e-54 Kirromycin trans PKS

Cyanobium sp. (06113) KC842298 CurA_AAT70096_mod 54 2e-66 Curacin KS PKS

Cyanobium sp. (06130) KC842364 LnmJ_AF484556_4T 55 6e-69 Leinamycin trans PKS


Cyanobium sp. (06134) KC842299 CurL_AAT70107_mod 52 9e-63 Curacin modular PKS

Cyanobium sp. (06184) KC842354 KirAII_CAN89632_5T 44 3e-48 Kirromycin trans PKS

Cyanobium sp. (07186) KC842302 CurL_AAT70107_mod 53 7e-66 Curacin modular PKS

KC842303 Strep_ZP_06279092_i 44 5e-50 Unknown iterative

Synechocystis salina (06099) KC842284 KirAII_CAN89632_5T 47 2e-50 Kirromycin trans PKS

Synechocystis salina (07173) KC842372 CurL_AAT70107_mod 54 3e-67 Curacin modular PKS

Chroococcidiopsis sp. (06174) KC842328 LOVAS_Q9Y8A5_i 45 1e-53 Lovastatin iterative PKS

KC842329 KirAI_CAN89631_1T 38 4e-44 Kirromycin trans PKS

Chroococcopsis sp. (07161) KC842343 KirAII_CAN89632_5T 47 3e-51 Kirromycin trans PKS

KC842344 StiG_Q8RJY0_1KSB 55 8e-59 Stigmatellin modular PKS

Chroococcopsis sp. (07187) KC842293 CurI_AAT70104_mod 63 3e-74 Curacin modular PKS


Hyella sp. (07179) KC842291 LnmJ_AF484556_4T 56 8e-67 Leinamycin trans PKS


Myxosarcina sp. (06146) KC842286 MtaF_Q9RFK6_1KSB 55 2e-66 Myxothiazol modular PKS

KC842287 KirAIV_CAN89634_9T 48 1e-50 Kirromycin trans PKS


Nodosilinea nodulosa* (06102) KC842304 EpoF_Q9L8C5_1mod 58 3e-63 Epothilone modular PKS

KC842305 MxaC_Q93TW9_1KSB 60 2e-65 Myxalamide modular PKS

Nodosilinea nodulosa* (06110) KC842285 bleom_AAG02357_H 47 1e-50 Bleomycin hybridKS Hybrid

2

Nodosilinea nodulosa* (06152) KC842316 MxaC_Q93TW9_1KSB 59 9e-74 Myxalamide modular PKS

Leptolyngbya fragilis (07167) KC842317 NosB_Q9RAH3_H 50 1e-59 Nostopeptolide hybridKS PKS

Leptolyngbya fragilis (07176) KC842330 CurI_AAT70104_mod 58 2e-64 Curacin modular PKS

Leptolyngbya mycoidea (06009) KC842331 StiB_Q8RJY5_1KSB 59 3e-71 Stigmatellin modular PKS

KC842332 JamP_AAS98787_H 66 2e-74 Jamaicamide hybridKS PKS

Leptolyngbya mycoidea (06108) KC842357 NosB_Q9RAH3_H 59 1e-66 Nostopeptolide hybridKS PKS

Leptolyngbya mycoidea (06118) KC842358 NosB_Q9RAH3_H 59 1e-64 Nostopeptolide hybridKS PKS

Leptolyngbya mycoidea (06126) KC842347 JamP_AAS98787_H 65 2e-61 Jamaicamide hybridKS PKS

Leptolyngbya saxicola (07132) KC842359 JamK_AAS98782_mod 61 1e-73 Jamaicamide modular PKS

KC842360 JamE_AAS98777_KS1 62 4e-69 Jamaicamide KS PKS

KC842361 CurI_AAT70104_mod 59 1e-71 Curacin modular PKS

Leptolyngbya saxicola (07170) KC842371 JamM_AAS98784_H 60 9e-82 Jamaicamide hybridKS PKS

Leptolyngbya sp. 1 (06121) KC842362 EpoC_Q9L8C8_H 62 1e-71 Epothilone hybridKS PKS

KC842363 CurI_AAT70104_mod 74 2e-95 Curacin modular PKS

Leptolyngbya sp. 2 (06188) KC842318 CurG_AAT70102_H 57 6e-68 Curacin hybridKS PKS

KC842319 CurA_AAT70096_mod 46 3e-50 Curacin KS PKS

Phormidium sp. 1 (06111) KC842337 LnmJ_AF484556_4T 46 7e-55 Leinamycin trans PKS

KC842338 EpoC_Q9L8C8_H 60 7e-66 Epothilone hybridKS PKS



Plectonema cf. radiosum (06105) KC842355 JamK_AAS98782_mod 70 5e-95 Jamaicamide modular PKS

KC842356 StiC_Q8RJY4_1KSB 67 2e-79 Stigmatellin modular PKS

Pseudanabaena aff. curta (07160) KC842368 JamE_AAS98777_KS1 63 4e-70 Jamaicamide KS PKS

KC842369 CurG_AAT70102_H 61 1e-82 Curacin hybridKS PKS

KC842370 MtaF_Q9RFK6_1KSB 62 2e-78 Myxothiazol modular PKS

Pseudanabaena aff. curta (07169) KC842326 CurG_AAT70102_H 61 7e-83 Curacin hybridKS PKS

KC842327 NosB_Q9RAH3_H 66 3e-76 Nostopeptolide hybridKS PKS

Pseudanabaena aff. persicina (07163) KC842289 MxaB_Q93TX0_1KSB 58 9e-72 Myxalamide modular PKS

Pseudanabaena cf. frigida (06144) KC842348 StiB_Q8RJY5_1KSB 52 3e-67 Stigmatellin modular PKS

KC842349 JamK_AAS98782_mod 59 4e-67 Jamaicamide modular PKS

KC842350 CurA_AAT70096_mod 58 3e-73 Curacin KS PKS

Pseudanabaena sp. (06194) KC842300 EpoC_Q9L8C8_H 60

7e-66 Epothilone hybridKS PKS


3

Pseudanabaena sp. 3 (06116) KC842341 EpoC_Q9L8C8_H 59 4e-62 Epothilone hybridKS PKS


Pseudanabaena sp. 3 (07190) KC842320 CurL_AAT70107_mod 63 3e-78 Curacin modular PKS


Romeria sp. (06013) KC842334 KirAI_CAN89631_1T 38 3e-44 Kirromycin trans PKS

KC842335 bleom_AAG02357_H 45 6e-51 Bleomycin hybridKS Hybrid

Schizothrix aff. septentrionalis (07164) KC842290 NosB_Q9RAH3_H 61 1e-76 Nostopeptolide hybridKS PKS

Calothrix sp. 1 (06100) KC842315 StiG_Q8RJY0_1KSB 67 2e-84 Stigmatellin modular PKS

Calothrix sp. 2 (06122) KC842309 StiG_Q8RJY0_1KSB 66 4e-83 Stigmatellin modular PKS



Calothrix sp. 2 (07177) KC842345 CurJ_AAT70105_mod 66 8e-85 Curacin modular PKS


Nostoc sp. 1 (06106) KC842306 LnmJ_AF484556_4T 56 3e-78 Leinamycin trans PKS



Nostoc sp. 1 (06150) KC842312 JamK_AAS98782_mod 65 1e-84 Jamaicamide modular PKS

KC842313 StiE_Q8RJY2_1KSB 46 3e-53 Stigmatellin modular PKS

KC842314 HSAF_ABL86391_i 48 1e-51 HSAFa iterative

Nostoc sp. 2 (06158) KC842366 LnmJ_AF484556_4T 45 2e-57 Leinamycin trans PKS

KC842367 MtaF_Q9RFK6_1KSB 59 2e-74 Myxothiazol modular PKS

Rivularia sp. (07159) KC842351 CurG_AAT70102_H 70 2e-90 Curacin hybridKS PKS

KC842352 EpoE_Q9L8C6_1mod 60 2e-75 Epothilone modular PKS


Scytonema sp. (07189) KC842295 StiG_Q8RJY0_1KSB 68 2e-86 Stigmatellin modular PKS


KC842297 StiE_Q8RJY2_1KSB 57 1e-56 Stigmatellin modular PKS a HSAF: heat stable antifungal factor. * Formerly assigned to Leptolyngbya cf. halophila [1, 2].

.

4

Table S2 Cyanobacterial A domain sequence based prediction of NRPS using the PKS/NRPS analysis software.

Organism (LEGE code) Accession

Number Database match ID

Pathway

product

E

value

Signature

sequencea

Predicted aab

Cyanobium sp. (06098) KC813175 gi||genome sequencing|PvdD-M2-Arg|Pyoverdin synthetase Pyoverdin 0.28 DAEYIGAI PvdD-M2-Arg

Cyanobium sp. (06109) KC813182 gi|6563399|gb|AAF17280.1|NosC-M3 Asp/Asn?|Nostopeptolide synthetase Nostopeptolide 0.038 DATKIGEV NosC-M3-Asp/Asn

Gloeocapsopsis cf. crepidinum (06123) KC813188 gi|5822841|dbj|BAA83992.1|McyA-M2-Gly|Microcistin synthetase A Microcystin 0.026 DLFNNALT McyA-M2-Gly

KC813189 gi|6563397|gb|AAF15891.2|NosA-M2-Ser|Nostopeptolide synthetase Nostopeptolide 0.017 DVWHISLI NosA-M2-Ser

Synechocystis salina (06099) KC813168 gi|1805421|dbj|BAA08983.1|SrfAB-M2 Asp|surfactin synthetaseB Surfactin 0.082 DATKVGHV SrfAB-M2-Asp

KC813169 gi|5822841|dbj|BAA83992.1|McyA-M2-Gly|Microcistin synthetaseA Microcystin 0.046 DLYNNALT McyA-M2-Gly

Synechocystis salina (07173) KC813193 gi||genome sequencing|PvdD-M3-Ser|Pyoverdin synthetase Pyoverdin 0.017 DVWHVSLI PvdD-M3-Ser

KC813194 gi|9715734|gb|CAC01604.1|AdpB-M3-Thr|Anabaenopeptilide synthetase B Anabaenopeptilide 0.15 DVWNMGLI AdpB-M3-Thr

Hyella sp. (07179) KC813174 gi|5822841|dbj|BAA83992.1|McyA-M2-Gly|Microcistin synthetaseA Microcystin 0.083 DLFNNAL- McyA-M2-Gly

Myxosarcina sp. (06146) KC813170 gi|5763943|CchH-M2-Thr|Coelichelin synthetase Coelichelin 0.006 DFWNIGMV CchH-M2-Thr

Phormidium sp. 1 (06111) KC813183 gi|6449056|gb|AAF08796.1|MycB-M3-Gln|mycosubtilin synthetase B Mycosubtilin 0.17 DAEDLGTV MycB-M3-Gln

Plectonema cf. radiosum (06105) KC813187 NO HIT - NOVEL A-DOMAIN SIGNATURE No hit VDWVVSLA

Pseudanabaena aff. curta (07160) KC813192 gi|1171129|gb|AAC44129.1|SafA-M1-3h4mPhe|saframycin Mx1 synthetase. Saframycin 0.47 DILXLGMV SafA-M1-Phe

Pseudanabaena aff. persicina (07163) KC813171 gi|2623773|gb|AAC45930.1|TycC-M4-Val|tyrocidine synthetase 3 Tyrocidine 0.015 DAFWLGGT TycC-M4-Val

KC813172 gi|6563399|gb|AAF17280.1|NosC-M2-Gly|Nostopeptolide synthetase Nostopeptolide 0.073 DILQLGLI NosC-M2-Gly

Pseudanabaena sp. (06194) KC813177 NO HIT - NOVEL A-DOMAIN SIGNATURE No hit DALWLGXX

KC813178 gi||genome sequencing|PvdD-M2-Arg|Pyoverdin synthetase Pyoverdin 0.14 DAEDIGTV PvdD-M2-Arg

Schizothrix aff. septentrionalis (07164) KC813173 NO HIT - NOVEL A-DOMAIN SIGNATURE No hit VDWVCALA

Calothrix sp. 1 (06100) KC813180 gi|2623771|gb|AAC45928.1|TycA-M1-D-/L-Phe|tyrocidine synthetase 1 Tyrocidine 0.050 DAWTIAAV TycA-M1-D/L-Phe

KC813181 gi|1171129|gb|AAC44129.1|SafA-M1-3h4mPhe|saframycin Mx1 synthetase Saframycin 1.7 DILFIGVV SafA-M1-Phe

Calothrix sp. 2 (07177) KC813184 NO HIT - NOVEL A-DOMAIN SIGNATURE No hit DAIDSGG-

Nostoc sp. 1 (06106) KC813176 gi|6563397|gb|AAF15891.2|NosA-M2-Ser|Nostopeptolide synthetase Nostopeptolide 0.017 DVWHISLI NosA-M2-Ser

Nostoc sp. 1 (06150) KC813179 NO HIT - NOVEL A-DOMAIN SIGNATURE No hit DPWAIGCI

Nostoc sp. 2 (06158) KC813190 gi|5822842|dbj|BAA83993.1|McyB-M1-Leu|Microcistin synthetase B Microcystin 0.008 DAWFLGNV McyB-M1-Leu

KC813191 gi|2623772|gb|AAC45929.1|TycB-M2-L-Phe/L-Trp|tyrocidine synthetase Tyrocidine 0.50 DALFVGAV TycB-M2-Phe/Trp

Rivularia sp. (07159) KC813185 gi|8250617|PhsB-M1-Ser|Phosphitricin synthetase Phosphitricin 0.009 DVWHFSLI PhsB-M1-Ser

KC813186 gi|6449056|gb|AAF08796.1|MycB-M4-Pro|mycosubtilin synthetase B Mycosubtilin 0.019 DVQFIAHV MycB-M4-Pro

a Eight variable amino acids of the signature sequences determined as described [3].

b Nomenclature of the reference compounds as described [4].

5

Fig. S1. Proposed structure dolastatin16 analogue ion of 760.

6

References

[1] R.B. Perkerson, J.R. Johansen, L. Kovacik, J. Brand, J. Kastovsky, D.A. Casamatta, A

unique Pseudanabaenalean (Cyanobacteria) genus Nodosilinea gen. nov. based on

morphological and molecular data, J. Phycol. 47 (2011) 1397-1412.

[2] Z.K. Li, J. Brand, Leptolyngbya nodulosa sp nov (Oscillatoriaceae), a subtropical marine

cyanobacterium that produces a unique multicellular structure, Phycologia 46 (2007) 396-401.

[3] T. Stachelhaus, H.D. Mootz, M.A. Marahiel, The specificity-conferring code of

adenylation domains in nonribosomal peptide synthetases, Chem. Biol. 6 (1999) 493-505.

[4] G.L. Challis, J. Ravel, C.A. Townsend, Predictive, structure-based model of amino acid

recognition by nonribosomal peptide synthetase adenylation domains, Chem. Biol. 7 (2000)

211-224.

CHAPTER V

General Discussion, Conclusions

and Future Perspectives

FCUP Characterization of cyanobacteria isolated from the Portuguese coast

101

General Discussion, Conclusions and

Future Perspectives

5.1. General Discussion and Conclusions

The work presented here constitutes a comprehensive study on marine

cyanobacteria from the intertidal zones of the Portuguese Continental coast,

contributing to overcome the lack of information about this group of organisms in this

temperate Atlantic region. It includes the isolation, cultivation and characterization of

the cyanobacterial strains and the assessment of their potential to fix atmospheric

nitrogen and to produce secondary metabolites.

The intertidal zones of nine beaches (five in the west and four in the south)

presenting different topographies, morphologies and wave regimens were sampled.

More than 100 cyanobacterial strains were retrieved and 60 isolates were

characterized using a polyphasic approach (Chapter II). Based on morphological

characters, 35 morphotypes (15 genera and 16 species) belonging to all cyanobacterial

Orders/Subsections with the exception of Stigonematales/Subsection V (filamentous

true-branching cyanobacteria) were identified. This was expected since representatives

of this Order/Subsection are mainly encountered in flowing waters of hot springs and

soils (Gugger & Hoffmann, 2004; Finsinger et al., 2008; Soe et al., 2011). The most

representative groups among the isolates were the filamentous non-heterocystous

forms, Oscillatoriales and the unicellular Chroococcales, with a clear predominance of

the first. It is worthwhile to notice that the number of isolates belonging to a certain

genus is much higher than to others, but that these numbers can be due either to their

ubiquity or to the fact that certain strains are easier to isolate. Analysing the strains

origin and their geographical distribution, no clear spatial distribution was found. The

different genera were distributed by all the beaches types - rocky beaches, beaches

with sand and rocks and sandy beaches with dispersed rocks - and they were not

restricted to any specific areas of the coast (northwest, southwest or south).

To complement the morphological identification, the 16S rRNA gene was partially

sequenced for all isolates. For the majority of the isolates there was a quite good

102 FCUP Characterization of cyanobacteria isolated from the Portuguese coast

correlation between the phenotypic and genotypic based identifications. However,

some discrepancies were observed, possibly resulting from limitations on the

information available in the databases and taxonomic constraints, since that it is well

known that the cyanobacterial taxonomy faces several problems (Taton et al., 2006;

Komárek et al., 2014). In the phylogenetic analysis, based on the 16S rRNA gene

sequences, the heterocystous cyanobacteria formed a coherent genetic cluster

(monophyletic group), while the unicellular and filamentous nonheterocystous were

intermixed throughout the tree, in agreement with previous phylogenies (Castenholz,

2001; Gugger & Hoffmann, 2004). Our analysis also showed that some isolates

morphologically identified as same taxon were in different positions along the tree,

highlighting the difficulty to identify cyanobacteria based on morphology only and the

importance of polyphasic studies. In addition, nine 16S rRNA gene sequences had less

than 97% similarity compared to the sequences available in databases, strongly

suggesting novel cyanobacterial diversity. It is widely recognized that although the 16S

rRNA gene is the most commonly used molecular marker it lacks resolution at the

species level, thus it is often combined with other less conserved molecular markers

(e.g. internal transcribed spacer, ITS, between the 16S rRNA and 23S rRNA genes) to

achieve a higher taxonomic resolution (Rocap et al., 2002; Casamatta et al., 2006;

Komárek, 2014). Therefore, to be able to further characterize our novel isolates, the

remaining part of 16S rRNA gene, the ITS and most of the 23S rRNA gene were

sequenced and additional phylogenetic analyses were performed (Chapter III).

Together with the morphological and ultrastructural data, this analysis showed that the

isolate LEGE 07459 is related with the genera Geminocystis and Cyanobacterium

(Korelusová et al., 2009), but presents distinct features that indicate that it could be a

representative of a new genus. The isolate LEGE 07179 previously identified as Hyella

sp. clustered with members of the Pleurocapsales/Subsection II, forming a distinct

branch. Since this is the first sequence available for members of this genus it was not

possible to further infer phylogenetic relationships. The assignment of the isolate LEGE

06188 to the genus Leptolyngbya was supported by the phylogenetic analysis using

both the 16S rRNA gene only and the 16S-ITS-23S gene sequences. In contrast the

other four filamentous strains LEGE 07167, LEGE 07176, LEGE 07157 and LEGE

06111, previously assigned to L. fragilis, L. mycoidea, Phormidium/Leptolyngbya,

respectively formed a distinct cluster of filamentous cyanobacteria that probably should

be assigned to a new genus firstly isolated from the Portuguese coast.

The potential of our isolates to fix atmospheric nitrogen was evaluated by the

presence of nif genes and/or heterocysts. The presence of nifH was detected in one

third of the cyanobacteria analysed (20 out of 60). The fraction of the putative N2-fixers


103

obtained suggested that these cyanobacteria may play an important role in N2 fixation

along the Continental Portuguese coast, as it is common in the marine environment,

particularly in the intertidal zones. These zones are harsh and often nutrient-limited and

the ability to fix nitrogen gives the cyanobacteria a competitive advantage, allowing

these organisms to become the predominant N2-fixers in the majority of the marine

microbial mats (Díez et al., 2007; Severin et al., 2010). Additional work, such as

growing the strains in medium with decreasing concentrations of combined nitrogen

until depletion and the evaluation of the nitrogenase activity using the acetylene

reduction assay was initiated for some strains and will be performed for the remaining

isolates.

The putative presence of toxin producers among our isolates was initially evaluated

by assessing the presence of the genes encoding proteins involved in the production of

the cyanotoxins microcystin (mcyA, mcyE), nodularin (ndaF) and saxitoxin (sxtI). No

amplicons were obtained but this result does not exclude the presence of the genes

since the majority of the primers used were designed based on sequences from

freshwater cyanobacteria and may not be the most appropriate for marine strains.

Moreover, previous studies demonstrated that extracts of cyanobacterial strains, also

isolated from the Portuguese coast, were toxic to marine invertebrates suggesting the

presence of toxic compounds (Martins et al., 2007; Frazão et al., 2010). Recently,

crude extracts of some of our isolates were analysed by LC-MS/MS directed to detect

microcystin and the MS fragmentation pattern obtained for the isolate LEGE 07167

(Oscillatoriales) matched the pattern of microcystin-LR (data not shown). Although

microcystin is mostly produced by freshwater cyanobacteria, there are a few evidences

that marine Synechococcus strains can also produce this hepatotoxin (Carmichael & Li,

2006). Further studies should be performed to validate this preliminary result, but if this

is confirmed constitutes the first report of a microcystin-producing filamentous

cyanobacterial strain from the temperate Atlantic region and may suggest that marine

toxin-producing strains might be more common than previously believed.

In addition, the potential of our isolates to produce other secondary metabolites was

also evaluated (Chapter IV). Taken into account that the majority of the secondary

metabolites produced by cyanobacteria are PKS and NRPS, a PCR screen for the

presence of the genes encoding type I PKS and NRPS pathways, targeting the KS and

A domains respectively, was performed. The data revealed that PKS and NRPS genes

were present in the majority of the cyanobacterial isolates tested, with the PKS genes

being more ubiquitous than the NRPS genes. The higher frequency of the genes was

found in filamentous strains (about 85%), which is in agreement with previous reports

(Ehrenreich et al., 2005). Furthermore, the sequences obtained were used in an in


silico prediction of the PKS and NRPS systems. The 89 KS-domains encoding

sequences led to the identification of biosynthetic gene clusters that code for 13

different compounds. However, all the identities were low (less than 80%) indicating

that the corresponding biosynthetic gene clusters are probably not yet characterized.

The 27 sequences that coded for A-domains led to the identification of biosynthetic

gene clusters responsible for the biosynthesis of ten different compounds. Five NRPS

A-domains from five different strains had no correspondence, suggesting once again

clusters not yet characterized. It is important to stress that these results are predictions

and, although this kind of analysis can have some limitations, it is a useful resource to

quickly access the potential of an organism to produce a certain metabolite. Therefore,

the interpretation of the data should be conservative but definitely highlight the potential

of our strains to produce a diverse range of compounds. Certainly, an increase in the

number of characterized cyanobacterial biosynthetic gene clusters that leads to the

production of secondary metabolites will improve the accuracy of the bioinformatics

predictions (Micallef et al., 2014).

To understand if the genes encoding the KS- and A-domains were transcribed under

routine laboratory conditions, RT-PCR analyses were carried out in a set of selected

strains. The results obtained demonstrated that under the conditions tested both PKS

and NRPS encoding genes were transcribed in the majority of the isolates.

To complement the previous results on the potential of our isolates to produce

secondary metabolites, crude extracts of 57 of our cyanobacterial isolates were

analysed by LC-MS/MS. Their MS/MS spectra together with the MS/MS spectra of

reference cyanobacterial compounds were used to generate a network. In the obtained

network it was possible to discern different clusters and connections between nodes of

our samples and reference compounds. Our samples contained compounds related to

naopopeptin and antanapeptin, an analogue of dolastatin 16 as well as malyngamide -

type molecules. In addition, a connection between microcystin, nodularin and nodes

from different isolates was also observed, indicating that some strains might be able to

produce compounds related with hepatotoxins. One of these isolates is the filamentous

strain LEGE 07167, for which we did an additional LC-MS/MS analysis targeting

microcystin (see above) and obtained a MS fragmentation pattern that matched

microcystin-LR. It was also possible to observe clusters composed only by nodes from

our samples, highlighting that our strains can produce different/novel compounds.

Furthermore, all MS spectra were checked for major peaks and the masses were

searched against the MarinLit database. For some masses no correspondence were

found in MarinLit suggesting once more compounds not yet identified and thus

deserving further investigation.


105

Overall, only a few network connections with the reference compounds were found.

This can be explained by the fact that our isolates are from a temperate region and the

standards used were mainly isolated from tropical cyanobacteria. Moreover, it is widely

documented that environmental factors can strongly influence the production of

secondary metabolites (Shalev‐Malul et al., 2008; Sorrels et al., 2009; Esquenazi et al.,

2011). As in other studies, molecular networking proved to be a valuable tool since it

helped to identify families of compounds based on structural and spectral similarities

(Yang et al., 2013; Winnikoff et al., 2014). Altogether, these results highlighted the

metabolic potential of our strains to produce diverse compounds and it will allow the

selection of the most promising strains for further studies. This work also demonstrated

that prospecting new cyanobacterial diversity is a valid approach for the discovery of

novel compounds.

5.2. Future perspectives

The results obtained in this work provided novel and important information regarding

the diversity of cyanobacteria from the Portuguese Atlantic coast as well as defined

new questions and unveiled new paths to explore.

(i) The presence of novel cyanobacterial diversity was reported in this study. These

results should be complemented with additional phylogenetic analysis and new

genera should be proposed based on the results obtained.

(ii) Since about 30% of our isolates are putative N2-fixers it will be interesting to perform

in situ assays to verify if these strains play a key role as diazotrophs in the

intertidal ecosystems.

(iii) In an extract of one of our marine isolates was detected microcystin-LR, and there

were indication that other strains might produce hepatoxins. These results should

be validated and the sequences obtained for these strains could be used as a

template to design primers aiming at improving the molecular methods to detect

toxic marine cyanobacteria.

(iv) It would also be relevant to use the extracts of a set of selected strains to perform

standard bioassays (e.g. cancer cell line cytotoxic assays), and to isolate and

structurally characterize the metabolites produced (NMR).


(v) It will be also interesting to explore the heterologous expression of the

cyanobacterial biosynthetic gene clusters (structurally unique and/or interesting

biologically active compounds) in cyanobacterial or other bacterial hosts.

(vi) Sequence the complete genome of some of our isolates (e.g. Geminocystis-like sp.

LEGE 07459, Hyella sp. LEGE 07179 and Leptolyngbya-like sp. LEGE 07167).

General References


109

References

Adams DG & Duggan PS (2008) Cyanobacteria–bryophyte symbioses. J Exp Bot 59:

1047-1058.

Araújo R, Bárbara I, Tibaldo M, Berecibar E, Tapia PD, Pereira R, Santos R & Pinto IS

(2009) Checklist of benthic marine algae and cyanobacteria of northern Portugal.

Bot Mar 52: 24-46.

Bachmann BO & Ravel J (2009) Chapter 8. Methods for in silico prediction of microbial

polyketide and nonribosomal peptide biosynthetic pathways from DNA sequence

data. Method Enzymol 458: 181-217.

Baran R, Ivanova NN, Jose N, Garcia-Pichel F, Kyrpides NC, Gugger M & Northen TR

(2013) Functional genomics of novel secondary metabolites from diverse

cyanobacteria using untargeted metabolomics. Mar Drugs 11: 3617-3631.

Barrios-Llerena ME, Burja AM & Wright PC (2007) Genetic analysis of polyketide

synthase and peptide synthetase genes in cyanobacteria as a mining tool for

secondary metabolites. J Ind Microbiol Biotechnol 34: 443-456.

Beattie KA, Kaya K & Codd GA (2000) The cyanobacterium Nodularia PCC 7804, of

freshwater origin, produces [L-Har 2] nodularin. Phytochemistry 54: 57-61.

Bergman B, Gallon J, Rai A & Stal L (1997) N2 fixation by non-heterocystous

cyanobacteria. FEMS Microbiol Rev 19: 139-185.

Bergman B, Ran L & Adams DG (2008a) Cyanobacterial-plant Symbioses: signalling

and development. The Cyanobacteria: Molecular Biology, Genomics and Evolution

(Herrero A & Flores E, eds.), pp. 447-473. Caister Academic Press, Norfolk, UK.

Bergman B, Zheng W-W, Klint J & Ran L (2008b) On the origin of plants and relations

to contemporary cyanobacterial-plant symbioses. Plant Biotech 25: 213-220.

Bergman B, Sandh G, Lin S, Larsson J & Carpenter EJ (2013) Trichodesmium–a

widespread marine cyanobacterium with unusual nitrogen fixation properties. FEMS

Microbiol Rev 37: 286-302.

Berman-Frank I, Lundgren P & Falkowski P (2003) Nitrogen fixation and photosynthetic

oxygen evolution in cyanobacteria. Res Microbiol 154: 157-164.

Böhme H (1998) Regulation of nitrogen fixation in heterocyst-forming cyanobacteria.

Trends Plant Sci 3: 346-351.

Boison G, Steingen C, Stal LJ & Bothe H (2006) The rice field cyanobacteria Anabaena

azotica and Anabaena sp. CH1 express vanadium-dependent nitrogenase. Arch

Microbiol 186: 367-376.

Boyd MR, Gustafson KR, McMahon JB, Shoemaker RH, O'Keefe BR, Mori T,

Gulakowski RJ, Wu L, Rivera MI & Laurencot CM (1997) Discovery of cyanovirin-N,

a novel human immunodeficiency virus-inactivating protein that binds viral surface

envelope glycoprotein gp120: potential applications to microbicide development.

Antimicrob Agents Chemoter 41: 1521-1530.


Bui HT, Jansen R, Pham HT & Mundt S (2007) Carbamidocyclophanes A-E,

chlorinated paracyclophanes with cytotoxic and antibiotic activity from the

Vietnamese cyanobacterium Nostoc sp. J Nat Prod 70: 499-503.

Calteau A, Fewer DP, Latifi A, Coursin T, Laurent T, Jokela J, Kerfeld CA, Sivonen K,

Piel J & Gugger M (2014) Phylum-wide comparative genomics unravel the diversity

of secondary metabolism in cyanobacteria. BMC genomics 15: 977.

Cardellina JH, Marner FJ & Moore RE (1979) Malyngamide A, a novel chlorinated

metabolite of the marine cyanophyte Lyngbya majuscula. J Am Chem Soc 101: 240-

242.

Carmichael WW & Li R (2006) Cyanobacteria toxins in the Salton Sea. Saline systems

2: 5. doi: 10.1186/1746-1448-1182-1185.

Casamatta D, Vis ML & Sheath R (2003) Cryptic species in cyanobacterial systematics:

a case study of Phormidium retzii (Oscillatoriales) using RAPD molecular markers

and 16S rDNA sequence data. Aquat Bot 77: 295-309.

Casamatta DA, Gomez SR & Johansen JR (2006) Rexia erecta gen. et sp. nov. and

Capsosira lowei sp. nov., two newly described cyanobacterial taxa from the Great

Smoky Mountains National Park (USA). Hydrobiologia 561: 13-26.

Castenholz RW (2001) Phylum BX. Cyanobacteria. Oxygenic Photosynthetic Bacteria.

Bergey’s Manual of Systematic Bacteriology Vol. 1 (Boone DR, Castenholz RW &

Garrity G, eds.), pp. 473-599. Springer, New York.

Cavalier-Smith T (2006) Cell evolution and Earth history: stasis and revolution. Phil

Trans R Soc B 361: 969-1006.

Challis GL, Ravel J & Townsend CA (2000) Predictive, structure-based model of amino

acid recognition by nonribosomal peptide synthetase adenylation domains. Chem

Biol 7: 211-224.

Chang ZX, Sitachitta N, Rossi JV, Roberts MA, Flatt PM, Jia JY, Sherman DH &

Gerwick WH (2004) Biosynthetic pathway and gene cluster analysis of curacin A, an

antitubulin natural product from the tropical marine cyanobacterium Lyngbya

majuscula. J Nat Prod 67: 1356-1367.

Chen M, Schliep M, Willows RD, Cai Z-L, Neilan BA & Scheer H (2010) A red-shifted

chlorophyll. Science 329: 1318-1319.

Chen M, Li Y, Birch D & Willows RD (2012) A cyanobacterium that contains chlorophyll

f – a red - absorbing photopigment. FEBS letters 586: 3249-3254.

Cheng YQ, Coughlin JM, Lim SK & Shen B (2009) Type I polyketide synthases that

require discrete acyltransferases. Method Enzymol 459: 165-186.

Choi H, Pereira AR, Cao Z, Shuman CF, Engene N, Byrum T, Matainaho T, Murray TF,

Mangoni A & Gerwick WH (2010) The hoiamides, structurally intriguing neurotoxic

lipopeptides from Papua New Guinea marine cyanobacteria. J Nat Prod 73: 1411-

1421.

Christiansen G, Fastner J, Erhard M, Börner T & Dittmann E (2003) Microcystin

biosynthesis in Planktothrix: genes, evolution, and manipulation. J Bacteriol 185:

564-572.


111

Codd GA, Morrison LF & Metcalf JS (2005) Cyanobacterial toxins: risk management for

health protection. Toxicol Appl Pharmacol 203: 264-272.

Conroy T, Guo JT, Hunt NH & Payne RJ (2010) Total synthesis and antimalarial

activity of symplostatin 4. Org Lett 12: 5576-5579.

Costa M, Costa-Rodrigues J, Fernandes MH, Barros P, Vasconcelos V & Martins R

(2012) Marine cyanobacteria compounds with anticancer properties: A review on the

implication of apoptosis. Mar Drugs 10: 2181-2207.

Costa M, Garcia M, Costa-Rodrigues J, Costa MS, Ribeiro MJ, Fernandes MH, Barros

P, Barreiro A, Vasconcelos V & Martins R (2014) Exploring bioactive properties of

marine cyanobacteria isolated from the Portuguese Coast: High potential as a

source of anticancer compounds. Mar Drugs 12: 98-114.

D'Agostino PM, Moffitt MC & Neilan BA (2014) Current knowledge of paralytic shellfish

toxin biosynthesis, molecular detection and evolution. Toxins and biologically active

compounds from microalgae Vol. 1 (Rossini GP, ed.) pp. 251-280. CRC Press.

De Figueiredo DR, Reboleira AS, Antunes SC, Abrantes N, Azeiteiro U, Goncalves F &

Pereira MJ (2006) The effect of environmental parameters and cyanobacterial

blooms on phytoplankton dynamics of a Portuguese temperate lake. Hydrobiologia

568: 145-157.

De Philippis R & Micheletti E (2009) Heavy metal removal with exopolysaccharide-

producing cyanobacteria. Heavy Metals in the Environment (Wang LK, Chen JP,

Hung Y-T & Shammas NK, eds.), pp. 89-122. CRC Press, Boca Raton.

DeRuyter YS & Fromme P (2008) Molecular structure of the photosynthetic apparatus.

The Cyanobacteria: Molecular biology, genomics and evolution (Herrero A & Flores

E, eds.), pp. 217-269. Caister Academic Press, Norfolk, UK.

Deusch O, Landan G, Roettger M, Gruenheit N, Kowallik KV, Allen JF, Martin W &

Dagan T (2008) Genes of cyanobacterial origin in plant nuclear genomes point to a

heterocyst-forming plastid ancestor. Mol Biol Evol 25: 748-761.

Dey B, Lerner DL, Lusso P, Boyd MR, Elder JH & Berger EA (2000) Multiple antiviral

activities of cyanovirin-N: blocking of human immunodeficiency virus type 1 gp120

interaction with CD4 and coreceptor and inhibition of diverse enveloped viruses. J

Virol 74: 4562-4569.

Díez B, Bauer K & Bergman B (2007) Epilithic cyanobacterial communities of a marine

tropical beach rock (Heron Island, Great Barrier Reef): diversity and diazotrophy.

Appl Environ Microbiol 73: 3656-3668.

Díez B & Ininbergs K (2014) Ecological importance of cyanobacteria. Cyanobacteria:

An Economic Perspective (Sharma NK, Rai AK & Stal LJ, eds.), pp. 41-63. Willey

Blackwell, UK.

Dittmann E, Fewer DP & Neilan BA (2013) Cyanobacterial toxins: biosynthetic routes

and evolutionary roots. FEMS Microbiol Rev 37: 23-43.

Eady RR (1996) Structure-function relationships of alternative nitrogenases. Chem Rev

96: 3013-3030.


Edwards DJ & Gerwick WH (2004) Lyngbyatoxin biosynthesis: sequence of

biosynthetic gene cluster and identification of a novel aromatic prenyltransferase. J

Am Chem Soc 126: 11432-11433.

Edwards DJ, Marquez BL, Nogle LM, McPhail K, Goeger DE, Roberts MA & Gerwick

WH (2004) Structure and biosynthesis of the jamaicamides, new mixed polyketide-

peptide neurotoxins from the marine cyanobacterium Lyngbya majuscula. Chem Biol

11: 817-833.

Ehrenreich IM, Waterbury JB & Webb EA (2005) Distribution and diversity of natural

product genes in marine and freshwater cyanobacterial cultures and genomes. Appl

Environ Microbiol 71: 7401-7413.

Engene N, Rottacker EC, Kastovsky J, Byrum T, Choi H, Ellisman MH, Komarek J &

Gerwick WH (2012) Moorea producens gen. nov., sp nov and Moorea bouillonii

comb. nov., tropical marine cyanobacteria rich in bioactive secondary metabolites.

Int J Syst Evol Microbiol 62: 1171-1178.

Esquenazi E, Jones AC, Byrum T, Dorrestein PC & Gerwick WH (2011) Temporal

dynamics of natural product biosynthesis in marine cyanobacteria. P Natl Acad Sci

USA 108: 5226-5231.

Falcón LI, Magallón S & Castillo A (2010) Dating the cyanobacterial ancestor of the

chloroplast. ISME J 4: 777–783.

Finsinger K, Scholz I, Serrano A, Morales S, Uribe‐Lorio L, Mora M, Sittenfeld A,

Weckesser J & Hess WR (2008) Characterization of true‐branching cyanobacteria

from geothermal sites and hot springs of Costa Rica. Environ Microbiol 10: 460-473.

Frazão B, Martins R & Vasconcelos V (2010) Are known cyanotoxins involved in the

toxicity of picoplanktonic and filamentous north atlantic marine cyanobacteria? Mar

Drugs 8: 1908-1919.

Galhano V, De Figueiredo DR, Alves A, Correia A, Pereira MJ, Gomes-Laranjo J &

Peixoto F (2011) Morphological, biochemical and molecular characterization of

Anabaena, Aphanizomenon and Nostoc strains (Cyanobacteria, Nostocales)

isolated from Portuguese freshwater habitats. Hydrobiologia 663: 187-203.

Garcia-Pichel F (2008) Molecular ecology and environmental genomics of

cyanobacteria. The cyanobacteria: Molecular biology, Genomics and Evolution

(Herrero A & Flores E, eds.), pp. 59-88. Caister Academic Press, Norfolk, UK.

Geitler L (1932) Cyanophyceae. Rabenhorst´s Kryptogamenflora von Deutschland,

Österreich und der Schweiz Vol. XIV (Kolkwitz R, ed.) pp. 1 - 1196. Akademish

Verlagsgesellschaft, Leipzig. Reprinted 1971, Johnson, New York.

Gerwick WH, Coates RC, Engene N, Gerwick L, Grindberg RV, Jones AC & Sorrels

CM (2008) Giant marine cyanobacteria produce exciting potential pharmaceuticals.

Microbe 6: 277-284.

Grindberg RV, Ishoey T, Brinza D, Esquenazi E, Coates RC, Liu W-t, Gerwick L,

Dorrestein PC, Pevzner P & Lasken R (2011) Single cell genome amplification

accelerates identification of the apratoxin biosynthetic pathway from a complex

microbial assemblage. Plos One 6: e18565.


113

Gugger MF & Hoffmann L (2004) Polyphyly of true branching cyanobacteria

(Stigonematales). Int J Syst Evol Microbiol 54: 349-357.

Gutierrez M, Suyama TL, Engene N, Wingerd JS, Matainaho T & Gerwick WH (2008)

Apratoxin D, a potent cytotoxic cyclodepsipeptide from Papua New Guinea

collections of the marine cyanobacteria Lyngbya majuscula and Lyngbya sordida. J

Nat Prod 71: 1099-1103.

Han B, Gross H, Goeger DE, Mooberry SL & Gerwick WH (2006) Aurilides B and C,

cancer cell toxins from a Papua New Guinea collection of the marine

cyanobacterium Lyngbya majuscula. J Nat Prod 69: 572-575.

Haverkamp TH, Schouten D, Doeleman M, Wollenzien U, Huisman J & Stal LJ (2008)

Colorful microdiversity of Synechococcus strains (picocyanobacteria) isolated from

the Baltic Sea. ISME J 3: 397-408.

Hoiczyk E & Hansel A (2000) Cyanobacterial cell walls: news from an unusual

prokaryotic envelope. J Bacteriol 182: 1191-1199.

Huang T-C, Lin R-F, Chu M-K & Chen H-M (1999) Organization and expression of

nitrogen-fixation genes in the aerobic nitrogen-fixing unicellular cyanobacterium

Synechococcus sp. strain RF-1. Microbiology 145: 743-753.

Ishida K, Welker M, Christiansen G, Cadel-Six S, Bouchier C, Dittmann E, Hertweck C

& de Marsac NT (2009) Plasticity and evolution of aeruginosin biosynthesis in

cyanobacteria. Appl Environ Microbiol 75: 2017-2026.

Iteman I, Rippka R, de Marsac NT & Herdman M (2000) Comparison of conserved

structural and regulatory domains within divergent 16S rRNA–23S rRNA spacer

sequences of cyanobacteria. Microbiology 146: 1275-1286.

Jochimsen EM, Carmichael WW, An J, et al. (1998) Liver failure and death after

exposure to microcystins at a hemodialysis center in Brazil. New Engl J Med 338:

873-878.

Jones AC, Gu L, Sorrels CM, Sherman DH & Gerwick WH (2009) New tricks from

ancient algae: natural products biosynthesis in marine cyanobacteria. Curr Opin

Chem Biol 13: 216-223.

Jones AC, Monroe EA, Eisman EB, Gerwick L, Sherman DH & Gerwick WH (2010)

The unique mechanistic transformations involved in the biosynthesis of modular

natural products from marine cyanobacteria. Nat Prod Rep 27: 1048-1065.

Jungblut A-D & Neilan BA (2006) Molecular identification and evolution of the cyclic

peptide hepatotoxins, microcystin and nodularin, synthetase genes in three orders of

cyanobacteria. Arch Microbiol 185: 107-114.

Kaas H & Henriksen P (2000) Saxitoxins (PSP toxins) in Danish lakes. Water Res 34:

2089-2097.

Kaneko T, Sato S, Kotani H, Tanaka A, Asamizu E, Nakamura Y, Miyajima N,

Hirosawa M, Sugiura M & Sasamoto S (1996) Sequence analysis of the genome of

the unicellular cyanobacterium Synechocystis sp. strain PCC 6803. II. Sequence

determination of the entire genome and assignment of potential protein-coding

regions. DNA Res 3: 109-136.


Kao C & Levinson S (1986) Tetrodotoxin, saxitoxin, and the molecular biology of the

sodium channel. Annals of the New York Academy of Sciences (Boland B, cullinan J

& Cohn T, eds.), pp. 1-445. New York Academy of Sciences, New York.

Kehr JC, Gatte Picchi D & Dittmann E (2011) Natural product biosyntheses in

cyanobacteria: A treasure trove of unique enzymes. Beilstein J Org Chem 7: 1622-

1635.

Kellmann R, Michali TK & Neilan BA (2008a) Identification of a saxitoxin biosynthesis

gene with a history of frequent horizontal gene transfers. J Mol Evol 67: 526-538.

Kellmann R, Mihali TK, Jeon YJ, Pickford R, Pomati F & Neilan BA (2008b)

Biosynthetic intermediate analysis and functional homology reveal a saxitoxin gene

cluster in cyanobacteria. Appl Environ Microbiol 74: 4044-4053.

Knoll AH (2008) Cyanobacteria and earth history. The Cyanobacteria: Molecular

Biology, Genomics, and Evolution (Herrero A & Flores E, eds.), pp. 1-20. Caister

Academic Press, Norfolk, UK.

Komárek J & Anagnostidis K (1989) Modern approach to the classification system of

Cyanophytes 4-Nostocales. Algological Studies/Archiv für Hydrobiologie,

Supplement Volumes 247-345.

Komárek J & Anagnostidis K (1998) Cyanoprokaryota, Part 1: Chroococcales.

Süsswasserflora von Mitteleuropa, Bd 19/1 (Ettl H, Gartner G, Heynig H &

Mollenhauer D, eds.), pp. 548. Gustav Fischer: Stuttgart.

Komárek J & Anagnostidis K (2005) Cyanoprokaryota-2. Teil/2nd part: Oscillatoriales.

Süsswasserflora von Mitteleuropa 19⁄ 2 (Büdel B, Krienitz L, Gärtner G & Schagerl

M, eds.), pp. 759. Elsevier/Spektrum, Heidelberg.

Komárek J (2010) Recent changes (2008) in cyanobacteria taxonomy based on a

combination of molecular background with phenotype and ecological consequences

(genus and species concept). Hydrobiologia 639: 245-259.

Komárek J (2014) Modern classification of cyanobacteria. Cyanobacteria: An Economic

Perspective (Sharma NK, Rai aK & Stal LJ, eds.), pp. 21-39. Willey Blackwell, UK.

Komárek J, Kaštovský J, Mareš J & Johansen JR (2014) Taxonomic classification of

cyanoprokaryotes (cyanobacterial genera) 2014, using a polyphasic approach.

Preslia 86: 295-335.

Konstantinidis KT & Tiedje JM (2007) Prokaryotic taxonomy and phylogeny in the

genomic era: advancements and challenges ahead. Curr Opin Microbiol 10: 504-

509.

Korelusová J, Kaštovský J & Komárek J (2009) Heterogeneity of the cyanobacterial

genus Synechocystis and description of a new genus, Geminocystis. J Phycol 45:

928-937.

Leão PN, Costa M, Ramos V, Pereira AR, Fernandes VC, Domingues VF, Gerwick

WH, Vasconcelos VM & Martins R (2013a) Antitumor activity of hierridin B, a

cyanobacterial secondary metabolite found in both filamentous and unicellular

marine strains. Plos One 8: e69562.


115

Leão PN, Ramos V, Goncalves PB, Viana F, Lage OM, Gerwick WH & Vasconcelos

VM (2013b) Chemoecological screening reveals high bioactivity in diverse culturable

portuguese marine cyanobacteria. Mar Drugs 11: 1316-1335.

Liberton M & Pakrasi H (2008) Membrane systems in cyanobacteria. The

Cyanobacteria: Molecular Biology, Genomics and Evolution (Herrero A & Flores E,

eds.), pp. 271-287. Caister Academic Press, Norfolk, UK.

Lima FP, Queiroz N, Ribeiro PA, Hawkins SJ & Santos AM (2006) Recent changes

in the distribution of a marine gastropod, Patella rustica Linnaeus, 1758, and their

relationship to unusual climatic events. J Biogeogr 33: 812-822.

Lindblad P, Lindberg P, Oliveira P, Stensjö K & Heidorn T (2012) Design,

engineering, and construction of photosynthetic microbial cell factories for renewable

solar fuel production. Ambio 41: 163-168.

Linington RG, Clark BR, Trimble EE, Almanza A, Ureña L-D, Kyle DE & Gerwick

WH (2008) Antimalarial peptides from marine cyanobacteria: isolation and structural

elucidation of gallinamide A. J Nat Prod 72: 14-17.

Lopes VR, Antunes A, Welker M, Martins RF & Vasconcelos VM (2010)

Morphological, toxicological and molecular characterization of a benthic Nodularia

isolated from Atlantic estuarine environments. Res Microbiol 161: 9-17.

Lopes VR, Ramos V, Martins A, Sousa M, Welker M, Antunes A & Vasconcelos VM

(2012) Phylogenetic, chemical and morphological diversity of cyanobacteria from

Portuguese temperate estuaries. Mar Environ Res 73: 7-16.

Luesch H, Moore RE, Paul VJ, Mooberry SL & Corbett TH (2001) Isolation of

dolastatin 10 from the marine cyanobacterium Symploca species VP642 and total

stereochemistry and biological evaluation of its analogue symplostatin 1. J Nat Prod

64: 907-910.

Magarvey NA, Beck ZQ, Golakoti T, Ding Y, Huber U, Hemscheidt TK, Abelson D,

Moore RE & Sherman DH (2006) Biosynthetic characterization and chemoenzymatic

assembly of the cryptophycins. Potent anticancer agents from Nostoc cyanobionts.

ACS Chemical Biology 1: 766-779.

Maldener I, Summers ML & Sukenik A (2014) Cellular differentiation in filamentous

cyanobacteria. The cell biology of cyanobacteria (Flores E & Herrero A, eds.), pp. 263-

291. Caister Academic Press, Norfolk, UK.

Malloy KL, Villa FA, Engene N, Matainaho T, Gerwick L & Gerwick WH (2010)

Malyngamide 2, an oxidized lipopeptide with nitric oxide inhibiting activity from a Papua

New Guinea marine cyanobacterium. J Nat Prod 74: 95-98.

Martins A, Moreira C, Vale M, Freitas M, Regueiras A, Antunes A & Vasconcelos V

(2011) Seasonal dynamics of Microcystis spp. and their toxigenicity as assessed by

qPCR in a temperate reservoir. Mar Drugs 9: 1715-1730.

Martins J, Saker ML, Moreira C, Welker M, Fastner J & Vasconcelos VM (2009)

Peptide diversity in strains of the cyanobacterium Microcystis aeruginosa isolated from

Portuguese water supplies. Appl Microbiol Biotechnol 82: 951-961.


Martins R, Fernandez N, Beiras R & Vasconcelos V (2007) Toxicity assessment of

crude and partially purified extracts of marine Synechocystis and Synechococcus

cyanobacterial strains in marine invertebrates. Toxicon 50: 791-799.

Medema MH, Blin K, Cimermancic P, de Jager V, Zakrzewski P, Fischbach MA,

Weber T, Takano E & Breitling R (2011) antiSMASH: rapid identification, annotation

and analysis of secondary metabolite biosynthesis gene clusters in bacterial and fungal

genome sequences. Nucleic Acids Res 39: doi: 10.1093/nar/gkr1466.

Medina RA, Goeger DE, Hills P, Mooberry SL, Huang N, Romero LI, Ortega-Barria

E, Gerwick WH & McPhail KL (2008) Coibamide A, a potent antiproliferative cyclic

depsipeptide from the panamanian marine cyanobacterium Leptolyngbya sp. J Am

Chem Soc 130: 6324-6325.

Méjean A, Mazmouz R, Mann S, Calteau A, Médigue C & Ploux O (2010) The

genome sequence of the cyanobacterium Oscillatoria sp. PCC 6506 reveals several

gene clusters responsible for the biosynthesis of toxins and secondary metabolites. J

Bacteriol 192: 5264-5265.

Méjean A & Ploux O (2013) A genomic view of secondary metabolite production in

cyanobacteria. Genomics of Cyanobacteria Vol. 65 (Chauvat F, Cassier-Chauvat, C.,

ed.) pp. 189-234. Elsevier Academic Press, San Diego.

Mevers E, Liu W-T, Engene N, Mohimani H, Byrum T, Pevzner PA, Dorrestein PC,

Spadafora C & Gerwick WH (2011) Cytotoxic veraguamides, alkynyl bromide-

containing cyclic depsipeptides from the marine cyanobacterium cf. Oscillatoria

margaritifera. J Nat Prod 74: 928-936.

Micallef ML, D'Agostino PM, Al-Sinawi B, Neilan BA & Moffitt MC (2014) Exploring

cyanobacterial genomes for natural product biosynthesis pathways. Mar Genomics 21:

1-12.

Mihali TK, Kellmann R & Neilan BA (2009) Characterisation of the paralytic shellfish

toxin biosynthesis gene clusters in Anabaena circinalis AWQC131C and

Aphanizomenon sp. NH-5. BMC Biochem 10: 8.

Miller SR & Castenholz RW (2000) The evolution of thermotolerance in hot spring

cyanobacteria of the genus Synechococcus. Appl Environ Microbiol 66: 4222-4229.

Minich SS (2012) Brentuximab vedotin: a new age in the treatment of Hodgkin

lymphoma and anaplastic large cell lymphoma. Ann Pharmacother 46: 377-383.

Moffitt MC & Neilan BA (2001) On the presence of peptide synthetase and polyketide

synthase genes in the cyanobacterial genus Nodularia. FEMS Microbiol Lett 196:

207-214.

Moffitt MC & Neilan BA (2004) Characterization of the nodularin synthetase gene

cluster and proposed theory of the evolution of cyanobacterial hepatotoxins. Appl

Environ Microbiol 70: 6353-6362.

Nakamura Y, Kaneko T, Sato S, Mimuro M, Miyashita H, Tsuchiya T, Sasamoto S,

Watanabe A, Kawashima K & Kishida Y (2003) Complete genome structure of

Gloeobacter violaceus PCC 7421, a cyanobacterium that lacks thylakoids. DNA Res

10: 137-145.


117

Neilan BA, Jacobs D & Goodman AE (1995) Genetic diversity and phylogeny of toxic

cyanobacteria determined by DNA polymorphisms within the phycocyanin locus.


Nishizawa T, Ueda A, Asayama M, Fujii K, Harada K-i, Ochi K & Shirai M (2000)

Polyketide synthase gene coupled to the peptide synthetase module involved in the

biosynthesis of the cyclic heptapeptide microcystin. J Biochem 127: 779-789.

Nunnery JK, Mevers E & Gerwick WH (2010) Biologically active secondary metabolites

from marine cyanobacteria. Curr Opin Biotechnol 21: 787-793.

Ochoa de Alda JA, Esteban R, Diago ML & Houmard J (2014) The plastid ancestor

originated among one of the major cyanobacterial lineages. Nat Commun 5: 4937.

Orme-Johnson W (1992) Nitrogenase structure: where to now? Science 257: 1639-

1640.

Pereira A, Cao ZY, Murray TF & Gerwick WH (2009) Hoiamide A, a sodium channel

activator of unusual architecture from a consortium of two Papua New Guinea

cyanobacteria. Chem Biol 16: 893-906.

Pereira AR, Kale AJ, Fenley AT, Byrum T, Debonsi HM, Gilson MK, Valeriote FA,

Moore BS & Gerwick WH (2012) The carmaphycins: new proteasome inhibitors

exhibiting an α, β‐Epoxyketone warhead from a marine cyanobacterium.

Chembiochem 13: 810-817.

Pereira S, Zille A, Micheletti E, Moradas‐Ferreira P, De Philippis R & Tamagnini P

(2009) Complexity of cyanobacterial exopolysaccharides: composition, structures,

inducing factors and putative genes involved in their biosynthesis and assembly.

FEMS Microbiol Rev 33: 917-941.

Pontes MT, Aguiar R & Pires HO (2005) A nearshore wave energy atlas for Portugal. J

Offshore Mech Arct 127: 249-255.

Post AF (2006) The genus Prochlorococcus. The Prokaryotes Vol. 4 (Dworkin M,

Falkow S, Rosenberg E, Schleifer K-H & Stackebrandt E, eds.), pp. 1099-1110.

Springer, New York.

Potts M (1999) Mechanisms of desiccation tolerance in cyanobacteria. Eur J Phycol 34:

319-328.

Rajaniemi P, Hrouzek P, Kaštovská K, Willame R, Rantala A, Hoffmann L, Komárek J

& Sivonen K (2005) Phylogenetic and morphological evaluation of the genera

Anabaena, Aphanizomenon, Trichormus and Nostoc (Nostocales, Cyanobacteria).

Int J Syst Evol Microbiol 55: 11-26.

Ramaswamy AV, Sorrels CM & Gerwick WH (2007) Cloning and biochemical

characterization of the hectochlorin biosynthetic gene cluster from the marine

cyanobacterium Lyngbya majuscula. J Nat Prod 70: 1977-1986.

Ramos V, Seabra R, Brito Â, Santos A, Santos CL, Lopo M, Moradas-Ferreira P,

Vasconcelos VM & Tamagnini P (2010) Characterization of an intertidal

cyanobacterium that constitutes a separate clade together with thermophilic strains.

Eur J Phycol 45: 394-403.

Rantala-Ylinen A, Känä S, Wang H, Rouhiainen L, Wahlsten M, Rizzi E, Berg K,

Gugger M & Sivonen K (2011) Anatoxin-a synthetase gene cluster of the


cyanobacterium Anabaena sp. strain 37 and molecular methods to detect potential

producers. Appl Environ Microbiol 77: 7271-7278.

Rippka R, Deruelles J, Waterbury JB, Herdman M & Stanier RY (1979) Generic

assignments, strain histories and properties of pure cultures of cyanobacteria. J Gen

Microbiol 111: 1-61.

Rocap G, Distel DL, Waterbury JB & Chisholm SW (2002) Resolution of

Prochlorococcus and Synechococcus ecotypes by using 16S-23S ribosomal DNA

internal transcribed spacer sequences. Appl Environ Microbiol 68: 1180-1191.

Rossi F & De Philippis R (2015) Role of cyanobacterial exopolysaccharides in

phototrophic biofilms and in complex microbial mats. Life 5: 1218-1238.

Rouhiainen L, Vakkilainen T, Siemer BL, Buikema W, Haselkorn R & Sivonen K (2004)

Genes coding for hepatotoxic heptapeptides (microcystins) in the cyanobacterium

Anabaena strain 90. Appl Environ Microbiol 70: 686-692.

Runnegar M, Berndt N, Kong S-M, Lee EY & Zhang L (1995) In vivo and in vitro

binding of microcystin to protein phosphatase 1 and 2A. Biochem Biophys Res

Commun 216: 162-169.

Schopf JW (2000) The fossil record: tracing the roots of the cyanobacterial lineage.

The ecology of cyanobacteria (Whitton BA & Potts M, eds.), pp. 13-35. Springer,

Netherlands.

Seabra R, Wethey DS, Santos AM & Lima FP (2011) Side matters: microhabitat

influence on intertidal heat stress over a large geographical scale. J Exp Mar Biol

Ecol 400: 200-208.

Severin I, Acinas SG & Stal LJ (2010) Diversity of nitrogen‐fixing bacteria in

cyanobacterial mats. FEMS Microbiol Ecol 73: 514-525.

Shalev‐Malul G, Lieman‐Hurwitz J, Viner‐Mozzini Y, Sukenik A, Gaathon A, Lebendiker

M & Kaplan A (2008) An AbrB‐like protein might be involved in the regulation of

cylindrospermopsin production by Aphanizomenon ovalisporum. Environ Microbiol

10: 988-999.

Shannon P, Markiel A, Ozier O, Baliga NS, Wang JT, Ramage D, Amin N, Schwikowski

B & Ideker T (2003) Cytoscape: a software environment for integrated models of

biomolecular interaction networks. Genome Res 13: 2498-2504.

Shen B (2003) Polyketide biosynthesis beyond the type I, II and III polyketide synthase

paradigms. Curr Opin Chem Biol 7: 285-295.

Sherwood AR & Presting GG (2007) Universal primers amplify a 23S rDNA plastid

marker in eukaryotic algae and cyanobacteria. J Phycol 43: 605-608.

Shi T & Falkowski PG (2008) Genome evolution in cyanobacteria: the stable core and

the variable shell. P Natl Acad Sci USA 105: 2510-2515.

Shih PM, Wu D, Latifi A, et al. (2013) Improving the coverage of the cyanobacterial

phylum using diversity-driven genome sequencing. P Natl Acad Sci USA 110: 1053-

1058.

Shishido TK, Humisto A, Jokela J, Liu L, Wahlsten M, Tamrakar A, Fewer DP, Permi P,

Andreote AP & Fiore MF (2015) Antifungal compounds from Cyanobacteria. Mar

Drugs 13: 2124-2140.


119

Short SM & Zehr JP (2005) Quantitative analysis of nifH Genes and transcripts from

aquatic environments. Method Enzymol 397: 380-394.

Singh P, Singh SS, Elster J & Mishra AK (2013) Molecular phylogeny, population

genetics, and evolution of heterocystous cyanobacteria using nifH gene sequences.

Protoplasma 250: 751-764.

Singh RK, Tiwari SP, Rai AK & Mohapatra TM (2011) Cyanobacteria: an emerging

source for drug discovery. J Antibiot (Tokyo) 64: 401-412.

Sivonen K & Jones G (1999) Cyanobacterial toxins. Toxic cyanobacteria in water: a

guide to their public health consequences, monitoring and management Vol. 1

(Chorus I & Bartram J, eds.), pp. 41-111. WHO, Spon Press, London, UK.

Sivonen K & Börner T (2008) Bioactive compounds produced by cyanobacteria. The

cyanobacteria: Molecular Biology, Genomics and Evolution (Herrero A & Flores E,

eds.), pp. 159-197. Caister Academic Press, Norfolk, UK.

Sivonen K, Leikoski N, Fewer DP & Jokela J (2010) Cyanobactins - ribosomal cyclic

peptides produced by cyanobacteria. Appl Microbiol Biotechnol 86: 1213-1225.

Soe KM, Yokoyama A, Yokoyama J & Hara Y (2011) Morphological and genetic

diversity of the thermophilic cyanobacterium, Mastigocladus laminosus

(Stigonematales, Cyanobacteria) from Japan and Myanmar. Phycol Res 59: 135-

142.

Sorrels CM, Proteau PJ & Gerwick WH (2009) Organization, evolution, and expression

analysis of the biosynthetic gene cluster for scytonemin, a cyanobacterial UV-

absorbing pigment. Appl Environ Microbiol 75: 4861-4869.

Stachelhaus T, Mootz HD & Marahiel MA (1999) The specificity-conferring code of

adenylation domains in nonribosomal peptide synthetases. Chem Biol 6: 493-505.

Stal LJ (2000) Cyanobacterial mats and stromatolites. The ecology of cyanobacteria:

Their diversity in time and space (Whitton BA & Potts M, eds.), pp. 61-120. Springer,

Netherlands.

Stal LJ & Zehr JP (2008) Cyanobacterial nitrogen fixation in the ocean: diversity,

regulation, and ecology. The Cyanobacteria: Molecular Biology, Genomics and

Evolution (Herrero A & Flores E, eds.), pp. 423-446. Caister Academic Press,

Norfolk, UK.

Swingley WD, Blankenship RE & Raymond J (2008) Insights into cyanobacterial

evolution from comparative genomics. The cyanobacteria: Molecular Biology,

Genomics, and Evolution (Herrero A & Flores E, eds.), pp. 21-43. Caister Academic

Press, Norfolk, UK.

Tamagnini P, Leitão E, Oliveira P, Ferreira D, Pinto F, Harris DJ, Heidorn T & Lindblad

P (2007) Cyanobacterial hydrogenases: diversity, regulation and applications. FEMS

Microbiol Rev 31: 692-720.

Tan LT (2010) Filamentous tropical marine cyanobacteria: a rich source of natural

products for anticancer drug discovery. J Appl Phycol 22: 659-676.

Tan LT (2013a) Pharmaceutical agents from filamentous marine cyanobacteria. Drug

Discov Today 18: 863-871.


Tan LT (2013b) Marine cyanobacteria: A prolific source of bioactive natural products as

drug leads. Marine Microbiology: Bioactive Compounds and Biotechnological

Applications (Kim S-K, ed.) pp. 59-81. Wiley-VCH, Germany.

Taori K, Paul VJ & Luesch H (2008) Structure and activity of largazole, a potent

antiproliferative agent from the Floridian marine cyanobacterium Symploca sp. J Am

Chem Soc 130: 1806-1807.

Taton A, Grubisic S, Ertz D, et al. (2006) Polyphasic study of Antarctic cyanobacterial

strains. J Phycol 42: 1257-1270.

Teruya T, Sasaki H, Fukazawa H & Suenaga K (2009) Bisebromoamide, a potent

cytotoxic peptide from the marine cyanobacterium Lyngbya sp.: isolation,

stereostructure, and biological activity. Org Lett 11: 5062-5065.

Thiel T (1993) Characterization of genes for an alternative nitrogenase in the

cyanobacterium Anabaena variabilis. J Bacteriol 175: 6276-6286.

Tillett D, Dittmann E, Erhard M, von Döhren H, Börner T & Neilan BA (2000) Structural

organization of microcystin biosynthesis in Microcystis aeruginosa PCC 7806: an

integrated peptide - polyketide synthetase system. Chem Biol 7: 753-764.

Tomitani A, Knoll AH, Cavanaugh CM & Ohno T (2006) The evolutionary diversification

of cyanobacteria: molecular–phylogenetic and paleontological perspectives. P Natl

Acad Sci USA 103: 5442-5447.

Valério E, Chambel L, Paulino S, Faria N, Pereira P & Tenreiro R (2009) Molecular

identification, typing and traceability of cyanobacteria from freshwater reservoirs.

Microbiology 155: 642-656.

Vasconcelos V (1999) Cyanobacterial toxins in Portugal: effects on aquatic animals

and risk for human health. Braz J Med Biol Res 32: 249-254.

Vestola J, Shishido TK, Jokela J, Fewer DP, Aitio O, Permi P, Wahlsten M, Wang H,

Rouhiainen L & Sivonen K (2014) Hassallidins, antifungal glycolipopeptides, are

widespread among cyanobacteria and are the end-product of a nonribosomal

pathway. P Natl Acad Sci USA 111: E1909-E1917.

Wang H, Fewer DP, Holm L, Rouhiainen L & Sivonen K (2014) Atlas of nonribosomal

peptide and polyketide biosynthetic pathways reveals common occurrence of

nonmodular enzymes. P Natl Acad Sci USA 111: 9259-9264.

Wase NV & Wright PC (2008) Systems biology of cyanobacterial secondary metabolite

production and its role in drug discovery. Expert Opin Drug Discov 3: 903-929.

Watrous J, Roach P, Alexandrov T, Heath BS, Yang JY, Kersten RD, van der Voort M,

Pogliano K, Gross H & Raaijmakers JM (2012) Mass spectral molecular networking

of living microbial colonies. P Natl Acad Sci USA 109: E1743-E1752.

Weber T (2014) In silico tools for the analysis of antibiotic biosynthetic pathways. Int J

Med Microbiol 304: 230-235.

Weber T, Blin K, Duddela S, Krug D, Kim HU, Bruccoleri R, Lee SY, Fischbach MA,

Müller R & Wohlleben W (2015) antiSMASH 3.0 - a comprehensive resource for the

genome mining of biosynthetic gene clusters. Nucleic Acids Res doi:

10.1093/nar/gkv1437.


121

Whitton BA & Potts M (2000) Introduction to the Cyanobacteria. The ecology of

cyanobacteria: their diversity in time and space (Whitton BA & Potts M, eds.), pp. 1-

11. Springer, Netherlands.

Wiegand C & Pflugmacher S (2005) Ecotoxicological effects of selected cyanobacterial

secondary metabolites: a short review. Toxicol Appl Pharmacol 203: 201-218.

Winnikoff JR, Glukhov E, Watrous J, Dorrestein PC & Gerwick WH (2014) Quantitative

molecular networking to profile marine cyanobacterial metabolomes. J Antibiot 67:

105-112.

Wrasidlo W, Mielgo A, Torres VA, Barbero S, Stoletov K, Suyama TL, Klemke RL,

Gerwick WH, Carson DA & Stupack DG (2008) The marine lipopeptide

somocystinamide A triggers apoptosis via caspase 8. P Natl Acad Sci USA 105:

2313-2318.

Yang JY, Sanchez LM, Rath CM, Liu X, Boudreau PD, Bruns N, Glukhov E, Wodtke A,

de Felicio R & Fenner A (2013) Molecular networking as a dereplication strategy. J

Nat Prod 76: 1686-1699.

Zehr JP, Mellon MT & Zani S (1998) New nitrogen-fixing microorganisms detected in

oligotrophic oceans by amplification of nitrogenase (nifH) genes. Appl Environ

Microbiol 64: 3444-3450.

Zehr JP (2011) Nitrogen fixation by marine cyanobacteria. Trends Microbiol 19: 162-

173.

Ziemert N, Ishida K, Weiz A, Hertweck C & Dittmann E (2010) Exploiting the natural

diversity of microviridin gene clusters for discovery of novel tricyclic depsipeptides.


Ziemert N, Podell S, Penn K, Badger JH, Allen E & Jensen PR (2012) The Natural

Product Domain Seeker NaPDoS: A phylogeny based bioinformatic tool to classify

secondary metabolite gene diversity. Plos One 7:e34064.

characterization of cyanobacteria isolated from the

Documents