Genomic and Morphological Diversity of Marine Planktonic Diatom

Genomic and morphological diversity of marine planktonic diatom-diazotroph associations — a continuum of integration and diversification through geological time Andrea Caputo Academic dissertation for the Degree of Doctor of Philosophy in Marine Ecology at Stockholm University to be publicly defended on Friday 22 February 2019 at 09.30 in P216. NPQ-huset, Svante Arrhenius väg 20.

Abstract Symbioses between eukaryotes and nitrogen (N2)-fixing cyanobacteria (or diazotrophs) are quite common in the plankton community. A few genera of diatoms (Bacillariophyceae) such as Rhizosolenia, Hemiaulus and Chaetoceros are well known to form symbioses with the heterocystous diazotrophic cyanobacteria Richelia intracellularis and Calothrix rhizosoleniae. The latter are also called diatom-diazotroph associations, or DDAs. Up to now, the prokaryotic partners have been morphologically and genetically characterized, and the phylogenetic reconstruction of the well conserved nifH gene (encodes for the nitrogenase enzyme) placed the symbionts in 3 clusters based on their host-specificity, i.e. het-1 (Rhizosolenia-R. intracellularis), het-2 (Hemiaulus-R. intracellularis), and het-3 (Chaetoceros-C- rhizosoleniae). Conversely, the diatom-hosts, major representative of the phytoplankton community and crucial contributors to the carbon (C) biogeochemical cycle, have been understudied. The first aim of this thesis was to genetically and morphologically characterize the diatom-hosts, and to reconstruct the evolutionary background of the partnerships and the symbiont integration in the host. The molecular-clock analysis reconstruction showed the ancient appearance of the DDAs, and the traits characterizing the ancestors. In addition, diatom- hosts bearing internal symbionts (with more eroded draft genomes) appeared earlier than diatom-hosts with external symbionts. Finally a blast survey highlighted a broader distribution of the DDAs than expected. The second aim of this thesis was to compare genetic and physiological characteristics of the DDAs symbionts with the other eukaryote-diazotroph symbiosis, i.e. prymnesiophyte-UCYN-A (or Candidatus Atelocyanobacterium thalassa). The genome comparison highlighted more genes for transporters in het-3 (external symbiont) and in the UCYN-A based symbiosis, suggesting that symbiont location might be relevant also for metabolic exchanges and interactions with the host and/or environment. Moreover, a summary of methodological biases that brought to an underestimation of the DDAs is reported. The third aim of this thesis was to determine the distribution of the DDAs in the South Pacific Ocean using a quantitative polymerase chain reaction (qPCR) approach and to outline the environmental drivers of such distribution. Among the het- groups, het-1 was the most abundant/detected and co-occurred with the other 2 symbiotic strains, all responding similarly to the influence of abiotic factors, such as temperature and salinity (positive and negative correlation, respectively). Globally, Trichodesmium dominated the qPCR detections, followed by UCYN-B. UCYN-A phylotypes (A-1, A-2) were detected without their proposed hosts, for which new oligonucleotides were designed. The latter suggested a facultative symbiosis. Finally, microscopy observations of the het-groups highlighted a discrepancy with the qPCR counts (i.e. the former were several order of magnitudes lower), leading to the idea of developing a new approach to quantify the DDAs. The fourth aim of this thesis was to develop highly specific in situ hybridization assays (CARD-FISH) to determine the presence of alternative life-stages and/or free-living partners. The new assays were applied to samples collected in the South China Sea and compared with abundance estimates from qPCR assays for the 3 symbiotic strains. Free-living cells were indeed detected along the transect, mainly at deeper depths. Free-living symbionts had two morphotypes: trichomes and single-cells. The latter were interpreted as temporary life-stages. Consistent co-occurrence of the 3 het-groups was also found in the SCS and application of a SEM model predicted positive interactions between the het groups. We interpreted the positive interaction as absence of intra-specific competition, and consistent with the previous study, temperature and salinity were predicted as major drivers of the DDAs distribution.

Keywords: phytoplankton, diatoms, cyanobacteria, diazotrophs, symbiosis, evolution, phylogenetics, confocal microscopy, qPCR, CARD-FISH, tropics, sub-tropics.

Stockholm 2019 http://urn.kb.se/resolve?urn=urn:nbn:se:su:diva-163027

ISBN 978-91-7797-556-4 ISBN 978-91-7797-557-1

Department of Ecology, Environment and Plant Sciences

Stockholm University, 106 91 Stockholm

GENOMIC AND MORPHOLOGICAL DIVERSITY OF MARINE PLANKTONIC DIATOM-DIAZOTROPH ASSOCIATIONS

Andrea Caputo

Genomic and morphological diversity of marine planktonic diatom-diazotroph associations a continuum of integration and diversification through geological time

Andrea Caputo ©Andrea Caputo, Stockholm University 2019

ISBN print 978-91-7797-556-4 ISBN PDF 978-91-7797-557-1

Cover image: from the left, 3-D Imaris reconstrucion of two chains of H. hauckii (in green) and R. intracellularis in yellow; R. clevei (purple) in symbiosis with two trichomes of R. intracellularis (in light green) Note. this picture is from paper I; asymbiotic R. intracellularis (red and green); C. rhizosoleniae culture (in purple). Pictures by A. Caputo.

Printed in Sweden by Universitetsservice US-AB, Stockholm 2019

...alla mia famiglia.

Sammanfattning

Symbioser mellan eukaryoter och di-kvävefixerande (N2) cyanobakterier (eller diazotrofer) är vanligt förekommande i planktonsamhällen. Några få genus av kiselalger (Bacillariophyceae) tillhörande Rhizosolenia, Hemiaulus och Chaetoceros ingår i välkända symbioser med heterocysta, diazotrofa cyanobakterier av arterna Richelia intracellularis och Calothrix rhizosoleniae. Även kallade ”diatom-diazotroph associations” eller DDAs. Fram till nu har den prokaryota partnern blivit morfologiskt och genetiskt karaktäriserad och genom fylogenetisk rekonstruktion av den mycket konserverade nifH genen (som kodar för enzymet nitrogenas) har symbionterna grupperats i tre kluster baserat på deras respektive värdcell, het- 1 (Rhizosolenia-R. intracellularis), het-2 (Hemiaulus-R. intracellularis), och het-3 (Chaetoceros-C. rhizosoleniae). Kiselalgs-värdcellerna å andra sidan, som utgör en stor del av fytoplanktonsamhället och är viktiga för kolets (C) biogeokemiska cykel, är understuderade. Avhandlingens första mål var att genetiskt och morfologiskt karaktärisera kiselalgs-värdcellen och att återskapa den evolutionära bakgrunden till partnerskapet och symbiontens integrering i värdcellen. Analys av den återskapade molekylära klockan visade den uråldriga uppkomsten av DDAs och de egenskaper som karaktäriserar deras stamfäder. Dessutom uppkom kiselalgs-värdceller med interna symbionter (med mer eroderade genom) tidigare än kiselalgs-värdceller med externa symbionter. Slutligen belyste en kartläggning med verktyget BLAST en bredare distribution av DDAs än förväntat. Avhandlingens andra mål var att jämföra de genetiska och fysiologiska karaktärerna hos DDA symbionterna med andra eukaryot-diazotrof symbioser, t.ex. prymnesiophyceae-UCYN-A (eller Candidatus Atelocyanobacterium thalassa). Jämförelsen av genomen belyste att het-3 (extern symbiont) och UCYN-A hade fler gener som kodade för transportproteiner, vilket tyder på att symbiontens fysiska placering i symbiosen är relevant för metaboliskt utbyte samt interaktioner med dess värdcell och/eller miljö. Dessutom visade en summering av provtagningsmetodiken en skevhet som lett till en underskattninga av DDAs. Avhandlingens tredje mål var att med hjälp av ”quantitative polymerase chain reaction” (qPCR) fastställa distributionen av DDAs i södra Stilla Havet och ge en översikt av de miljöfaktorer som gav upphov till distributionen. Inom het-gruppen var het-1 mest abundant, och även den mest detekterade, men den förekom tillsammans med de andra två symbiotiska stammarna. Alla tre stammarna responderade likvärdigt på abiotiska miljöfaktorer såsom temperatur och salinitet (positiv respektive negativ korrelation). Ur ett globalt perspektiv var det Trichodesmium som dominerade qPCR detektionerna, följt av UCYN-B. UCYN-A fylotyper (A-1, A-2) detekterades utan deras föreslagna värdceller, för vilka nya oligonukleotider konstruerades. Den senare föreslog en fakultativ symbios. Slutligen belyste mikroskopiering en skillnad mellan antalet observationer av het-gruppen i mikroskop och antalet detekterade med qPCR (det förstnämnda var flera storleksordningar mindre) vilket ledde till idén att utveckla ett nytt sätt att kvantifiera DDAs. Avhandlingens fjärde mål var att utveckla särskilt specifika in situ hybridiseringsmetoder (CARD-FISH) för att verifiera existensen av alternativa livsstadier och/eller frilevande partners. Den nya metoden användes till prover från Sydkinesiska Havet och jämfördes sedan med den uppskattade abundansen från qPCR av de tre symbiotiska stammarna. Frilevande celler detekterades längs provtaningstransektet, främst på djupet. Frilevande symbionter hade två morfer: trikomer och encelliga. Det sistnämnda tolkades som ett temporärt livsstadium. Konsekvent sam- förekomst av 3-het-grupperna fanns också i SCS och applicering av en SEM- modell förutsagde positiva interaktioner mellan het-grupperna. Vi tolkade den positiva interaktionen som frånvaro av intra-specifik konkurrens, och i överensstämmelse med föregående studie, var temperatur och salthalt förutspådda som huvudförare för DDA-fördelningen.

Nyckelord: fytoplankton, kiselalger, cyanobakterier, diazotrofer, symbios, evolution, fylogenetik, konfokalmikroskopi, qPCR, CARD-FISH, tropisk, sub-tropisk TABLE OF CONTENTS

LIST OF PUBLICATIONS 5 ABBREVIATIONS 7 INTRODUCTION 9 Nitrogen fixation in the oceans 10

Marine diazotrophic cyanobacteria 11

Diatom Diazotroph Associations – DDAs 16

N2 fixation rates in the oceans 18

Factors influencing marine N2 fixation 18

DDAs evolutionary records 21

Microscopy observations and symbiont location 22

DDA symbiont diversity and sequenced genomes 25

Diatom-hosts diversity and molecular markers 25

DDAs abundance and ecological distribution 28

Other eukaryote-diazotroph associations 30 AIMS OF THE THESIS 32

COMMENTS ON METHODOLOGY 33

Sampling areas and conditions 33

Cultures collections 34

Microscopy 36

In situ hybridization 38

Molecular analyses 40

Quantitative polymerase chain reaction (qPCR) 42

Piecewise Structural Equation Model (SEM) 42 Phylogenetic analysis 44

RESULTS AND DISCUSSION 45 Re-evaluation of symbiont location and morphology using advanced microscopy 45 In situ hybridization determines DDAs distribution and host-symbiont specificity 47

Molecular-clocks outline the DDAs appearance and the ancestors’ traits 50

Sequence-based survey broaden the DDAs ecological distribution 54

CONCLUSIONS 56

FUTURE PERSPECTIVES 57

ACKNOWLEDGMENTS 64

REFERENCES 67

List of publications This thesis is based on the following papers:

I. Caputo A., Nylander J.A.A., and Foster R.A. (2019) The genetic diversity and evolution of diatom-diazotroph associations highlights traits favoring symbiont integration. FEMS Microbiology Letters 365: 1–11. doi:10.1093/femsle/fny297.

II. Caputo A., Stenegren M., Pernice M.C., and Foster R.A. (2018) A short comparison of two marine diazotrophic symbioses highlights an un-quantified disparity. Frontiers in Marine Science 5 (2): 1–8. doi: 10.3389/fmars.2018.00002

III. Stenegren M., Caputo A., Berg C., Bonnet S., and Foster R.A. (2018) Distribution and drivers of symbiotic and free-living diazotrophic cyanobacteria in the Western Tropical South Pacific. Biogeosciences 15 (5): 1559–1578. doi: 10.5194/bg-15-1559-2018

IV. Caputo A., Steiger M., Pernice M.C., Stenegren M., Montoya J.P., Subramaniam A., and Foster R.A. Asymbiotic host and symbionts in a widely distributed N2 fixing planktonic symbiosis identified by new CARD-FISH assays. Manuscript.

My contributions to the papers: (I) – Experimental design, sample collection in the South Pacific, confocal microscopy, PCR, phylogenetic analysis with assistance, data processing, major part in writing and revisions. (II) – Organized the joint co-authored perspective paper, contributed to data collection, presentation and writing. (III) – Sample collections, DNA extraction and qPCR analysis at sea, microscopy analysis, contributed to the writing. (IV) – Experimental design, double CARD-FISH optimization, oligonucleotide designs, microscopy analysis and execution, data processing, lead in writing.

Additional peer-reviewed articles to which the author of the thesis contributed during his PhD studies:

(i). Pennesi C., Caputo A., Lobban C., Poulin M., and Totti C. (2017) Morphological discoveries in the genus Diploneis (Bacillariophyceae) from the tropical west Pacific, including the description of new taxa. Diatom Research 32 (2): 195–228. doi: 10.1080/0269249X.2017.1343752

(ii). Spungin D., Belkin N., Foster R.A., Stenegren M., Caputo A., Pujo- Pay M., Leblond N., Dupouy C., Bonnet S., and Berman-Frank I. (2018) Programmed cell death in diazotrophs and the fate of organic matter in the Western Tropical South Pacific Ocean during the OUTPACE cruise. Biogeosciences. doi: 10.5194/bg-2018-3

Abbreviations

BNF Biological Nitrogen Fixation CalSC Calothrix rhizosoleniae draft genome (associated with C. compressus) CARD- CAtalyzed Reporter Deposition FISH Fluorescence In Situ Hybridization DAPI 4′,6-diamidino-2-phenylindole DDA Diatom Diazotroph Association DIN Dissolved Inorganic Nitrogen DON Dissolved Organic Nitrogen FISH Fluorescence In Situ Hybridization GOGAT Glutamine oxoglutarate aminotransferase het-1 Richelia intracellularis associated with Rhizosolenia clevei het-2 Richelia intracellularis associated with Hemiaulus spp. het-3 Calothrix rhizosoleniae associated with Chaetoceros compressus Mbp Megabase pair Mya Million years ago

N2 Di-nitrogen

NH3 Ammonia + NH4 Ammonium nifH Nitrogenase iron protein component II - NO3 Nitrate PAR Photosynthetically active radiation PCR Polymerase Chain Reaction PFA Paraformaldehyde PON Particulate Organic Nitrogen qPCR Quantitative-PCR rbcL RuBisCO gene RintHH R. intracellularis draft genome (associated with H. haukii) RintHM R. intracellularis draft genome (associated with H. membranaceus) RintRC R. intracellularis draft genome (associated with R. clevei)

SEM Piecewise Structural Equation Model SIMS Secondary Ion Mass Spectrometry TEM Transmission Electron Microscopy TL Transmitted light UCYN Unicellular cyanobacteria

Introduction

Endosymbiotic events deeply changed life on our planet. One of the earliest endosymbioses involved the engulfment of a primitive cyanobacterium by an ancient eukaryote, giving rise to an ancestral algae and its photosynthetic organelle, the chloroplast (Cavalier-Smith, 2000). Green algae, red algae, and Glaucophytes (plus land plants) still preserve the original chloroplast (phylum Archeplastida), whereas modern microalgal phyla undergone various endosymbiotic events, resulting into multiple number of chloroplasts and membranes (e.g. phylum Stramenopiles) (Keeling, 2013). A few genera of marine and freshwater diatoms (phylum Stramenopiles, class

Bacillariophyceae) form symbiosis with di-nitrogen (N2) fixing cyanobacteria.

The N2 reduction to ammonia provide an available form of nitrogen (N) to their host, sustaining their metabolism. Early evidences of diatom fossils from the Late Cretaceous (100-66 Mya) coinciding with isotopic signatures of N2- fixing cyanobacteria, suggested the ancient nature of these symbioses (Sachs and Repeta, 1999; Kashiyama et al., 2008; Bauersachs et al., 2010). Given the evolutionary records, the internal location of the symbiont (in most diatom- hosts), and their function as N-source, the symbiont might be evolving into a nitroplast. Hence, the intricate nature of these symbioses and their fundamental role in the past and present carbon (C) and N biogeochemical cycles make them interesting to study.

This PhD thesis focuses on the planktonic association between diatoms and N2 fixing cyanobacteria, also known as Diatom Diazotrophs Associations (DDAs), and seeks to acquire genetic, phylogenetic, evolutionary, morphological and ecological knowledge on these symbioses, re-evaluating previous approaches and proposing alternative methods.

Nitrogen fixation in the oceans About half of the photosynthesis on Earth is performed by phytoplankton in the oceans (Karl et al., 2012). Vast regions of the surface oceans are nutrient depleted, thus the photosynthetic C sequestration from the atmosphere to deep waters, is influenced by the load of new N to the ecosystem. In the open ocean, inputs of N to the euphotic zone can either diffuse from deep waters as nitrate - (NO3 ; McCarthy and Carpenter, 1983; Chavez and Toggweiler, 1995) or, alternatively, through the biological N2 fixation (BNF) (Karl et al., 1997; Capone et al., 2005) (Fig. 1). BNF has a high energy demand, considering the

16 ATP required to reduce inert atmospheric di-nitrogen (N2; N≡N) into a biologically available form, ammonia (NH3):

+ - N2 + 8H + 8e + 16ATP > 2NH3 + H2 + 16ADP + 16 Pi

Fig. 1. Carbon (C) and Nitrogen (N) pumps in the open oceans. With permission from Sohm et al., 2011.

Tropical and subtropical oceanic waters, often characterized by oligotrophic conditions, contribute to most of the N2 fixation in the world oceans; hence open oceans have a major role in the global N biogeochemical cycle (Karl et al., 2002, 2012; Montoya et al., 2004; Foster et al., 2013). The enzyme that 10 catalyzes the reduction of atmospheric N2 to ammonia is the nitrogenase, composed of two subunits, both with a high iron (Fe) requirement (38 atoms of Fe in total), and encoded by a suite of nif genes. The iron molybdenom (FeMo) subunit is encoded by nifDK, and the Fe subunit is encoded by nifH

(Kim and Rees, 1992). The former is highly conserved among N2 fixing cyanobacteria, and its phylogeny mostly congruent with the 16S rRNA reconstruction (Zehr et al., 2003). Indeed, nifH has become an essential marker for phylogenetic, taxonomical and quantification analyses of diazotrophs.

Marine diazotrophic cyanobacteria

The organisms capable of N2 fixation are also known as diazotrophs (“di”, two; “azo”, nitrogen; “troph”, pertaining to nourishment). In the open ocean, much of the BNF is attributed to a diverse group of cyanobacterial lineages (Fig. 2; Table 1): filamentous non-heterocystous Trichodesmium spp., heterocystous symbiotic forms (Richelia intracellularis and Calothrix rhizosoleniae) associated with diatoms, and a few groups of unicellular types: UCYN-A, UCYN-B and UCYN-C.

Fig. 2. Marine diazotrophic cyanobacteria lineages and their implications in the biogeochemical cycles. From the left: Crocosphaera watsonii (UCYN-B), Trichodesmium spp., R. intracellularis and C. rhizosoleniae, and UCYN-A. The arrows represent the gas exchanges between atmosphere and ocean for each symbioses (yellow=exchanges relative to the cyanobacteria; grey=exchanges relative to the eukaryotic hosts). Note. In red the hosts´ chloroplasts. Organisms are not to scale, for example UCYN-A is estimated as 1-2 µm and its host 15-25 µm. Modified from Thompson and Zehr, 2013.

Table 1. Summary of the photosynthetic marine diazotrophic cyanobacteria and their main physiological and genetic traits. Note. Due to limited information the other UCYN-A (A3-A6) lineages have not been included; Trichodesmium spp. has been categorized as “free-living” although the bacterial community associated with it (Ruoco et al., 2016). Note. + stands for presence; – for absence; f=filamentous; u=unicellular. Modified from Zehr, 2011.

- -

Reference

Hilton et et Hilton al., 2013 et Hilton al., 2013 Walworth et 2015 al., al., et Tripp 2010 et Bombar al., 2014 et Bench al., 2011

, 7.8 Mb, , 7.8

genome, 3.2 3.2 genome,

8501,

Genome

HH01, draft HH01, coding 56% GC, Mb, 34% 6.0 genome, draft SC01, coding 76% GC, Mb, 39% T. erythraeum coding 64% 34% GC, 81% GC, 31% 1.44 Mb, coding 79.3% GC, 31% 1.47 Mb, coding watsonii C. 79% GC, 6.2 Mb, 37% coding reported. Not

Day Day Day Day Day

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Time of Time

otic

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spp.

A1 A2 B C

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fixing cyanobacteria

N intracellularis Richelia rhizosoleniae Calothrix Trichodesmium UCYN UCYN UCYN UCYN

Based on the phylogenetic reconstruction of partial fragments of 16S rRNA, nifH and hetR genes (the latter encodes for heterocyst and akinete differentiation), R. intracellularis and C. rhizosoleniae clustered in a well- defined group, also known as “het-group”, further branching depended on the associated diatom-host, i.e. het-1, for R. intracellularis associated with R clevei, het-2A for R. intracellularis associated with H. haukii, het-2B for R. intracellularis associated with H. membranaceus and het-3 for C. rhizosoleniae associated with C. compressus (Fig. 3; Janson et al., 1999; Foster and Zehr, 2006). The driver of such specificity and if the host has a selective process remains unknown. Three additional partial gene markers were sequenced only for the het-3 symbiont, i.e. rpnB, rbcL, narB (encoding for ribonuclease, large subunit RuBisCO, and Nitrate Reductase, respectively; Foster et al., 2010).

Recently 6 sub-lineages of UCYN-A have been described based on nifH sequences, 2 of which (UCYN-A1 and A2) are symbiotic with single celled eukaryotic microalgae (Fig. 3; Zehr et al., 2008; Tripp et al., 2010; Thompson et al., 2012; Hagino et al., 2013; Bombar et al., 2014; Farnelid et al., 2016; Turko-Kubo et al., 2017). The closest cultured representative of UCYN-B is Crocosphaera watsonii, which lives singly, in colonies, and also reported in symbioses with diatoms (Carpenter and Janson, 2000; Zehr et al., 2007; Webb et al., 2009; Foster et al., 2011, 2013). Less is known about UCYN-C other than it often co-occurs with the other cyanobacterial diazotrophs and is phylogenetically closely related to the unicellular diazotroph Cyanothece ATCC51142 (Langlois et al., 2005; Foster et al., 2007; Turk-Kubo et al., 2015).

The pelagic diazotrophic domain also includes non-cyanobacterial N2 fixers (NCDs), mostly represented by heterotrophic bacteria (e.g. Alpha- and Gammaproteobacteria), and in less extent Archaea and anaerobic bacteria (Riemann et al., 2010). Although their contribution to the BNF is on average lower than 1 nmol N L-1 d-1 (Moisander et al., 2017), nifH studies have estimated the NCDs abundance such as up to 107 nifH copies L-1 (Halm et al., 2012; Moisander et al., 2012; Farnelid et al., 2013; Loescher et al., 2014; Bentzon-Tilia et al., 2015; Shiozaki et al., 2014; Bombar et al., 2016). Hence, a substantial role in the oceanic C and N budgets is outlined, and potentially comparable to the cyanobacterial contribution.

Fig. 3. Diazotrophic cyanobacteria nifH nucleotide phylogenetic reconstruction (45 strains; 359 bp) highlights R. intracellularis and C. rhizosoleniae clustering according to their host specificity. The three main clusters are highlighted (het-1=red; het- 2=green; het-3=blue). Note. The symbiotic unicellular UCYNA-1, -2, are highlighted in yellow. The numbers on the nodes represent the bootstrap values inferred with neighbor-joining iteration and Jukes-Cantor correction. Outgroup: Trichodesmium erythraeum (IMS 101). Phylogenetic reconstruction by Andrea Caputo.

Diatom Diazotroph Associations – DDAs The first who coined the term “symbiosis” (derived from Greek “sýn-”, together; “bíōsis”, living) was the German biologist Albert Bernhard Frank in late 1877, describing crustose lichens as the tight association between a fungus and a blue-green algae (cyanobacteria). Frank defined symbiosis as “where two different species live on or in one another” and suggested the term “mycorrhiza” for the mutualistic partnership between the fungi and the roots of the trees (Frank, 1877). Common also to terrestrial plants are nitrogen- fixing symbioses, for example between ferns and diazotrophic cyanobacteria, e.g. Azolla-Nostoc (Papaefthimiou et al., 2008), or between actinorhizal plants and N2 fixing Gram positive soil actinobacteria, e.g. Alnus-Frankia (Benson and Silvester, 1993). In the plankton, partnerships are commonly observed, especially in oligotrophic waters, where nutrient limitation drives different organisms to form symbioses (Decelle et al., 2015). Often cyanobacteria have been observed living with diverse eukaryotic hosts, such as radiolarians, tintinnids, dinoflagellates, silicoflagellates, haptophytes, and diatoms (Carpenter and Foster, 2002; Foster et al., 2006). The filamentous heterocystous diazotrophic cyanobacteria R. intracellularis and C. rhizosoleniae have been observed in symbioses with different genera of centric diatoms: Rhizosolenia, Hemiaulus, Chaetoceros, and more rarely with Guinardia (Gómez et al., 2005; Foster and Zehr, 2006; Foster and O’Mullan, 2008). Collectively the latter are referred to as diatom-diazotroph associations, DDAs (Fig. 4).

Fig. 4. Confocal images of three open-ocean DDAs. (A) Valve apex of symbiotic Rhizosolenia with two trichomes of Richelia (yellow) and the chloroplast of the diatom (dark green), Note the silica frustule is visible; (B) a chain of 4 Hemiaulus cells with 6 trichomes of Richelia (yellow) and the chloroplasts of the diatoms (in green). Note the larger diameter for the terminal heterocyst over the vegetative cells of Richelia; (C) a chain of Chaetoceros with 8 filaments of Calothrix (yellow; 1 is less visible) and a contaminating Trichodesmium sp. filament (green). Zeiss LSM 780 equipped with 488 and 561 laser lines [note. (A) and (C) were imaged with additional Transmitted Light]. Scale bars 10 μm. Legend: c (chloroplast), h (heterocyst), v (vegetative cell). Photos by Andrea Caputo. 16

Within the definition of DDAs are included symbiosis between diatoms and unicellular diazotrophic cyanobacteria. For example, in few occasions the marine centric diatom Climacodium frauenfeldianum has been observed in symbiosis with unicellular cyanobacteria morphologically and phylogenetically (16S rRNA sequence) similar to C. watsonii´s (Carpenter and Janson, 2000). Moving to the freshwaters, diatoms belonging to the family Rhopalodiaceae, i.e. Rhopalodia gibba (Ehrenberg) Otto Müller (Fig. 5; Prechtl et al., 2004; Kneip et al., 2008; Nakayama et al., 2011), R. gibberula (Ehrenberg) Otto Müller (Nakayama and Inagaki, 2017), Epitemia sorex Kützing (Nakayama et al., 2011), E. turgida (Ehrenberg) Kützing (Nakayama et al., 2014), host intracellular “spheroid bodies” of cyanobacterial origin. The genome sequencing of the “spheroid bodies” in E. turgida (EtSB) and R. gibberula (RgSB) revealed the presence of nitrogenase genes and, at the same time, the absence of photosynthetic genes (Nakayama et al., 2014; Nakayama and Inagaki, 2017). These evidences highlighted a streamlined genome probably due to the internal location and a strict host-dependency. Hence, the genome reduction of the spheroid bodies could represent an evolutionary process leading to the origin of a “nitroplast”, i.e. an N2-fixing organelle for its host diatom.

Fig. 5. Confocal micrograph of R. gibba (chloroplast in red; frustule in gray) and the internal spheroid bodies (in green). Scale bar: 10 µm. With permission from Trapp et al., 2012.

N2 fixation rates in the oceans.

In open oceans, DDAs contribute significantly to the N and C biogeochemical cycles (Venrick et al., 1974; Mague et al., 1974; Carpenter et al., 1999, 2004; Karl et al., 2002, 2012; Montoya et al., 2004; Foster et al., 2013). Field based measures of single-cell N2 fixation rates for R. intracellularis associated with H. hauckii has been estimated in the range of 1.15–50.4 fmol N cell−1 h−1

(Foster et al., 2011). The latter are in fact comparable to the single-cell N2 fixation rates of other widespread diazotrophic cyanobacteria. For example, the single-cell N2 fixation rates of the filamentous Trichodesmium have been estimated between 0.92–28.7 fmol N cell−1 h−1 (Orcutt et al., 2001; Luo et al., 2012; Martinez-Perez et al., 2016), which are also comparable to the single- cell UCYN-A1 associated with the prymnesiophyte Braarudosphaera bigelowii (0.5–9.16 fmol N cell−1 h−1; Martinez-Perez et al., 2016), and Crocosphaera watsonii (or UCYN-B) estimated between 2.10×10-3–2.31 fmol N cell−1 h−1 (Foster et al., 2013; Krupke et al., 2013). Additionally, due to their high sinking rates, the DDAs are responsible for the rapid transfer of diazotroph-derived nitrogen (DDN) to the deep sea (Subramaniam et al., 2008; Karl et al. 2012). According to bio-volume and C content estimates of the symbiotic heterocystous cyanobacteria, the DDA symbionts contribute up to 6% of the total biomass of the marine diazotrophic cyanobacteria, ranging from 3.3–16 μg C m−3 (Luo et al., 2012). Nevertheless, given the major C production and export by the diatom-host genera estimated through fossil records (Kemp and Villareal, 2013), a re-evaluation of the DDAs actual contribution to the global C biogeochemical cycle in the modern oceans should be considered.

Factors influencing marine N2 fixation

Marine N2 fixation and distribution of N2 fixers are influenced by a variety environmental factors and the extent of such limitation varies depending on the diazotrophic species.

Oxygen (O2) has a big impact on N2 fixation, since it irreversibly deactivates the nitrogenase enzyme (Postgate, 1982). Diazotrophic cyanobacteria have evolved different strategies to prevent the inhibition of the nitrogenase enzyme by the O2 evolved during photosynthesis: specialized cells, temporal and/or spatial separation. R. intracellularis and C. rhizosoleniae are heterocystous cyanobacteria, hence they are characterized by a terminal heterocyst, i.e. an

N2-fixing specialized cell lacking the oxygenic Photosystem II and composed 18 of unique glycolipids and a polysaccharide layer which decreases the O2 permeability (Kumar et al., 2010). Thus, the nitrogenase can function during the day without being inhibited by the O2 evolved from the adjacent photosynthetic vegetative cells. Conversely, a temporal shift has been adopted by most unicellular photosynthetic diazotrophs (e.g. C. watsonii, or UCYN- B, Cyanothece spp.), which use the ATP generated by the PSI during the day to reduce the atmospheric N2 at night, and keeping the nitrogenase activated by respiring the photosynthetic O2 (Gallon, 1992; Sherman et al., 1998; Berman-Frank et al., 2003; Dron et al., 2012) (Table 1). An exception is represented by the unicellular UCYN-A, which lacks photosystem II (Zehr et al., 2008; Tripp et al., 2010), hence the nitrogenase does not risk deactivation by O2 evolved during photosynthesis (Bothe et al., 2010). A combination of the previous strategies has been proposed for the filamentous non- heterocystous Trichodesmium spp., which developed both a spatial and temporal separation. Although the N2 fixation still occurs during the light phase of the diel cycle, the enzyme nitrogenase is localized to a portion of the trichome called “diazocyte”, characterized by higher level of respiratory enzymes (Capone et al., 1997; Berman-Frank et al., 2001; Bergman et al., 2013).

Among other abiotic factors (e.g. salinity, PAR), temperature influences N2 fixation and the ecological distribution of the diazotrophs. Due to the fact that high water temperature decreases the O2 dissolution, non-heterocystous diazotrophic cyanobacteria should be constrained to the tropical and subtropical areas, whereas heterocystous N2-fixing cyanobacteria (e.g. Richelia/Calothrix) were thought to be favored at higher latitudes (Staal et al., 2003; Stal, 2009). Indeed, the distribution of Trichodesmium spp. is largely constrained to the 20-25 C isotherms (Capone et al., 1997; Breitbarth et al., 2007). A similar temperature-dependent distribution has been recently outlined for the unicellular diazotrophic cyanobacterium UCYN-C, which showed a better adaptation to warmer waters than the other UCYN strains (Berthelot et al., 2017). Conversely, previous studies on UCYN-A detected its presence at higher latitudes and depths (Needoba et al., 2009; Moisander et al., 2010; Farnelid et al., 2016; Martínez-Pérez et al., 2016), outlining a broader distribution than expected and undermining the canonical view of diazotrophy limited to the warm oligotropic gyres.

Nutrient availability, mainly iron (Fe) and phosphorous (P), is also crucial for

N2 fixation. As previously mentioned, due to high Fe requirement, the nitrogenase enzyme is directly affected by dissolved or available Fe concentrations. As a matter of fact, the North Atlantic Ocean is naturally 19 fertilized by Aeolian Saharan dust rich in Fe, which promotes diazotrophic activity, and registers among the highest rates of areal N2 fixation (Moore et al., 2009; Chappell et al., 2012; Snow et al., 2015). Nevertheless, similar rates have been estimated in the North Pacific Ocean, which is less affected by iron supply, suggesting that other drivers may limit BNF (Luo et al., 2014; Weber and Deutsch, 2014). Phosphorous, on the other hand, is fundamental for other energy rich molecules, such as ATP and NADPH, and can represent a limiting factor for the diazotrophic activity in open oceans (Dhyrman and Haley, 2006; Hynes et al., 2009). To overcome such limitation, some diazotrophic cyanobacteria developed strategies to efficiently assimilate P from the environment; for example, recent studies on R. intracellularis associated with Rhizosolenia (het-1) reported the ability of the cyanobacteria to assimilate dissolved organic phosphorous (DOP; Girault et al., 2013), favoring also the

N2 fixation in oligotrophic waters (Meyer et al., 2016). Similarly, 3- Trichodesmium can assimilate both inorganic phosphate (PO4 ; Dhyrman et al., 2002) and phosphonate compounds (Dhyrman et al., 2006); whereas low- P conditions triggers in Crocosphaera WH8501 an enzymatic cascade which 3- increases its affinity for PO4 (Dhyrman and Haley, 2006). Alternatively, diazotrophs can be simultaneously limited by Fe and P; for example, previous studies in the Pacific Ocean (Hynes et al., 2009; Chappell et al., 2012), corroborated by more recent laboratory studies, identified Trichodesmium as Fe/P co-limited (Walworth et al., 2016, 2018), suggesting that likely other diazotrophic cyanobacteria might be equally affected.

Less studied than the above mentioned nutrients is the role of vitamins and their influence on N2 fixation. Most phytoplanktonic eukaryotes, diatoms included, need vitamin B12, or cobalamin, for their growth (Haines and Guillard, 1974). Marine diazotrophic cyanobacteria (e.g. Crocosphaera) can indeed provide vitamin B12 to larger eukaryotes (e.g. diatoms; Bonnet et al., 2010), which they can use to synthesize methionine, and, likely, supply DOM to the bacteria. (Amin et al., 2012). Hence, vitamin B12 availability seems a crucial factor for the establishment of diatom-N2 fixing cyanobacteria symbiosis, especially in oligotrophic waters.

DDAs evolutionary records

In both past and modern oceans, diatoms have contributed significantly to global primary production (Nelson et al., 1995; Field et al., 1998). Moreover, the siliceous diatom cell walls (or frustules) are well preserved in the sediments, and offer a valuable record and a documented ‘glimpse’ into the past. For example, according to paleo-ecological studies on Mediterranean sapropels (i.e. sediment records rich in organic matter) and Arctic Ocean sediments, diatom derived primary production has been fundamental in the late Cretaceous and Paleocene periods (53-65 Mya), as well as more recently, in the late Quaternary (approx. 1 Mya) (Kemp et al., 1999; Bauersachs et al., 2010; Davies and Kemp, 2016). The Cretaceous era has been characterized by an intense climate change, which severely affected the oceans and its living forms. The increase of greenhouse gases (mostly carbon dioxide, CO2 and methane, CH4) caused an increase of both atmospheric and oceanic temperatures (up to 42 ºC), resulting in vast anoxic areas, high sea level, an undefined pycnocline, and mesoscale water circulation (Hay, 2008). During the Cretaceous the composition of sapropel laminate sediments was largely characterized by alternating mats of genera Rhizosolenia sp. and Hemiaulus spp. (Kemp et al., 1999; Bauersachs et al., 2010; Davies and Kemp, 2016), which are 2 of the 3 DDA host lineages (Fig. 6).

Fig. 6. Scanning electron microscope micrographs of (A) Rhizosolenia sp., (B) Chaetoceros spp. resting spores, and (C) Hemiaulus cf. gleseri chain. Parallel light microscopy images showing the symbiosis between (D) R. clevei and R. intracellularis, (E) C. compressus and C. rhizosoleniae, (F) Hemiaulus sp. and R. intracellularis. Note. A, B, and C are with permission from Davies and Kemp, 2016; D, E, and F are photos by Andrea Caputo. Scale bars: A, D, E (10 µm), B (2 µm), and C (20 µm), F (5 µm).

The layers enriched with Rhizosolenia and Hemiaulus microfossils were also found to possess isotopic signatures (e.g. δ15N range -6.6 -3.9%) indicative of intense N2 fixation (Sachs and Repeta, 1999; Kashiyama et al., 2008). Additionally, the unique glycolipid composition characterizing the endosymbiotic heterocystous cyanobacteria has been recently characterized for 2 of the 3 het-groups (i.e. het-1 and 2; Bauersachs et al., 2010; Schouthen et al., 2013; Bale et al., 2018). Hence, the co-occurrence of fossilized glycolipids produced by N2 fixing cyanobacteria and the conspicuous presence of two genera of symbiotic diatoms from the same geological era suggested an early occurrence of the DDAs.

Microscopy observations and symbiont location

The earliest microscopy observation of a DDA dates back to the beginning of 1900s when the Danish botanist Carl Emil Hansen Ostenfeld and biologist Johannes Schmidt described the genus Richelia associated with the diatom Rhizosolenia Brightwell (Ostenfeld and Schmidt, 1901). Few years later the German botanist Ernst Lemmermann reported C. rhizosoleniae living associated with the diatoms Hemiaulus Ehrenberg and Chaetoceros Ehrenberg (Lemmermann, 1905). Given the similar shape and the symbiotic nature of the two cyanobacteria, their taxonomical identity has been confused and re-interpreted in the coming years (Fig. 7; Gómez et al., 2005). 22

Fig. 7. Micrographs of (A, B) R. intracellularis in symbiosis with R. clevei, and (C) C. rhizosoleniae associated with C. compressus. From Gómez et al., 2005.

Light (LM) and electron microscopy (TEM) were the first techniques adopted to identify and analyze phytoplankton. While standard light microscopy (LM) was useful to identify Rhizosolenia/Chaetoceros-Richelia symbiosis, the symbionts in Hemiaulus, placed at the center of the valve, were hard to distinguish from the host protoplasm (Fig. 8; Villareal, 1991, 1992; Ferrario et al., 1995).

© Inter-Research 1991 Fig. 8. Top: Light micrograph showing a chain of Hemiaulus sp. Note: the brightness come from the diatom protoplast, whereas the symbionts are not visible. Bottom: epifluorescence picture of the same chain: R. intracellularis trichomes are clearly visible in yellow, and the host chloroplasts in red. Scale bar: 8 µm. With permission from Villareal, 1991.

Hence, epi-fluorescent microscopy became a pre-requisite to detect and identify the DDAs. In an earlier TEM study, combined immunolabelling with antibodies against proteins expected in the symbiont (e.g. nitrogenase, phycoerythrin) showed that R. intracellularis was located in the periplasm (i.e. the space between the cell wall and the cell membrane) in R. clevei diatoms (Janson et al., 1995). Gas vesicles were also absent in R. intracellularis vegetative cells (Janson et al., 1995). The same endobiotic location has been presumed to resemble the symbiosis with R. clevei in Hemiaulus diatoms, hence has been unresolved and not investigated. Unique among the DDAs is the Chaetoceros-Calothrix symbioses, where the symbiont is truly external, transversally attached to the intercellular space of the diatom chain at the terminal heterocyst (Norris, 1961). The draft genome of C. rhizosoleniae highlighted its similarity with the free-living cyanobacteria (e.g. size and content), although a few N-related pathways were still missing (i.e. urea transporters, urease, and GS inactivating factor; Hilton et al., 2013), likely due to its symbiotic nature. In addition, the number of trichomes varies depending on the length of the Chaetoceros chain (Gomez et al., 2005). Interestingly, the symbiont seems also capable of living freely, and 2 years after isolation the morphology of the Calothrix cells changed dramatically, i.e. longer trichomes than the symbiotic ones, and observations of intercalary heterocysts (Foster et al., 2010). Still unresolved is the mechanism by which the symbiont is transmitted to the diatom-host. Few studies on Richelia/Calothrix symbioses observed free- living cyanobacteria attached to the diatom frustule, albeit no penetration has ever been documented (Karsten, 1907; Norris, 1961; Sournia, 1970; Villareal, 1989, 1990; Gómez et al., 2005). Nevertheless, R. clevei-R. intracellularis culture experiments highlighted asynchronous growth cycles of host and symbiont, leading to asymbiotic hosts (Villareal 1989, 1990). It was suggested that symbionts propagation occurred either vertically, through the diatom cytoplasmic transfer during auxospurolation (i.e. size restoration through a specialized cell, the auxospore; Round et al., 1990), or horizontally, hence acquired from the environment (Villareal 1989, 1990). Field observations of H. hauckii-R. intracellularis symbiosis confirmed the symbiont vertical transmission, although in synchronicity with the symbiont division (Zeev et al., 2008). Hence, the modality by which the symbiont transmission occurs and if it varies in different DDAs, e.g. depending on the symbiont location and/or on the diatom cell symmetry, remain unknown.

DDA symbiont diversity and sequenced genomes. Beside single-gene phylogenetic approaches, draft genomes of four strains of Richelia/Calothrix have been reported (Hilton et al., 2013; Hilton, 2014). A major outcome of the genome sequencing was that genome size and content was related to location (Hilton et al. 2014). For example, the genome of the epi-symbiotic Calothrix SC (CalSC: 5.97 Mbp) is different in size and content than the endosymbiotic Richelia associated with Hemiaulus spp. (RintHM: 3.24 Mbp; RintHH: 2.21 Mbp). Conversely, the periplasmic Richelia associated with Rhizosolenia has a quite comparable genome size to the external symbiont (RintRC: 5.4 Mbp), although a few key pathways are still missing. In fact, the reduced genome of the internal R. intracellularis symbiont lacks important N acquisition pathways, such glutamine:2-oxoglutarate aminotransferase (GOGAT; note. this has not been reported for RintRC), transporters (i.e. ammonium, nitrate and urea) and key enzymes (i.e. urease, nitrite and nitrate reductase) typical of free-living cyanobacteria and also of the external symbiont genome (CalSC). As reported in other eukaryote-diazotroph symbiosis, e.g. UCYNA- prymnesiophyte (Tripp et al., 2010), environment and life history can shape and streamline the prokaryote genome (Giovannoni et al., 2014). In addition, the symbiont location plays a central role in genome erosion, especially if the symbiont is internal and isolated from the environment (Moran, 2003). Indeed, the smaller, albeit draft, genome of R. intracellularis associated with Hemiaulus spp. is evidence of a streamlining process due to an obligated symbiotic lifestyle (Hilton et al., 2013).

Diatom-hosts diversity and molecular markers. Up to now, the diversity of the diatom hosts have not been investigated and little is known about their phylogeny. Three plastidial genomes within the Rhizosolenia genus have been recently sequenced: Rhizosolenia imbricata (Sabir et al., 2014), R. setigera and R. fallax (Yu et al., 2018) (Fig. 9). Interestingly, R. imbricata and R. fallax reported photosynthetic gene loss (psaE, psaI and psaM, central genes of PSI), although still present in the earlier diverging R. setigera (Sabir et al., 2014; Yu et al., 2018). As previously hypothesized by Sabir et al. (2014), the reason behind this gene loss could be either a gene transfer to the nucleus, or horizontal/lateral gene transfer (HGT/LGT) to an internal symbiont, although no proof of either hypothesis are currently available. Another group from the University of Tsukuba (Japan), which studies the freshwater symbiotic diatoms of the family Rhopalodiaceae, stated the successful genome sequencing of the diatom host Epithemia adnata and claimed evidences of

HGT (Nakayama and Inagaki, 2016), however the genome is not publically available yet. Recent metatranscriptomic studies, e.g. the global Marine Microbial Eukaryote Transcriptome Sequencing Project (MMETSP; Keeling et al., 2014), provided high throughput information useful for downstream analysis on the symbiotic diatoms. For example, using the transcriptome of Rhizosolenia setigera as reference, Harke et al. (2018) analyzed the coordination between the metabolic pathways of the diatom-host Rhizosolenia. sp. and the symbiont R. intracellularis, and confirmed the interconnection of diatom-symbiont, especially when it comes to N and C metabolisms, as well as cell cycle.

Fig. 9. Phylogenetic reconstruction of 65 diatom rbcL-18S rRNA concatenated sequences (554 bp-410 bp) highlighting the Rhizosolenia clade (in red); the sequences derived from host DDAs are in blue. The tree has been inferred with a MCMC algorithm, and branch support are reported as percent probability; substitution rate bar=0.05; outgroup=Bolidomonas mediterranea. Modified from paper I.

Two of the most commonly used barcodes to resolve diatom phylogeny are the 18S SSU rDNA (Theriot et al., 2010; Zimmermann et al., 2011; Luddington et al., 2012; Guo et al., 2015) and rbcL (Theriot et al., 2010; Hamsher et al., 2011; MacGillivary and Kaczmarska, 2011; Kermarrec et al., 2013; Guo et al., 2015). Few other barcodes (e.g. COI, ITS, psbC) have been used, although mainly as concatenated genes to SSU and rbcL (Theriot et al., 2010; Guo et al., 2015; Trobajo et al., 2017). The 18S rDNA of eukaryotes contains nine variable regions (V1-V9) (Fig. 10; Neefs et al., 1993).

Fig. 10. Map of 18S rRNA gene. The variable regions (V1-V9) are marked in black, and the bp length is reported below. Modified from Ishaq et al., 2014.

In silico studies have shown that the V6 region is more conservative than the V2, V4, and V9 regions, which are considered the best biomarkers for eukaryotic biodiversity assessments (Hadziavdic et al., 2014). Meta-barcode studies on costal plankton have compared V4 and V9 regions and identified an equal number of Operational Taxonomic Units (OTUs) in the class Bacillariophyceae (diatoms) independent of the barcode used, suggesting that both regions are well represented in the databases (Tragin et al., 2018). Hence, the next criteria for choosing a valid biomarker is the resolution at genus/species level, and thus far the V4 region (the largest and variable of the 18S rRNA regions) showed a high resolution in the class Bacillariophyceae (Zimmermann et al., 2011; Luddington et al., 2012). Similarly, the large (L) subunit of the RubisCO enzyme (encoded by rbcL gene) is more conservative, if compared to the small subunit (encoded by rbcS; Xu and Tabita, 1996). Particularly, the ID form of the rbcL showed a substantial taxonomical resolution within the chromophytic phytoplankton (Paul et al., 1999, 2000; Mann et al., 2001; Wawrik et al., 2002; Tamura et al., 2005; MacGillivary and Kaczmarska, 2011; Li et al., 2013; Li et al., 2016).

DDAs abundance and ecological distribution

The marine DDAs have been largely observed in tropical and sub-tropical waters, especially in areas with mesohaline low-nutrient surface conditions. In these areas DDA abundances range from a few cells/L to high density, or bloom densities. These “hot-spots” are generally represented by areas where seasonally oceanic waters (high in salinity, but low nutrients) meet fresh-water (low salinity, high nutrient concentrations), such as near major river plumes, e.g. Amazon (Carpenter et al., 1999; Foster et al., 2007; Stenegren et al., 2017), Congo (Foster et al., 2009), Mekong (Bombar et al., 2011), Mississippi (Knapke, 2012), and Niger (Tovar-Sanchez et al., 2006; Foster et al. 2009). In brackish environments, e.g. the Baltic Sea, heterocystous cyanobacteria can also dominate the diazotrophic community, however these are free-living populations (Laamanen, 1997). Interestingly, symbiotic R. intracellularis and C. rhizosoleniae have not been reported in brackish waters, but largely observed in open oceans in symbiosis with diatoms, where they represent the only filamentous heterocystous cyanobacteria in direct competition with the widespread filamentous non-heterocystous N2-fixing cyanobacteria Trichodesmium. The DDAs have mostly been reported and quantified through microscopy observations and in the last decade by qPCR assays targeting the symbionts nifH gene (Table 2). The general condition(s) that constrain and influence DDA abundances and distribution is unresolved and difficult to study. Whereas diatoms typically thrive in coastal environments, characterized by high-nutrient supply (Goldman, 1993), an earlier study on R. clevei-R. intracellularis cultures suggested that these conditions negatively affect the symbiosis, especially when N-replete, resulting in the cyanobacteria expulsion by the diatom-host (Villareal, 1990). In addition, given that most modern N2- fixing heterocystous cyanobacteria reside in brackish or benthic habitats and are limited by high temperatures (Staal et al., 2003), the uniqueness of these associations lies in the fact that they thrive in environments where either partner would not grow if free-living (note: this apply to the internal symbioses), suggesting that perhaps their current habitat is a result of an ancient process of co-evolution and adaptation.

Table 2. Summary of the DDAs maximum abundances (cells L-1) in different oceans, detected both by microscopy and nifH qPCR. Tropical NA=Tropical North Atlantic; NPSG=North Pacific Subtropical Gyre; SP=South Pacific; SCS=South China Sea. Note: nm=not measured. From paper IV.

Study area Max Max References abundances abundances qPCR microscopy Tropical het-1: 105 het - 1: 10 3 Villareal, 1991, 1994; NA het-2: 109 het - 2: 10 3 Carpenter et al., 1999; het-3: 104 het-3: - Foster and Zehr, 2006; Foster et al., 2007, 2009; Subramaniam et al., 2008; Goebel et al., 2010; Großkopf et al., 2012; Ratten et al., 2015; Stenegren et al., 2017 NPSG het-1: 103 het - 1: 10 3 Mague et al., 1974; het-2: 103 het - 2: 10 2 Marumo and Asaoka, het-3: 106 het-3:<102 1974; Venrick et al., 1974; (cells/m2) Heinbokel, 1986; Vaillancourt et al., 2003; Ferrario et al., 1995; Gómez et al., 2005; Foster and Zehr, 2006; Wilson et al., 2008; Kitajima et al., 2009; Foster et al., 2010; Villareal et al., 2012; Follet et al., 2018 SP het-1: 105 het - 1: <50 Moisander et al., 2010, het-2: 104 het - 2: nm 2014; Turk-Kubo et al., het-3: 103 het-3: nm 2015 SCS het-1: 106 het - 1: 103 Chen et al., 2004; Chen, het-2: 104 het - 2 : nm 2005; Grosse et al., 2010; het-3: 103 het-3: nm Bombar et al., 2011; Zhang et al., 2011 Indian het-1: 104 het -1: 102 Kulkarni et al., 2010; Ocean het-2: 102 het-2: 102 Padmakumar et al., 2010; het-3: nm het-3: nm Jabir et al., 2013; Madhu et al., 2013; Manjumol et al., 2018 Mediter- het-1: nm het-1:<102 Zeev et al., 2008 ranean Sea het-2: nm het-2:<102

Other eukaryote-diazotroph associations

A similar marine eukaryote- diazotroph symbiosis to the DDAs is the one between the widespread unicellular diazotrophic cyanobacterium, Candidatus Atelocyanobacterium thalassa (UCYN-A) and a prymnesiophyte closely related to Braarudosphaera bigelowii Gran and Braarud (Fig. 11; Thompson et al., 2012). Also in this case the prokaryotic symbiont has been subjected to a significant genome reduction (UCYN-A=1.44 Mbp vs. the unicell Cyanothece ATCC Fig. 11. Double CARD-FISH assay targeting the prymnesiophyte host B. 51142, 5.4 Mbp) (Welsh et al., bigelowii (in green) and the symbiont 2008; Thompson et al., 2012), UCYN-A (in purple). Blue fluorescence including the reduction of key corresponds to DAPI staining. With genes for photosynthesis permission from Krupke et al., 2014. common to all other cyanobacteria (i.e. absence of photosystem II, Ribulose-1,5-bisphosphate carboxylase/oxygenase), and other fundamental pathways, including the tricarboxylic acid cycle (TCA) (Zehr et al., 2008; Tripp et al., 2010). A phylogenetic reconstruction based on the cyanobacteria sequence dated the appearance of the symbiotic UCYN-A ca. 90 Mya, in the Late Cretaceous (Cornejo-Castillo et al., 2016). Interestingly, the molecular dating of the symbiont coincided with the fossil evidence for its suggested host, Braarudosphaera spp. (Siesser et al., 1992; Bown et al., 2004), outlining an ancient co-evolution process between the two symbiotic partners. Thus, considering the similar nature of these eukaryote-diazotroph associations, common traits and differences have been summarized in Table 3.

Table 3. Comparison between DDAs, UCYN-A–prymnesiophyte and Rhopalodiaceae symbiosis. Unkn= unknown. Modified from Caputo et al., 2018 (paper II). *Hilton, 2014; **Hilton et al., 2013; ¤Zehr et al., 2008; ¤¤Tripp et al., 2010; ¤¤¤Nakayama et al., 2014.

Symbiont Host Environment Symbiont Symbiont Symbiont Host Photo- N2 lineages morphotype location genome genome synthesis fixation size-Mbp size R. R. clevei marine heterocystous endobiont 5.4* unkn + + intracellularis filamentous (periplasmic (het-1) space)

R. Hemiaulus marine heterocystous presumed 3.24 ** unkn + + intracellularis spp. filamentous endobiont 2.21 (het-2)

C. rhizosoleniae C. marine heterocystous external 5.97** unkn + + (het-3) compressus filamentous

A. thalassa B. marine unicellular endobiont 1.44¤ unkn - + (UCYN-A1) bigelowii relative A. thalassa B. marine unicellular Endobiont 1.48¤¤ unkn - + (UCYN-A2) bigelowii symbiosome

EtSB E. turgida fresh-water unicellular endobiont 2.79¤¤¤ unkn - potential

Aims of the thesis

Several previous studies on the DDAs have focused their attention on the prokaryotic partner, leaving little to no information about the diatom-hosts´ genetic identity, cellular location of the symbiont, and co-evolution. Moreover, the DDAs ecological distribution has been so far based on microscopy observations performed on a limited number of epi-fluorescent microscopy observations and/or qPCR assays targeting only the cyanobacterial partner. The evolutionary background and the ecological drivers that led to DDAs, are also unknown. In order to fill the abovementioned gaps of knowledge, this thesis aimed to:

1. Determine the genetic identity of the diatom-hosts and outline the evolutionary history that led to the modern-day DDAs, analyzing also morphological key traits and re-evaluating their ecological distribution.

2. Summarize similarities and discrepancies between two marine eukaryote-diazotroph symbioses, i.e. DDAs and prymnesiophyte- UCYN-A, and highlight biases that determined an underestimation of the DDAs populations.

3. Investigate the abundances and conditions conducive to diazotrophic cyanobacteria distribution in two regions of the South Pacific Ocean and in other ocean basins.

4. Develop a double catalyzed reporter deposition (CARD)-FISH assay specific for the DDAs in order to identify and quantify the presence of alternative life-stages (e.g. akinetes and/or auxospores) and/or free-living partners, and determine the drivers of distribution in environmental samples from the South China Sea.

Comments on methodology

Sampling areas and conditions

The samples processed in this thesis have been collected from different locations in the tropical and sub-tropical oceans, during several oceanographic expeditions (Fig. 12). In paper I samples for microscopy and genetic diversity were sampled from North and South West Pacific Ocean (NPAC and SWP), Western Tropical North Atlantic (WTNA), and South China Sea (SCS). Additionally, a short-lived (2 months) enrichment culture of H. hauckii-R. intracellularis was also processed. In paper III all the samples processed were collected in the Western Tropical South Pacific (WTSP) during the OUTPACE cruise (Oligotrophy to UlTra-oligotrophy PACific Experiment; Feb-Apr 2015). In paper IV samples used for in situ hybridization and qPCR analyses have been collected in the South China Sea (SCS).

The North and South Pacific oceans are characterized by a low concentration of nutrients (e.g. dissolved inorganic nitrogen and phosphate close to detection limit) and warm surface temperatures (25-30 ºC), where the diazotrophs represent one of the major sources of primary productivity and the only organisms capable of reducing atmospheric N2 to an available source of N, i.e. ammonium (Karl et al., 2012; Böttjer et al., 2017). On the other hand, the WTNA and SCS represent more dynamic environments, especially due to the influence of the Amazon and Orinoco Rivers (in the WTNA) and the Mekong River (in the SCS). Hence, when extensive inputs of riverine waters, low in salinity and nutrient-rich, meets the salty oligotrophic oceanic waters, creates intermediate conditions optimal for the diatom diazotrophs ecological success.

Fig. 12. A. Summary of the sampling locations and stations (red dots); B. South China Sea (SCS); C. North Pacific (NPAC); D. Western Tropical North Atlantic (WTNA); E. South West Pacific (SWP). Images by Andrea Caputo using ODV (Ocean Data View 4.7.1).

Cultures collections

Control samples for papers I and IV utilized cultured isolates of 3 strains of diatoms: Phaeodactylum tricornutum Bohlin 1898 (courtesy of Sara Rydberg Laboratory, SU, Sweden), Skeletonema marinoi Zingone & Sarno 2005 (provided by Prof. Monika Winder, SU, Sweden), and Chaetoceros socialis Lauder 1864 (provided by the Culture Collection of Algae and Protozoa, Scotland, UK); 5 strains of cyanobacteria: Calothrix rhizosoleniae (SC01; Foster et al., 2010), Crocosphaera watsonii WH8501 (provided by Prof. Jonathan P. Zehr, University of California, Santa Cruz), and Anabaena variabilis ATTC 29413, Nostoc punctiforme ATCC 29133, Nostoc sp2 (N. Moss2; Warshan et al., 2017) (provided by Prof. Ulla Rasmussen, SU, Sweden); an enrichment culture of H. hauckii-R. intracellularis (courtesy of Andrew E. Allen Laboratory; JCVI, La Jolla, CA USA). The conditions for culture maintenance are summarized in Table 4. 34

STRAIN MEDIUM T LIGHT/ STUDY (ºC) DARK P. tricornutum f/2 + Si 20 12:12 Paper I

S (Guillard, light:dark, at C. socialis 1975) 26 30 μmol Paper IV photons m−2 −1

DIATOM S. marinoi 20 s Paper IV

Ca. YBC II 26 16:8 Paper IV rhizosoleniae (Chen et al., light:dark 1996) cycle, at 30 μmol photons m−2 −1 s

A.variabilis BG110 20 continuous Paper IV ATTC 29413 (Rippka et light, at 35 al., 1979) μmol N.punctiforme photons m−2 Paper IV ATCC 29133 s−1

CYANOBACTERIA Nostoc sp2 Paper IV 16:8 C.watsonii YBC II 26 light:dark, at Paper IV WH8501 (Chen et al., 30 μmol 1996) photons m−2 s−1 H. hauckii-R. modified 26 16:8 Paper I intracellularis RMP light:dark

S medium cycle, at 30-

(Webb et 60 µmol

DDA al., 2001) photons m-2 s-1

Table 4. Summary of culture strains used in this thesis and maintenance conditions.

Microscopy

The first approach with microscopy (paper III) used a standard epifluorescence microscope (Olympus BX60) equipped with blue (460–490 nm) and green (545–580 nm) excitation wavelengths. These filter sets excite, respectively, the chlorophyll a (in both diatoms and cyanobacteria), and the phycobiliproteins (e.g. phycocyanin and phycoerythrin) typical of cyanobacteria. However widefield microscopy has a limited resolution due to the out of focus light, and detects single fluorophores. Thus, in order to both increase the resolution of the imaging and to combine channels, confocal microscopy was performed. In paper I, a Zeiss LSM 780 confocal microscope, equipped with 405, 488 and 561 laser lines was used. The 488 and 561 laser lines were simultaneously used to visualize the chlorophyll of the host diatoms and cyanobacterial symbionts (Ex430 nm/Em680-720 nm), and the phycobilins of the cyanobacteria (Ex561 nm/Em 562-672 nm). Additionally, the 405 laser line was adopted to visualize the nuclei/nucleoids, previously stained with DAPI (Ex358 nm/Em406-568 nm). Bit depth was set at 16, in order to visualize fine details, and voxel (i.e. the smallest unit of the sampled 3D volume) at 0.73 µm. Imaging was consistently conducted using the range indicator tool in order to prevent oversaturated pixels. Moreover, in order to resolve the 3D location of the symbiont in each symbioses, orthogonal views (xy, yz, xz) were imaged with the ZEN software (Zeiss), and further processed using the Contour Surface tool in IMARIS v.8.1 (Bitplane). The cyanobacteria autofluorescence was also used to measure the cell dimensions, whereas additional transmitted light was adopted to measure the diatoms cell diameter. Notably, in order to ensure that the fixation step using paraformaldehyde (PFA) was not creating any artifact (e.g. plasmolysis) culture samples of H. hauckii-R. intracellularis were imaged without fixative. The symbiont cellular location was further verified using transmitted electron microscopy (TEM; Zeiss EM 906). Previously fixed (3% PFA) samples of H. hauckii-R. intracellularis were embedded in LR-White resin (acrylic resin), and sectioned on an ultramicrotome (MT-7000 ULTRA, RMC). Ultrathin sections (60 nm) were further incubated with primary antibodies (anti- dinitrogenase reductase and antiphycoerythrin), followed by secondary antibodies (goat anti-rabbit IgG) and conjugated with 10 nm gold spheres (Janson et al., 1995; Foster et al., 1996). A confocal microscopy approach was also adopted in paper IV, using a Zeiss LSM 800. In the latter, in situ hybridized samples were simultaneously imaged with 488 (for visualizing the Alexa 488 dye; Ex488 nm/Em505-535 nm), and 640 nm (for detecting the Alexa 647 dye; Ex633 nm/Em657-687 nm) diode lasers, and cell abundances

36 enumerated. Common to all confocal microscopy observations was the slide preparation, using a 170 μm thick coverslip (#1.5) and a ProLong® Diamond Antifade Mountant (Molecular Probes). Thus, matching the reflective index of sample and objective, the point spread function (i.e. the light distortion) was reduced, and the resolution improved.

In situ hybridization

In paper IV a double catalyzed reporter deposition in situ hybridization (CARD-FISH) protocol was optimized and adopted for quantifying diatom- hosts and the heterocystous cyanobacteria strains (het-1, het-2, het-3). Abundances were estimated for each partner in both symbiotic and free-living cell states. The CARD-FISH protocol (Pernthaler and Amann, 2004; Pernthaler et al., 2004) was initially tested using universal prokaryotes/eukaryotes probes on the diatom and cyanobacteria cultures (diatoms: S. marinoii, C. socialis; cyanobacteria: C. rhizosoleniae, A. variabilis, N. punctiforme, N. sp2.). Subsequently, a CARD-FISH protocol was optimized for the same control cultures prior to application to the field collected samples. The rationale behind using a CARD-FISH approach, instead of the classic FISH, was a preference for the longer and stronger fluorescence intensity given by the horseradish peroxidase (HRP) labeled probes. However, HRP-labeled probes are much bigger than most in situ probes, thus require a more intense permeabilization step. Given the silica cell wall of the host diatoms, an aggressive permeabilization was required; similarly, the membranes around the cyanobacterial heterocysts are thicker than the respective vegetative cells, which indeed were harder to penetrate. The steps optimized in our CARD-FISH protocol for DDAs are summarized in Table 5. Step Description

Fixation 1% PFA for 1h at RT Embedding 0.1 % low-gelling-point agarose at 38-40 ºC Permeabilization (i). 10 mg mL-1 lysozyme for 1h at 35 ºC diatom (ii). 1% SDS for 1h at RT Permeabilization (i). 10 mg mL-1 lysozyme for 1h at 37 ºC cyanobacteria (ii). 1:10 Proteinase K for 1h at 50 ºC (iii). 1% SDS for 1h at 50 ºC Hybridization HB + 50 ng µL-1 probe for 3h at 35 ºC -1 TSA reaction AB + 0.5% H2O2 + 4 µg mL fluorescently labeled tyramide Slide preparation 1:100 DAPI/antifading mounting medium

Table 5. The various steps and description of the final optimizations for the double CARD-FISH assay. PFA=paraformaldehyde; SDS=sodium dodecylsulfate; RT=room temperature; HB=hybridization buffer; AB=Amplification buffer; DAPI= 4',6- diamidin-2-fenilindolo. From paper IV. 38

Based on the 18S and 16S rRNA sequences obtained in paper I, new HRP- labeled probes were designed for each of the symbiotic hosts (coupled with Alexa-647 fluorophore) and cyanobacteria (Alexa-488), respectively (Table 6).

Probe Target Sequence (5´-3´) Reference name HetCF Richelia CTACAAGACTWGA This study intracellularis GTRCGTTC Calothrix rhizosoleniae Rcle Rhizosolenia AGYGATCRGTCCR This study clevei GCTCT Hhem Hemiaulus hauckii AGCGATCRGTCCG This study GCRCT Ccom Chaetoceros AGGATCCGGTCCG This study compressus ACCTTTT Helpc-a Helper probe GTAGTCGCATTTCT This study C. compressus GGCGAG Helpc-b Helper probe GGTGGGTTCTTGGG This study C. compressus TCTCTT C. Competitor probe GTTTCGGTCCAGTA Krupke et watsonii- Crocosphaera GCAC al., 2013 732 watsonii WH8501 Eub 338 Most Bacteria GCTGCCTCCCGTAG Amann et GAGT al., 1990 Non 338 Antisense of ACTCCTACGGGAG Wallner et EUB338 GCAGC al., 1993 519r SSU universally GWATTACCGCGGC Embley et conserved KGCTG al., 1992. sequence 519r Antisense of 519r CAGCMGCCGCGGU Miller and complem AAUWC Scholin, ent 2000

Table 6. Summary of the 16S rRNA (HetCF) and 18S rRNA oligonucleotide probes (Rcle, Hhau, Ccom) used in this study, together with helpers, competitors, and positive/negative controls. Note. All the probes (except HelpC-A, B) were 5´-HRP conjugated. From paper IV.

Molecular analyses

In paper I single-gene Sanger sequencing was performed on amplicons (18S rRNA and rbcL) derived from samples containing 4 different symbiotic diatom-hosts (i.e. Rhizosolenia clevei, Hemiaulus hauckii, Chaetoceros compressus and Climacodium frauenfeldianum). The PCR products amplified a partial fragment of the V4 region (324 bp) of the small subunit of the 18S rRNA and in separate reactions the ID form of the large subunit of the ribulose-1,5-bisphosphate carboxylase/oxygenase (RuBisCO; rbcL; Table 7). The rationale behind using the V4 was the following reasons: (i) the V4 region has a larger database of sequences for comparison and includes several large scale metagenomic studies (e.g. the circumnavigation expedition Malaspina; Duarte, 2015); (ii) the V4 region is longer than the V9 ( ̴ 360 bp vs. ̴160 bp), thus it covers a better variability of the 18S rRNA gene; (iii) previous studies showed that the V4 has a high phylogenetic resolution in the phylum Stramenopiles (Pernice et al., 2013), specifically in the class Bacillariophyceae (Zimmermann et al., 2011). In addition to the single-gene sequencing of the diatom-hosts, the respective symbiotic cyanobacteria (R. intracellularis, C. rhizosoleniae, and C. watsonii) were amplified using the 16S rRNA and the nifH biomarkers (Table 7). Noteworthy, prior to the polymerase chain reaction (PCR) amplification, a few samples containing single symbiotic hosts were processed first with a whole genome amplification (WGA) step. Applied on single-cells, WGA provides a sufficient quantity of replicated DNA for downstream analysis, although it also increases the biases rate (Sabina and Leamon, 2015). The sequences obtained in this study for both partners, together with the 16S rRNA and nifH sequences from a previous complementary study (Foster and Zehr, 2006), were blasted on NCBI (selective for environmental sequences), KEGG MGENES (www.genome.jp/mgenes/) and in the IMNGS databases. Sequences with similarity ≥ 98% were selected and the respective sampling sites noted. The number of clones for each sequence and the distribution of the blast hits were shown on a global map. The abovementioned sequences were also part of a phylogenetic molecular-clock trait based analysis to trace the ancestor of phenotypes common to modern day populations.

Primer Code Target Approx. Reference Sequence fragment (5´-3´) length (bp) rbcL AK rbcL 554 Paul et al., GATGATGA (Form (Form 2000; RAAYATTA ID)-F/R ID) Li et al., 2013 ACTC ATTTGDCC ACAGTGDA TACCA D512for AL 18S 390-410 Zimmermann ATTCCAGC 18S rRNA et al., 2011 TCCAATAG (V4 CG D978rev region) GACTACGA 18S TGGTATCT AATC nifH-F O nifH 324 Olson et al., CGTAGGTT 1998* GCGACCCT AAGGCTGA nifH-R GCATACAT CGCCATCA TTTCACC 27F AA 16S 1000 Lane, 1991 AGAGTTTG rRNA ATCMTGGC TCAG 1492R TACGGTTA CCTTGTTA CGACTT

Table 7. Summary of the primers used in paper I. * primer set specific for cyanobacterial nifH fragments. From paper I (Supporting Information Table S2).

Quantitative polymerase chain reaction (qPCR)

In paper III and IV qPCR analyses was used to enumerate the DDAs in depth- profile samples. In paper III, nifH primer and probe sets (Table 8) were used to quantify the major lineages of diazotrophic cyanobacteria (Trichodesmium, R. intracellularis, C. rhizosoleniae, UCYN-A1, -B, and -C). Additionally, UCYN-A2 and the UCYN-A1 and A2 hosts were also detected. Notably, the UCYN-A1 host´s primer/probe set was newly designed, based on the 18S rRNA sequence reported from station Aloha (Thompson et al., 2012). On the other hand, paper IV focused exclusively on the nifH quantification of the 3 het groups. The qPCR assays, initially performed on the ship in paper III, were subsequently verified in the laboratory. The choice of re-running the qPCR samples was due to multiple reasons: (i) the diazotrophic relative abundance was a discriminating factor for the selecting the stations during the expedition, thus a shorter protocol (i.e. shorter DNA extraction protocol, a shorter qPCR conditions with corresponding assay chemistry) was adopted for the “on board” qPCR; (ii) the qPCR assays relies on the quality of the DNA extract, which varies depending both on the specific sample and the extraction procedure, thus replicates and/or complementary techniques are preferred. Hence, in our study epi-fluorescent microscopy was adopted to verify cell abundances and cell integrity. A further caveat of the qPCR assay is the probe specificity, which can over/under estimate the target. Thus, cross-reactivity tests at different conditions were conducted. Specifically, fast vs. standard mode were tested: holding, denaturation and annealing steps were run at 95º C for 20 s, followed by 45 cycles of 95º C for 1 s and 60º C for 20 s (i.e. fast mode), and compared with longer intervals (standard mode; 11 min and 40 s, 14 s, and 40 s, respectively); additionally, two different annealing temperatures were tested, i.e. 64 vs. 60 ºC.

Piecewise Structural Equation Model (SEM). In order to understand the interactions among biotic and abiotic factors characterizing a water mass, in relation to the DDAs´ abundance and distribution, a SEM model was adopted in paper IV. The SEM is a multivariate statistical model, which predicts the relative strength (or importance) of each variable and how they influence (positively or negatively, directly or indirectly) a certain relationship (Grace, 2006). Hence, besides disentangling complex ecological networks, a further strength of this approach is the causality, i.e. it creates a framework for linking back an ecological observation to a theoretical concept (Grace et al., 2010).

Phylotype Forward Probe Reverse Reference Primer 5' to 3' Primer 5' to 3' 5' to 3´ UCYN-A1 AGCTATA TCTGGTGG ACCACGA Church et ACAACG TCCTGAGC CCAGCAC al. 2005 TTTTATG CTGGA ATCCA CGTGA UCYN-A2 GGTTACA TCTGGTGG ACCACGA Thompson ACAACG TCCTGAGC CCAGCAC et al. 2014 TTTTATG CCGGA ATCCA TGTTGA UCYN-B CGTAATG TGTGCTGG CACGACC Moisander CTCGAA TCGTGGTA AGCACAA et al. 2010 GGGTTTG T CCAACT A het-1 CGGTTTC TCCGGTGG AATACCA Church et CGTGGTG TCCTGAGC CGACCCG al. 2005 TACGTT CTGGTGT CACAAC het-2 TGGTTAC TCTGGTGG AATGCCG Foster et CGTGATG TCCTGAGC CGACCAG al. 2007 TACGTT CTGGTGT CACAAC het-3 CGGTTTC TCTGGTGG GAGCGGG Foster et CGTGGC TCCAGAA TGTCGGA al. 2007 GTACGTT CCTGGTGT GACGGAT Tricho- GACGAA CATTAAGT CGGCCAG Church et desmium GTATTGA GTGTTGAA CGCAACCT al. 2005 AGCCAG TCTGGTGG A GTTTC TCCTGAGC UCYN-A1 AGGTTTG CTGGTAG GAGCGGG paper III Host CCGGTCT AACTGTCC TGTCGGA GCCGAT T GACGGAT UCYN-A2 GGTTTTG CTGGTGCG GAGTGGG Thompson Host CCGGTCT AGCGTCCT TGTCGGA et al. 2014 GCCGTT TCCT GACGGAT

Table 8. Summary of the primer and probe sets used in papers III and IV. Note. In grey the ones uniquely used in paper III. From paper III (Supplement, Table S1).

Phylogenetic analyses

In paper I, the sequences retrieved from the diatom-hosts (18S rRNA, rbcL) and for the cyanobacteria (16S rRNA, nifH) were concatenated in two separate alignments in order to increase the phylogenetic accuracy. An independent branch rate relaxed-clock model (IGR; Lepage et al., 2007) was applied and combined with a fossilized birth-death process (FBD; Zhang et al., 2016) on both concatenated alignments. Phylogenetic trees were inferred using MrBayes v.2.3.7 (Ronquist et al., 2012). Noteworthy, the selection of species for the alignments was based both on the phylogenetic relatedness to the species of interest and on the availability in NCBI of sequences derived from the same strain/genome. Genes from other known symbiosis were also part of the alignments (e.g. UCYN-A; Rhopalodiaceae diatoms and the respective cyanobionts). Thus, our dataset was restricted to 65 species of diatoms and 49 of cyanobacteria. In addition, the rationale of choosing a relaxed-clock model analysis was to maximize the robustness and the biological strength of the analysis, after testing alternative models (e.g. strict-clock). Additionally, in order to calibrate the trees multiple “clocks” were selected from the literature and applied. Based on the clock-tree analysis, an ancestral (character) state reconstruction (ASR) of a selective number of traits (morphological and environmental) for the diatom-hosts and the symbiotic cyanobacteria was performed. A total of 3 and 5 traits were selected, respectively, and 2 ̶ 4 character states were assigned, based on both observations and literature searches. ASR can be fundamental for determining how traits (e.g. metabolic, morphological, ecological) have evolved along an ancestor´s descendent line till the modern organisms, and explain what determined the acquisition, development, or inheritance of a certain character, e.g. bearing a symbiont.

In order to identify the phylogenetic closest strains that could potentially cross-hybridize our targets, a Bayesian Inference was performed in paper IV prior to the probe design. The latter was ran using Geneious® 9.1.6, setting number of cycles, sample frequency and portion of chain discarded in default mode (11·105, 200 and 500 respectively). For the diatom 18S rRNA probes an alignment with 137 sequences from the Protist Ribosomal Reference database (PR2; Guillou et al., 2012) was performed and the closely related species to those of interest (95-99% sequence identity) were checked for probe mismatches. Similarly, the newly designed probe (HetCF) was tested using the TestProbe tool in SILVA database (one of the largest ribosomal RNA database currently available), and the strains with high sequence similarity (>98%) verified for cross-reactivity.

Results and Discussion

Re-evaluation of symbiont location and morphology using advanced microscopy Around 40 samples, 3 oceans and more than 300 symbiotic cells were analyzed in paper I in order to review the symbiont location in the diatom- hosts. As previously described, the approach adopted to investigate the latter was confocal microscopy, due to its high resolution power and the 3D reconstruction tool. Analyzing the orthogonal reconstructions of H. hauckii- R. intracellularis, ca. 90% (215 of 238 cells) of the symbionts were observed (partially to fully) surrounded by the chloroplast auto-fluorescence of the diatom-host (Fig. 13).

Fig. 13. Confocal micrograph of a H. hauckii-R. intracellularis symbiosis simultaneously imaged with 488 and 561 laser lines: (a) z-stack image, (b) orthogonal reconstruction in xy, xz and yz, (c) 3D reconstruction using the Contour Surface tool of the software IMARIS (Bitplane). White arrows show the diatom-host chloroplast fluorescence (in green), surrounding the symbiont (in orange). Scale bars: 5 µm. From paper I.

Additionally, the absence of artifacts due to fixation (e.g. membrane dysplasia, chloroplast disruption) was positively verified on culture samples of H. hauckii-R. intracellularis without chemical fixation, which showed the same pattern visualized in the PFA fixed samples, i.e. the symbionts were partially to completely surrounded by the diatom chloroplasts (paper I, Suppl. Fig. 1b,

45 e). Moreover, 4',6-diamidin-2-fenilindolo (DAPI) staining showed the symbiont in close proximity to the host diatom nucleus, confirming a cytosolic location of the cyanobacteria in the diatom-host (paper I, Fig. 3). This finding was further verified by Transmission Electron Microscopy (TEM) on a cross- section of a H. hauckii-R. intracellularis cell (paper I, Suppl. Fig. 3). Prior to this study, the location of the symbiont in Hemiaulus spp. was assumed to resemble that of Rhizosolenia, i.e. with the symbiont(s) located in the space between the plasmalemma (i.e. periplasmic space) and the silicified cell wall (Villareal, 1992; Janson et al., 1995). However, in some earlier observations of Hemiaulus spp.-R. intracellularis symbiosis were carried out with widefield microscopy (phase contrast filter), and most of the time they were not able to resolve the symbiont location, due to the diatom chloroplasts masking the symbiont fluorescence (Ferrario et al., 1995). In our study, the same confocal approach was adopted when observing Rhizosolenia-Richelia and Chaetoceros-Calothrix symbiosis, and the symbiont location was consistent with the literature, i.e. in Rhizosolenia the symbionts were located between the plasmalemma and the frustule with the terminal heterocyst angled towards the diatom valve (Sundström, 1984; Villareal, 1989), whereas in Chaetoceros the symbionts were externally attached with the terminal heterocyst between two diatoms frustules (Norris, 1961; Gómez et al., 2005). Additionally, the number of trichomes of Calothrix per Chaetoceros-chain was similar to the latter study (Gomez et al., 2005), suggesting that attachment space might be fundamental for the episymbionts, and perhaps host-regulated. In addition to resolving the symbiont location in Hemiaulus spp., morphometric parameters were measured for both partners, and a trend showing the presence of longer trichomes in wider cell diameter hosts outlined, suggesting that the host-might control the symbiont growth. In paper III epifluorescence microscopy was adopted to verify the diazotroph abundances determined with qPCR assays on samples collected in the South West Pacific. Microscopy counts were performed for the previously microscopy identifiable diazotrophic cyanobacteria, i.e. Trichodesmium, het- groups, and UCYN-B. Interestingly, a significant discrepancy between nifH gene copies and microscopy cell counts was encountered. However, considering that (i) in some cyanobacteria, including heterocystous types, polyploidy occurs and can be influenced by cell cycle (Griese et al., 2011; Sukenik et al., 2012; Sargent et al., 2016), (ii) more than one nifH gene copy per organism occurs in some cyanobacteria, (iii) heterocystous cyanobacteria possess a genome copy in each cell along the filament (e.g. vegetative cell, heterocysts), (iv) probe specificity can be low and lead to cross-hybridization, (v) extraction efficiency is not 100% nor equal across all targets, and (vi) 46 sampling biases, over/under estimations of a particular target can occur. In addition to cell counts, cells conditions were also checked and linked to environmental parameters. Noteworthy, at LDA, 16 m depth, many free Richelia-like cells were observed. This observation became the main impetus of the next study (paper IV).

In situ hybridization determines DDA distribution and abundance.

Results from Paper I and III left some open questions that have been mostly addressed in paper IV, e.g. what is the best approach for estimating the DDAs without risking to over/under-estimate their abundance? Can the symbionts and hosts live freely? Do they have multiple life stages and other phenotypes during these life stages? In an attempt to address these questions an ad hoc in situ hybridization assay was designed, aimed to specifically bind the rRNA of the two partners for subsequent visualization with confocal microscopy (Fig. 14).

Fig 14. Confocal simultaneous imaging of the newly designed 18S rRNA (purple; Alexa 647, Ex633 nm/Em657-687 nm) and 16S rRNA (green: Alexa 488, Ex488 nm/Em505-535 nm) double CARD-FISH assays specific for DDAs. (a) R. clevei-R. intracellularis, (b) asymbiotic R. clevei, (c) asymbiotic R. intracellularis/C. rhizosoleniae, (d) Hemiaulus spp.-R. intracellularis, (e) asymbiotic Hemiaulus spp., (f) asymbiotic R. intracellularis, (g) C. compressus-R. intracellularis, (h) asymbiotic C. compressus, (i) asymbiotic R. intracellularis [TL]. Scale bars: a-c, g-h, 10 μm; d- f, i, 7 μm. Picture from paper IV.

The first challenge was designing a double CARD-FISH assay with a permeabilization step intense enough to penetrate the multiple layers of both partners, and, at the same time, maintain the cell integrity (Table 5, page 41). The next challenge was designing specific probes for both partners and use fluorophores that would not interfere with the natural autofluorescence of both cell types. In total 70 samples from 18 stations in the SCS were processed with the newly designed CARD-FISH assays. The abundance of the DDAs by the CARD-FISH assays was compared with nifH abundances for the 3 het- symbiont strains by the commonly used qPCR assays. Asymbiotic diatom- hosts and cyanobacteria were found almost at every station, although at low abundances (<50 cells/L). Conversely, the symbiotic DDAs showed over all higher abundances (except for Chaetoceros-Calothrix which max abundance was <50 cells/L). Albeit the orders of magnitude between the 2 methods varied greatly (up to 108), similar patterns with depth and salinity were observed between the CARD-FISH and qPCR. In fact, symbiosis were more abundant in the upper euphotic zone (0-20 m) and decreased with depth, whereas the asymbiotic cells followed an opposite trend (paper IV, Fig. 4a, b). Additionally, high nutrients concentrations and low salinities seemed also to be conducive to symbioses breakdown. No alternative stages or auxospores were observed for the diatom-hosts, aside from asymbiotic cells. Conversely, the asymbiotic cyanobacteria, both found as entire trichomes and single-cells, suggested the possibility of an intermediate life-stage. Moreover, the three different DDAs (Rhizosolenia, Hemiaulus, Chaetoceros) co-occurred at each detection depth, which was consistent with the positive interactions outlined among the three DDAs by our Piecewise SEM model, suggesting a lack of intra-specific competition, likely due to intra-specific variability. Our observations were not the first evidences of asymbiotic partners (e.g. Sournia, 1970; Marumo and Asaoka, 1974; Villareal, 1990; Gómez et al., 2005; Foster et al., 2007, 2011; White et al., 2007; Grosse et al., 2010; Bombar et al., 2011; Villareal et al., 2014), although never before have DDAs been identified and quantified using a CARD-FISH approach. Although these findings may undermine the cyanobacteria host-specificity described in previous studies

(Janson et al., 1999; Foster and Zehr, 2006), the cyanobacteria expulsion by the diatom-hosts could also be explained by a nutrient-replete environment (Villareal, 1990), or other environmental stressors (e.g. salinity, temperature) which might cause the symbiont release, similar to what occurs in other systems, e.g. Cnidarian-dinoflagellates symbiosis (Weis, 2008). Finally, sample processing could also cause cells to dislodge, although the vertical and horizontal patterns of distribution of the asymbiotic cells shown in our study strongly support the intermediate life-stage hypothesis.

Molecular-clocks outline the DDAs appearance and the ancestors’ traits.

In paper I single-gene sequencing of the diatom-hosts (18S rRNA and rbcL) and the respective symbionts (16S rRNA and nifH) was performed. The concatenated alignment of the symbiotic heterocystous cyanobacteria confirmed the clades described in previous studies (i.e. het-1, -2, -3; Fig. 15A; Janson et al., 1995; Foster and Zehr, 2006), and broadened each group with more sequences.

Fig. 15A. 16S rRNA-nifH concatenated molecular-clock and ancestral state reconstruction of the symbiotic cyanobacteria (in bold). The grey bars represent the uncertainties and red bars the calibrated nodes. Two traits are displayed for each node (life history and environment), and the pie charts indicate the trait probability. Picture from paper I.

New to the field is the report of the nifH sequence of the symbiont of C. frauenfeldianum. Previous studies only reported its 16S rRNA sequence (Carpenter and Janson, 2000), and the concatenation of the two sequences (nifH-16S) placed this strain closely related to C. watsonii WH8501 (Fig. 15A).

Fig. 15B. 18S rRNA-rbcL molecular-clock and ancestral state reconstruction of the diatom-hosts (in bold). The grey bars represent the uncertainties and red bars the calibrated nodes. Two traits are displayed for each node (colony formation and life history), and the pie charts indicate the trait probability. Picture from paper I.

The respective diatom partners were clustering separately from each other, although it was expected due to the phylogenetic distances between the genera they belong to (Fig. 15B). Consistently, the diatom-host C. frauenfeldianum, belonging to the family Hemiaulaceae, clustered in the Hemiaulus clade. Hence, the symbiosis between C. frauenfeldianum and Crocosphaera, thus far poorly studied due to its rare nature (Carpenter and Janson, 2000), has finally a genetic identity for both partners. Additionally, using molecular-clocks, i.e. fossil records and/or geological events, an estimate of the geological age of diatom-hosts and symbionts was outlined (Fig. 15A, B). The first appearance of the diatom-hosts was estimated in the Cretaceous, ca. 100 Mya, consistent with the fossil records from previous studies (Agnese and Clark, 1994; Kemp et al., 1999), and coincident with evidence for the presence of N2-fixing cyanobacteria (Rau et al., 1987; Kemp et al., 1999; Kuypers et al., 2004; Kashiyama et al., 2008). Interestingly, the symbiotic cyanobacteria divided from their sister clade much earlier, ca. 1500 Mya, although the major differentiation occurred between 200-100 Mya. In addition, the diatoms with internal symbionts (R. clevei and H. hauckii) have been estimated to be more ancient than the external-bearing associations (i.e. Chaetoceros-Calothrix) (Fig. 15B), suggesting the symbiont internalization might be part of an evolutionary process. Moreover, given the eroded (draft) genome of the “truly” internal symbiont (het-2) compared to the epiphytic strain (het-3), suggest that symbiont cellular location might be relevant for host control and genome streamlining. In order to understand the evolutionary background of the DDAs, other planktonic eukaryote- diazotroph symbioses were also compared (paper II; Table 9). The genomic comparison performed in paper II using an on-line annotation tool (i.e.RAST, Rapid Annotation using Subsystem Technology) for the het symbionts and the 2 UCYN-A symbionts estimated a lower number of transporters in the internal symbionts (het-2 the lowest) than het-3, suggesting a larger investment in the genome dedicated to transport/ exchanges (e.g. nutrients) for the external symbiont. Altogether, these evidences suggest that the location of the diazotrophic symbionts in the eukaryotic hosts and their evolutionary history might have influenced differently streamlining of symbiont genomes. Complementary to the molecular-clock model, an ancestor state reconstruction was performed in paper I, for both cyanobacteria-symbiont and diatom-host. According to the phylogenetic reconstruction, the key traits that characterized the DDA ancestors and perhaps determined their ecological success were: short terminal heterocystous trichomes (for the cyanobacteria, paper I, Fig. S7-11),

Symbiont R. intracellularis R. intracellularis C. rhizosoleniae A. thalassa A. thalassa EtSB

Lineages (het-1) (het-2) (het-3) (UCYN-A1) (UCYN-A2) - B. bigelowii Host R. clevei Hemiaulus spp. C. compressus B. bigelowii E. turgida relative Periplasmic Endobiont Location Endobiont External Endobiont Endobiont space symbiosome Approx. Age 50 60 30 125 125 25 (Mya) Genome 3.24** size 5.4* 5.97** 1.44¤ 1.48¤¤ 2.79¤¤¤ 2.21 (Mbp)

Table 9. Comparison between het-symbiosis and other eukaryote-diazotroph associations. Note. “Genome size” refers to the prokaryotic partner. Modified from paper II. * Hilton, 2014; **Hilton et al., 2013; ¤Zehr et al., 2008; ¤¤Tripp et al., 2010; ¤¤¤Nakayama et al., 2014.

53 a strongly silicified frustule, and the formation of chains (for the diatoms, paper I, Fig. S4-6). Interestingly, the cyanobacteria ancestor was likely adapted to fresh-water, which was consistent with a recent study dating the appearance of marine cyanobacteria in the late Proterozoic (Uyeda et al., 2016; Sánchez-Baracaldo et al., 2017). This finding suggested that perhaps the transition of the cyanobacteria to the open-ocean environment might have occurred later, in symbiosis with their hosts.

Sequence-based survey broadens the DDA ecological distribution Thus far, the detection of DDAs, using molecular and microscopy based approaches, has largely confined the distribution of the symbiosis to the tropical and sub-tropical oceans, within the 18ºC isotherms (Table 2, pg 29). Using the new sequences obtained in paper I (18S rRNA, rbcL, 16S rRNA, and nifH) a nucleotide BLAST survey against several databases including environmental and genome libraries (e.g. NCBI, KEGG MGENES, IMNGS) was performed. Most of the hits (min of 98% nucleotide similarity and 350 bp min coverage) corresponded to areas where the DDAs have been previously observed and/or quantified (e.g. the tropical and sub-tropical belt; Fig. 16). Rather low was the total number of hits (ca. 300), considering several large scale sampling efforts of the global ocean currently available (GOS, TARA, Malaspina) coupled with high-throughput sequencing. However, in paper II the sampling procedures of the global surveys were reviewed and we identified the use of pre-filtration steps (i.e. 20–200 μm mesh) necessary for excluding larger eukaryotes (e.g. fish larvae), likely excluded the DDA templates. In addition, reviewing ca. 50 nifH qPCR studies, less than 30% of the reports targeted the 3 het-groups (paper II), whereas UCYN-A was quantified in 96% of the studies, suggesting a global under-estimation of the DDAs due to sample processing biases. Somewhat surprising was that ca. 7% of the sequence-hits were retrieved in atypical environments, such as high latitudes, the gut content of copepods, and anoxic environments, suggesting that the distribution of DDAs might be broader than previously thought. Noteworthy, the areas where only one partner was detected could be explained as (i) a less-obligatory nature of the symbioses than previously thought (e.g. free-living states), (ii) limitation of the database (e.g. different region of the same genetic barcode or un-sampled regions), (iii) taxonomical resolution of the molecular markers used. Hence, comparative approaches (e.g. CARD- FISH) and a further genetic characterization of the DDAs might address this caveats.

Fig. 16. BLAST survey of diatom-host (18S rRNA, rbcL) and symbiont sequences (16S rRNA, nifH). Genes and organisms are color-coded and follow the legend on the right. The oceans show the surface temperatures (white-grey bar on the right); 18ºC and 25ºC isotherms, and ocean acronyms are reported on the map. Modified from paper I.

Conclusions

The association between marine diatoms and di-nitrogen fixing cyanobacteria has been known for over a century (Ostenfled and Schmidt, 1901), nevertheless still little is known about the eukaryotic hosts and how the two partners interact. This thesis aimed to cover some gaps of knowledge by coupling genetic information with microscopy tools, and contextualizing the findings with the diazotrophic community of the open-ocean. Confocal microscopy coupled with 3D reconstruction showed the internal location of the symbiont Richelia intracellularis in the diatom-hosts H. hauckii, with additional evidence by TEM. This finding revealed a continuum of integration between diatom and cyanobacteria, from the externally attached symbiont, to “truly” endosymbiont, passing through an intermediate state of “quasi-internal” (i.e. the periplasmic location). Consistent with the symbiont cellular location, the geological appearance of the DDAs suggested a gradual symbiont integration, i.e. internal symbionts were estimated earlier than the external one. The latter outlines a co-evolutionary process and key-traits conducive to the DDAs ecological success. Symbiont cellular location is also important for nutrient transport between host and symbiont, indeed the epi-symbionts (i.e. het-3, UCYN-A) showed a higher number of genes (normalized to genome size) involved in transport, compared to the internal symbionts (i.e. het-1, het-2). These evidences suggest that internal symbionts are likely metabolically dependent on the host. Additionally, due to methodological and sample processing biases (e.g. pre- filtration steps), the DDAs seem to be underestimated. Consistently, the gene- based survey suggested a wider distribution of the symbiotic partners than previously predicted, and likely wider ecological optima. The optimized double CARD-FISH assay proved to be an effective means to simultaneously detect host and symbionts, overcoming some of the biases characterizing the nifH qPCR assays, e.g. not being able to distinguish the symbiotic state. Indeed, the in situ approach showed the asymbiotic status of the DDAs partners, suggesting that the qPCR assays performed so far (targeting exclusively the cyanobacterial partner), might have overestimated the symbioses. Noteworthy, the asymbiotic single-cell Richelia/Calothrix suggested a different life-stage than the filamentous one, although further evidences are needed. Conversely, an auxospore stage of the diatom-hosts has not been detected. Finally, in 2 very different regions of the world’s oceans (SCS, SWP), DDAs persist, although when sampled for this thesis, were at background densities. 56

Nevertheless, the conditions driving abundances and distributions are similar, and salinity and depth seem to be the main players. This thesis represent a step more towards understanding the intricate symbiosis between diatoms and N2-fixing cyanobacteria, although some pieces of this fascinating puzzle are still missing.

Future perspective

This thesis disclosed morphological, genetic, phylogenetic and ecological aspects of the DDAs that thus far were unknown, or misinterpreted. Nevertheless, as Voltaire declaimed “Judge a man by his questions rather than by his answers”, this thesis also raised some more questions, such as:

What role have host and environment been playing in the symbiont genome streamlining? The new insights on the symbiont location and the geological cascade of symbiotic events described in paper I, outline an on-going co-evolutionary process leading to the symbiont genome streamlining, likely retaining exclusively their N2 fixation capability. Similar systems, e.g. UCYN-A– prymensiophyte (Thompson et al., 2012) and Rhopalodiaceae–“Spheroid bodies” (Nakayama et al., 2014), already show photosynthetic gene loss by the symbiont, and the latter study hypothesized a horizontal gene transfer (HGT) between the two partners. Interestingly, the plastidial genome sequencing of a closely related Rhizosolenia, showed the lack of some photosynthetic genes in the diatom, suggesting a possible HGT to either the nucleus or an endosymbiont (Sabir et al., 2014; Yu et al., 2018). Thus, in order to clarify the real nature of these symbioses and investigate for HGT and/or lateral gene transfer (LGT) between diatom and symbiont further sequencing is needed. The Richelia/Calothrix genomes are partially sequenced and remain in draft form (857, 90, 941 and 620 contigs, het-1, 2A, 2B, and 3, respectively), hence longer read sequencing (e.g. using a PacBio platform) would correct the current assemblies and, likely, complete the gaps in the genomes. A parallel approach should be adopted for the diatom-hosts, and similar to what has been recently achieved for the R. clevei transcriptome (Harke et al., 2018), using the plastidial genome of a closely related Rhizosolenia, e.g. R. setigera, as reference genome, build a new assembly. Alternatively, single-cell techniques might be adopted to isolate the symbiosis and sequence their 57 genome/transcriptome. For example, through laser capture microdissection (LCM) a single-cell can be excised from an environmental sample and isolated in a tube for downstream analyses. Finally, the comparison between symbiotic strains and asymbiotic ones (e.g. Nostoc punctiforme PCC 73102 for the cyanobacteria, and Thalassiosira pseudonana for the diatoms) could be useful to examine for HGT/LGT.

To which extent is the host controlling the metabolic exchanges with the endosymbionts? In paper I confocal microscopy and TEM showed the cytosolic location of the symbiont in the diatom-host Hemiaulus, suggesting a higher degree of interaction between the two partners. In addition, in paper II the number of transporters in R. intracellularis (het-2) was estimated lower than the other symbionts (het-1, het-3), outlining a different pathway of metabolite transfer and communication between the prokaryotic partner and the diatom-host.

Previous studies showed the transfer of fixed N2 by the cyanobacteria to the diatom-host (Foster et al., 2011), and suggested a C exchange in the opposite direction (Hilton et al., 2013; Harke et al., 2018). Nevertheless, the mechanism by which the C metabolites are transferred to the symbiont is still unknown, as well as how the different cellular locations of the symbiont can influence this exchange. Hence, further transcriptomic and proteomics studies would lead to a better understanding of how symbiont and host metabolisms are interwoven and how it varies in the endo- vs epi-symbiosis. Alternatively, the level of expression and co-regulation in the two partners could be investigated using spatial in situ RNA techniques, e.g. mRNA-FISH (Femino et al., 1998; Raj et al., 2006, 2008; Larsson et al., 2010; Ke et al., 2013; Muller et al., 2013).

For example, targeting the symbiont nifH gene (for nitrogenase and N2 fixation) simultaneously with a probe designed to target the diatom-host amtB gene (for ammonium transport; Satinsky et al., 2017) would visually highlight the pathway that the fixed N2 follows after the diazotrophic reduction to NH3, while highlighting coordination between host and symbiont. Nevertheless, + this assumption might be adequate if we consider an active transport of NH4 ; alternatively NH3, a small gas molecule, might simply diffuse through the cyanobacteria membranes (Ritchie and Gibson, 1987), and reach the diatom- host cytosol. Another pathway that could determine host-symbiont coordination would be targeting mRNAs involved in cell growth and cell cycle. In paper IV the detection of asymbiotic host partners suggested the

58 possibility of a horizontal infection of the symbiont, hence symbiont and host growth might not be coordinated as previously described (Zeev et al., 2008). Thus, targeting the diatom silica deposition proteins (SAPs; Tesson et al., 2017) in conjunction with the cyanobacteria FtsZ involved in cell-division (Dohert and Adams, 1995; Zhang et al., 1995) could address these questions. Thus, the mRNA-FISH can be an excellent technique to localize mRNA molecules on a single-cell level, reducing the bias and caveats introduced by other approaches (e.g. RT-qPCR).

What factors control the development of a singular terminal heterocyst in Richelia/Calothrix? The ancestral state reconstruction presented in paper I outlined the terminal heterocyst of the symbiotic cyanobacteria as one of the traits acquired through the evolution, and perhaps in correspondence of their transition from freshwater environments to a marine symbiotic life-style. Moreover, considering that R. intracellularis and C. rhizosoleniae are the only marine cyanobacteria with a terminal heterocyst strongly suggests a relationship with their symbiotic status. However the heterocyst differentiation and the mechanism behind the formation of just one terminal N2 fixing cell is still unknown. A recent study on Anabaena discovered NtcA and HetR as the main transcriptional factors responsible for the intercalary heterocyst differentiation (Flores et al., 2018). Hence, it would be interesting to investigate if the same factors are also involved in the heterocyst differentiation in Richelia/Calothrix and which gene regulation determines the formation of just one terminal cell. Prerequisite for such approach is the cyanobacteria persistence in culture, and, to date, the only symbiotic N2-fixing cyanobacteria successfully isolated is Calothrix rhizosoleniae SC01 (Foster et al., 2010). Nonetheless, through the heterologous expression of heterocyst differentiation genes, e.g. hetR, in Anabaena mutants it would also be possible to investigate the expression of R. intracellularis. Additionally, comparing the level of expression of cyanobacterial heterocyst differentiation genes with the host transcriptome (note. also the diatom-hosts evaded cultures) could represent a means to investigate if this process is somewhat regulated by the diatom-host or, perhaps, by environmental factors (e.g. light).

Can we reassign the species designation to C. rhizosoleniae? And how micro-diverse are the strains within the het-group?

The sequences generated in paper I and the phylogenetic clustering of the symbiotic cyanobacteria was consistent with previous studies (Janson et al., 1999; Foster and Zehr, 2006). Nevertheless, as pointed out earlier by Gómez et al. (2005), there was and there still is taxonomical confusion in the Richelia group. Besides the fact that R. intracellularis and C. rhizosoleniae share very similar morphologies and both seem to be the only marine symbiotic filamentous cyanobacteria with terminal heterocysts, C. rhizosoleniae does not phylogenetically cluster with any of the strains belonging to its genus (Fig. 17).

Fig. 17. Bayesian reconstruction of 51 cyanobacteria 16S rRNA sequences (1500 bp) showing the symbiotic C. rhizosoleniae (highlighted in light blue) clustering together with the Richelia sequences (highlighted in rose), and a part from the Calothrix cluster (highlighted in green). Branch support is reported as % probability. Scale bar=0.04; outgroup= Trichodesmium erythraeum IMS 101. Modified from paper I.

Indeed, its 16S rRNA sequence is 98% identical to R. intracellularis and just 90–92% to Calothrix spp. Thus, a taxonomical revision of the strain Calothrix rhizosoleniae should be considered. In addition, in paper I we highlight sequences with high similarity (98-100%) to ours from atypical environments. Given the ancient evolutionary background of Richelia/Calothrix strains, and the adaptation they likely undergone to survive in extreme environments (e.g. at polar regions), genetic micro-diversity among the symbiotic strains would be expected. A first attempt to verify the symbiotic heterocystous cyanobacteria micro-diversity has been performed using nifH environmental sequences (Fig. 18; Note. database curated by Dr. Carl T. Vigil Stenman).

Fig. 18. Bayesian reconstruction of the nifH nucleotide phylogenesis (362 bp) showing the micro-diversity among 92 environmental Richelia and Calothrix sequences. The reference sequences (or clusters) coming from the genomes are color coded (red=het-1; green=het-2, B; blue=het-3). Subtrees with distance < 0.042 have been collapsed, and number of sequences are reported in brackets. Branch support is reported as % probability. Outgroup=Trichodesmium erythraeum IMS 101. Scale bar=0.03. Phylogenetic reconstruction by Andrea Caputo.

Indeed, the phylogenetic reconstruction suggests more diversity than previously shown, although limited to the partial sequence of a single gene (362 bp). Hence, concatenating multiple genes and reconstructing larger portions of the genomes of the diatom-hosts as well would give a better estimate of the micro-diversity of these understudied symbioses.

How accurately can we estimate the DDAs absolute abundance? In paper II we summarized some of the biases that led to an underestimation of the DDAs, e.g. multiple pre-filtration steps employed during pan global surveys. An additional caveat of the DDAs quantification has been highlighted in paper III, i.e. cross-reactivity between het-1 and het-2 nifH oligonucleotides, specifically when qPCR assays run at 60ºC annealing temperature, both in fast and standard mode. Hence, in order to verify the abundance of Richelia/Calothrix quantified by nifH qPCR assays, a parallel set of primers/probes targeting another conservative gene, e.g. hetR, could provide a more accurate quantification of the symbionts. In addition, given that thus far the molecular quantification of the DDAs has been estimated exclusively based on the presence of the symbiont, the 18S rRNA and rbcL sequences obtained in this thesis could be used to design qPCR oligonucleotides specific for the diatom-hosts. Thus, simultaneous runs of symbiont and host primer/probes on environmental sample could estimate the DDAs actual abundance, and, indirectly used to estimate their contribution to the C and N biogeochemical cycles. However, with all qPCR based estimates it is important to recognize that these are relative abundances not absolute and caution should be used when referring to gene abundances as species or population abundances.

Does the symbiosis cell-division involve intermediate life-stages? In paper IV, the newly developed CARD-FISH assays showed the symbiotic partners in a free-living state, and likely the cyanobacteria in a different life stage (i.e. single-cell). Interestingly, several closely related strains to Richelia/Calothrix can form resting stages, (e.g. akinetes, important also as glycogen and proteins storage), and/or hormogonia, small motile filaments typical, for example, of most Nostoc and Calothrix (Flores and Herrero, 2010). Nevertheless, none of such morphotypes have been, so far, described for the marine symbiotic N2-fixing cyanobacteria. Previous studies occasionally reported observations of free-living symbionts (Sournia, 1970; Marumo and

Asaoka, 1974; Villareal, 1990; Gómez et al., 2005; Foster et al., 2007, 2011; White et al., 2007), yet in a filamentous state or with poor cell integrities. Thus, applying the ad hoc double CARD-FISH assay for DDAs developed in this thesis on samples from different oceans, and different environmental conditions, could estimate the magnitude of such phenomena on a larger scale. Alternatively, assuming the successful isolation and maintenance of DDAs into cultures, long-term experiments under different stressor(s) (e.g. temperature, salinity, pH) could be performed and the differentiation of the symbiosis into other morphotypes investigated. Moreover, the assay could also be a tool to examine if free-living symbionts represent a transition state prior to a new host infection (Gómez et al., 2005), or if they are capable to live independently from their host, as demonstrated for C. rhizosoleniae (Foster et al., 2010).

Acknowledgments During the last four years I have had the great opportunity of meeting incredible people, visiting remote places, sailing the oceans, and learn. It is beyond words to express how much I have learnt throughout my PhD, both as scientist and as person. And the credit for all of this belongs to my supervisor, Rachel Ann Foster. Thank you Rachel for the extraordinary experience that this four-year trip has represented for me and for my future.

Monika (Winder) and Ulla (Rasmussen), you have been supportive, understanding, caring and cheering since day 1. I have been extremely lucky to have you by my side. THANKS!

Peter (Hambäck), it is also for your support and your trust if today I am here, achieving this life goal that four years ago was as far as possible from me. Thank you for believing in me!

Chiara (Pennesi), an inspiration for all scientists, and especially for me. I will never thank you enough for being such a great mentor and friend since 2011. Grazie infinite Kia!

Anders (Löfgren), I am extremely glad I had the opportunity of meeting you and be your mentee. Thanks for all the insightful advice, and for listening to me all those hours! Ett stor tack Anders!

Edouard (Pesquet) and Ulla (Rasmussen), thanks for revising this thesis and for all the constructive feedback that dramatically improved my work. And thanks also to the Subject Head, Jonas (Gunnarson) for making the all defense process happening. Thanks!

Johan (Nylander), thank you so much for your advice on the phylogenetic work! I really appreciate the time and effort you have put in it. Thanks!

Massimo and Mercedes, in such short time you have been truly incredible friends. I know I can always count on you as you can count on me! Thanks for being so amazing! Os quiero mucho!

Marcus and Sofie, thanks for sharing your knowledge with me, for all the good times we had together, and for supporting each other unconditionally. It meant incredibly much to me! Miriam and Philip, thanks for being such smart and joyful students, I learnt so much from you too, Thanks! Lastly I want to thank whom has shortly being in the lab but left a special memory, Lotta, Theo, Narin, and Andreas, thank you guys!

Denis, Lina, Sandra, and Paul, you are the ones I called “The originals”. I could list a million reasons why you have been so amazingly important for me from the very beginning, but I am sure you already know. Thanks just for being You!

The Spanish mafia & co. (Note. I have avoided to use our WhatsApp group name) Sara, Sonia, David, Dimi, and Jörg, thank you for the fun time, the dances, and the hilarious (always boarder line) conversations at lunch.

Pil and Mariana, Tovetorp would not have been the same without you! Thanks also to all the students that made my teaching experience so overflowing! Lastly many thanks to the course leader Niklas Jenz which gave me this incredible opportunity (and thanks for the super fun volleyball games!).

Tom, Elina, Diana, DD, Marco, Alfred, Alma, Konrad, Peter, Aleksandra, Van, thanks to all of you for being, in different unique ways, very special to me.

My previous and current office mates, Jens and Laura, thanks for the nice chats and for keeping (or at least trying to) alive our plants!

The Equality Group members, thanks for valuing my ideas, for achieving goals together and for being such an inspiration for me. Thank you!

I could keep listing all the great people I have met during the last four years, but I would like to thank altogether the previous and current employees and students of Botan and DEEP. THANKS!

Mar, Kyle, Andy, Tasha, Tara, Kendra, the TMPs Nina and Yubin, and all the scientists and crew of the OUTPACE, HOT#290, and TriCoLim expeditions. Sailing with you has been one of the best experiences of my life!

Finally, the biggest thanks goes to my Italian-Swedish family. You have been the reason why I have never given up. You have been my safe harbor, the light in the darkest days, and my very first supporters, no matter what. Grazie mamma, papá e Giovi per esserci stati sempre! Thank you Amo for being by my side every single day and for being the best supporter ever! I feel extremely lucky!

References Agnese D.J.D. and Clark D.L. (1994) Siliceous Microfossils from the Warm Late Cretaceous and Early Cenozoic Arctic Ocean. J Paleontol 68: 31–47. Amann R.I., Binde, B.J., Olson R.J., Chisholm S.W., Devereux R., and Stahl D.A. (1990) Combination of 16S rRNA-targeted oligonucleotide probes with flow cytometry for analyzing mixed microbial populations. Appl Environ Microbiol 56: 1919–1925. Amin S. A., Parker M.S., and Armbrust E.V. (2012) Interactions between diatoms and bacteria. Microbiol Mol Biol Rev 76(3): 667–684. Bale N.J., Villareal T.A., Hopmans E.C., Brussaard C.P., Besseling M., Dorhout D., et al. (2018) C5 glycolipids of heterocystous cyanobacteria track symbiont abundance in the diatom Hemiaulus hauckii across the tropical North Atlantic. Biogeosciences 15(4): 1229–1241. Bauersachs T., Speelman E.N., Hopmans E.C., Reichart G.J., Schouten S., and Damste J.S.S. (2010) Fossilized glycolipids reveal past oceanic N2 fixation by heterocystous cyanobacteria Proc. Natl. Acad. Sci. U.S.A. 107(45): 19190–19194. Bench S.R., Ilikchyan I.N., Tripp H.J., and Zehr J.P. (2011) Two strains of Crocosphaera watsonii with highly conserved genomes are distinguished by strain-specific features. Front Microbiol 2 (261): 1–13 Benson D.R., and Silvester WB. (1993) Biology of Frankia strains, actinomycete symbionts of actinorhizal plants. Microbiol Rev 57 (2): 293–319. Bentzon-Tilia M., Severin I., Hansen L.H., and Riemann L. (2015) Genomics and ecophysiology of heterotrophic nitrogen-fixing bacteria isolated from estuarine surface water. MBio 6:e00929. Bergman B., Sandh G., Lin S., Larsson J., and Carpenter E.J. (2013) Trichodesmium– a widespread marine cyanobacterium with unusual nitrogen fixation properties. FEMS Microbiol rev 37 (3): 286–302. Berman-Frank I., Cullen J., Hareli Y., Sherrell R., and Falkowski P.G. (2001) Iron availability, cellular iron quotas, and nitrogen fixation in Trichodesmium. Limnol Oceanogr 46: 1249–1277. Berman-Frank I., Lundgren P., and Falkowski, P. (2003) Nitrogen fixation and photosynthetic oxygen evolution in cyanobacteria. Res Microbiol 154 (3): 157– 164. Berthelot H., Benavides M., Moisander P.H., Grosso O., and Bonnet S. (2017) High‐ nitrogen fixation rates in the particulate and dissolved pools in the Western Tropical Pacific (Solomon and Bismarck Seas). Geophys Res Lett 44 (16): 8414– 8423. Bombar D., Heller P., Sanchez-Baracaldo P., Carter B.J., and Zehr J.P. (2014) Comparative genomics reveals surprising divergence of two closely related strains of uncultivated UCYN-A cyanobacteria. ISME J 8: 2530–2542. 67

Bombar D., Moisander P.H., Dippner J.W., Foster R.A., Voss M., Karfeld B., et al. (2011) Distribution of diazotrophic microorganisms and nifH gene expression in the Mekong River plume during intermonsoon. Mar Ecol Prog Ser 424: 39–52. Bombar D., Paerl R.W., and Riemann L. (2016) Marine Non-Cyanobacterial Diazotrophs: Moving beyond Molecular Detection. Trends Microbiol 24: 916– 927. Bonnet S., Webb E.A., Panzeca C., Karl D.M., Capone D.G., and Wilhelmy S.A.S. (2010) Vitamin B12 excretion by cultures of the marine cyanobacteria Crocosphaera and Synechococcus. Limnol Oceanog 55(5): 1959–1964. Bothe H., Tripp H.J., and Zehr J.P. (2010) Unicellular cyanobacteria with a new mode of life: the lack of photosynthetic oxygen evolution allows nitrogen fixation to proceed. Arch Microbiol 192 (10): 783–790. Bown P.R., Lees J.A., and Young J.R. (2004) Calcareous nannoplankton evolution and diversity through time. In: Coccolithophores. Springer, Berlin, Heidelberg, pp. 481-508. Böttjer D., Dore J.E., Karl D.M., Letelier R.M., Mahaffey C., Wilson S.T., et al. (2017). Temporal variability of nitrogen fixation and particulate nitrogen export at Station ALOHA. Limnol Oceanogr 62(1): 200–216. Breitbarth E., Oschlies A., and LaRoche J. (2007) Physiological constraints on the global distribution of Trichodesmium - effect of temperature on diazotrophy. Biogeosciences 4(1): 53–61. Capone D.G., Burns J.A., Montoya J.P., Subramaniam A., Mahaffey C., Gunderson T., et al. (2005) Nitrogen fixation by Trichodesmium spp.: An important source of new nitrogen to the tropical and subtropical North Atlantic Ocean. Global Biogeochem Cycles 19 (2): 1–17. Capone D.G., Zehr J., Paerl H., Bergman B., and Carpenter E.J. (1997) Trichodesmium: A globally significant marine cyanobacterium. Science 276: 1221– 1229. Caputo A., Stenegren M., Pernice M.C., and Foster R.A. (2018) A short comparison of two marine diazotrophic symbioses highlights an un-quantified disparity. Front Mar Sci 5 (2): 1–8. Carpenter E.J. (1983) Nitrogen fixation by marine Oscillatoria (Trichodesmium) in the world's oceans. In: Carpenter E.J. and Capone D.G. (Eds.), Nitrogen in the Marine Environment. Academic Press, New York. Carpenter E.J. and Foster R.A. (2002) Marine Cyanobacterial Symbioses. In: Rai A.N., Bergman B., and Rasmussen U. (eds) Cyanobacteria in Symbiosis. Kluwer Academic Publishers, Dordrecht, pp. 11–17. Carpenter E.J. and Janson S. (2000) Intracellular cyanobacterial symbionts in the marine diatom Climacodium frauenfeldianum (Bacillariophyceae). J Phycol 36: 540–544.

Carpenter E.J. and McCarthy J.J. (1975) Nitrogen fixation and uptake of combined nitrogenous nutrients by Oscillatoria (Trichodesmium) thiebautii in the western Sargasso Sea. Limn Oceanog 20: 389–401. Carpenter E.J., Montoya J.P., Burns J., Mulholland M.R., Subramaniam A., and Capone D.G. (1999) Extensive bloom of a N2-fixing diatom/cyanobacterial association in the tropical Atlantic Ocean. Mar Ecol Prog Ser 185: 273–283. Carpenter E.J., A. Subramaniam A., and Capone D.G. (2004) Biomass and primary productivity of the cyanobacterium, Trichodesmium spp., in the southwestern tropical N. Atlantic Ocean. Deep Sea Res 51 (1): 173–203. Chappell P.D., Moffett J.W., Hynes A.M., and Webb E.A. (2012) Molecular evidence of iron limitation and availability in the global diazotroph Trichodesmium. ISME J 6(9): 1728–1739. Chavez F.P. and Toggweiler J.R. (1995) Physical estimates of global new production: the upwelling contribution. In: Upwelling in the ocean: modern processes and ancient records, Summerhayes CP et al. (eds). John Wiley and Sons, Chichester, 313–320. Chen Y.L. (2005) Spatial and seasonal variations of nitrate-based new production and primary production in the South China Sea. Deep Sea Res Part I Oceanogr Res Pap 52: 319–340. Chen Y.L., Chen H.Y., Karl D.M., Takahashi M. (2004). Nitrogen modulates phytoplankton growth in spring in the South China Sea. Cont Shelf Res 24: 527– 541. Church M.J., Jenkins B.D., Karl D.M., and Zehr J.P. (2005) Vertical distributions of nitrogen-fixing phylotypes at Stn Aloha in the oligotrophic North Pacific Ocean, Aquat Microb Ecol 38: 3–14. Cornejo-Castillo F.M., Cabello A.M., Salazar G., Sa P., Lima-mendez G., Hingamp P., et al. (2016) Cyanobacterial symbionts diverged in the late Cretaceous towards lineage-specific nitrogen fixation factories in single-celled phytoplankton. Nat Commun 7: 1–9. Davies A. and Kemp A.E. (2016) Late Cretaceous seasonal palaeoclimatology and diatom palaeoecology from laminated sediments. Cretac Res 65: 82–111. Decelle J., Colin S., and Foster R.A. (2015) Photosymbiosis in Marine Planktonic Protists. In: Marine Protists. Springer Japan, Tokyo, 465–500. Doherty H.M. and Adams D.G. (1995) Cloning and sequence of ftsZ and flanking regions from the cyanobacterium Anabaena PCC 7120. Gene 163(1): 93–96. Dron A., Rabouille S., Claquin P., Le Roy B., Talec A., and Sciandra A. (2012) Light– dark (12: 12) cycle of carbon and nitrogen metabolism in Crocosphaera watsonii WH8501: relation to the cell cycle. Environ Microbiol 14 (4): 967–981. Duarte C. M. (2015) Seafaring in the 21st century: the Malaspina 2010 Circumnavigation Expedition. Limnol Oceanogr Bull 24(1): 11–14.

Dugdale R.C. and Goering J.J. (1967) Uptake of new and regenerated forms of nitrogen in primary productivity. Limnol Oceanogr 12 (2): 196-206. Dugdale R.C., Menzel D.W., and Ryther J. H. (1961) Nitrogen fixation in the Sargasso Sea. Deep-Sea Res 7: 297–300. Dunthorn M., Klier J., Bunge J., and Stoeck T. (2012) Comparing the hyper‐variable V4 and V9 regions of the small subunit rDNA for assessment of ciliate environmental diversity. J Eukaryot Microbiol 59(2): 185–187. Dyhrman S.T., Chappell P.D., Haley S.T., Moffett J.W., Orchard E.D., Waterbury J.B., et al. (2006) Phosphonate utilization by the globally important marine diazotroph Trichodesmium. Nature 439 (7072): 68–71. Dyhrman S.T. and Haley S.T. (2006) Phosphorus scavenging in the unicellular marine diazotroph Crocosphaera watsonii. Appl Environ Microbiol 72(2): 1452–1458. Dyhrman S.T., Webb E.A., Anderson D.M., Moffett J.W., and Waterbury J.B. (2002) Cell-specific detection of phosphorus stress in Trichodesmium from the western North Atlantic. Limnol Oceanogr 47:1832–1836. Embley T.M., Finlay B.J., Thomas R.H., and Dyal, P.L. (1992) The use of rRNA sequences and fluorescent probes to investigate the phylogenetic positions of the anaerobic ciliate Metopus palaeformis and its archaeobacterial endosymbiont. J Gen Microbiol 138: 1479–1487. Farnelid H., Bentzon-Tilia M., Andersson A.F., Bertilsson S., Jost G., Labrenz M., et al. (2013). Active nitrogen-fixing heterotrophic bacteria at and below the chemocline of the central Baltic Sea. ISME J 7, 1413–1423. Farnelid H., Turk-Kubo K., Muñoz-Marín M., Zehr J. (2016) New insights into the ecology of the globally significant uncultured nitrogen-fixing symbiont UCYN-A. Aquat Microb Ecol 77: 125–138. Femino A.M., Fay F.S., Fogarty K., and Singer R.H. (1998) Visualization of single RNA transcripts in situ. Science 280: 585–590. Ferrario, M.E., Villafane, V., Helbling, W., and Hansen, O.H. (1995) The occurrence of the symbiont Richelia in Rhizosolenia and Hemiaulus in North Pacific. Rev Bras Bio 55: 439–443. Field C.B., Behrenfeld M.J., Randerson J.T., and Falkowski P. (1998) Primary production of the biosphere: integrating terrestrial and oceanic components. Science 281(5374): 237–240. Flores E. and Herrero A. (2010) Compartmentalized function through cell differentiation in filamentous cyanobacteria. Nat Rev Microbiol 8(1): 39. Flores E., Picossi S., Valladares A., and Herrero A. (2018) Transcriptional regulation of development in heterocyst-forming cyanobacteria. Biochim Biophys Acta Gene Regul Mech.

Follett C.L., Dutkiewicz S., Karl D.M., Inomura K., and Follows M.J. (2018) Seasonal resource conditions favor a summertime increase in North Pacific diatom– diazotroph associations. ISME J 12: 1543–1557. Foster R.A., Carpenter E.J., and Bergman B. (2006) Unicellular cyanobionts in open ocean dinoflagellates, radiolarians, and tintinnids: Ultrastructural characterization and immuno-localization of phycoerythrin and nitrogenase. J Phycol 42: 453–463. Foster R.A., Goebel N.L., and Zehr J.P. (2010) Isolation of Calothrix rhizosoleniae (cyanobacteria) strain SC01 from Chaetoceros (Bacillariophyta) spp. diatoms of the subtropical North Pacific Ocean. J Phycol 46: 1028–1037. Foster R.A., Kuypers M.M.M., Vagner T., Paerl R.W., Musat N., and Zehr J.P. (2011) Nitrogen fixation and transfer in open ocean diatom-cyanobacterial symbioses. ISME J 5: 1484–1493. Foster R.A., Subramaniam A., Mahaffey C., Carpenter E.J., Capone D.G., and Zehr J.P. (2007) Influence of the Amazon River plume on distributions of free-living and symbiotic cyanobacteria in the western tropical north Atlantic Ocean. Limnol Oceanogr 52: 517–532. Foster R.A., Subramaniam A., and Zehr J.P. (2009) Distribution and activity of diazotrophs in the Eastern Equatorial Atlantic. Environ Microbiol 11(4): 741–750.

Foster R.A., Sztejrenszus S., and Kuypers M.M. (2013) Measuring carbon and N2 fixation in field populations of colonial and free‐living unicellular cyanobacteria using nanometer‐scale secondary ion mass spectrometry. J Phycol 49 (3): 502– 516. Foster R.A. and O’Mullan G.D. (2008) Nitrogen-Fixing and Nitrifying Symbioses in the Marine Environment. Nitrogen Mar Environ 1197–1218. Foster R.A and Zehr J.P. (2006) Characterization of diatom-cyanobacteria symbioses on the basis of nifH, hetR and 16S rRNA sequences. Environ Microbiol 8: 1913– 1925. Frank A.B. (1877) Über die biologischen Verhältnisse des Thallus einiger Krustflechten. Cohn Beitr Biol Pflanz 2:123–200

Gallon J.R. (1992) Reconciling the incompatible: N2 fixation and O2. New Phytol 122 (4): 571–609. Giovannoni S.J., Thrash J.C., and Temperton B. (2014) Implications of streamlining theory for microbial ecology. ISME J 8(8): 1553–1565. Goldman J.C. (1993) Potential role of large oceanic diatoms in new primary production. Deep Sea Res Part 1 Oceanogr Res Pap 40(1): 159–168. Gómez F., Furuya K.E.N., Takeda S., and Go F. (2005) Distribution of the cyanobacterium Richelia intracellularis as an epiphyte of the diatom Chaetoceros compressus in the western Pacific Ocean. J Plankton Res 27: 323–330. Grace J.B. 2006. Structural equation modeling and natural systems. Cambridge University Press, Cambridge, UK. 71

Grace J.B., Anderson T.M., Olff H., and Scheiner S.M. (2010). On the specification of structural equation models for ecological systems. Ecol Monogr 80(1): 67–87. Griese M., Lange C., and Soppa J. (2011) Ploidy in cyanobacteria. FEMS Microbiol. Lett. 323: 124–131. Grosse J., Bombar D,, Doan H,N,, Nguyen L,N,, andVoss M. (2010) The Mekong River plume fuels nitrogen fixation and determines phytoplankton species distribution in the South China Sea during low- and high-discharge season. Limnol Oceanogr 55: 1668–1680. Großkopf T., Mohr W., Baustian T., Schunck H., Gill D., Kuypers M.M. et al. (2012) Doubling of marine dinitrogen-fixation rates based on direct measurements. Nature 488 (7411): 361–364. Gruber N. (2016) Elusive marine nitrogen fixation. Proc Natl Acad Sci 113 (16): 4246–4248. Guillard R.R.L. (1975) Culture of phytoplankton for feeding marine invertebrates. In: Smith W.L. and Chanley M.H (Eds.) Culture of Marine Invertebrate Animals. Plenum Press, New York, USA, pp 26-60 Guillou L., Bachar D., Audic S., Bass D., Berney C., Bittner L., et al. (2012) The Protist Ribosomal Reference database (PR2): a catalog of unicellular eukaryote small sub-unit rRNA sequences with curated taxonomy. Nucleic Acids Res 41(1): 597-604. Guo L., Sui Z., Zhang S., Ren Y., and Liu Y. (2015) Comparison of potential diatom ‘barcode’ genes (the 18S rRNA gene and ITS, COI, rbcL) and their effectiveness in discriminating and determining species taxonomy in the Bacillariophyta. Int J Syst Evol Microbiol 65(4): 1369–1380. Hadziavdic K., Lekang K., Lanzen A., Jonassen I., Thompson E.M., and Troedsson C. (2014) Characterization of the 18S rRNA gene for designing universal eukaryote specific primers. PloS one 9(2): e87624. Hagino K., Onuma R., Kawachi M., and Horiguchi T. (2013) Discovery of an endosymbiotic nitrogen-fixing cyanobacterium UCYN-A in Braarudosphaera bigelowii (Prymnesiophyceae). PLoS ONE 8 (12): 1–11. Haines K.C. and Guillard R.R. (1974) growth of vitamin B12‐requiring marine diatoms in mixed laboratory cultures with vitamin B12‐producing marine bacteria. J Phycol 10(3):245–252. Halm H., Lam P., Ferdelman T.G., Lavik G., Dittmar T., LaRoche J., et al. (2012). Heterotrophic organisms dominate nitrogen fixation in the South Pacific Gyre. ISME J 6: 1238–1249. Hamsher S.E., Evans K.M., Mann D.G., Poulíčková A., and Saunders G.W. (2011). Barcoding diatoms: exploring alternatives to COI-5P. Protist 162(3): 405–422.

Harke M.J., Frischkorn K.R., Haley S.T., Aylward F.O., Zehr J.P., and Dyhrman S.T. (2018) Periodic and coordinated gene expression between a diazotroph and its diatom host. The ISME J: 1–14. Heinbokel J.F. (1986) occurrence of Richelia Intacellularis (cyanophyta) within the diatoms Hemiaulus haukii adn H. membranaceus off Hawaii. J Phycol 22(3): 399– 403. Hilton J.A. (2014) Ecology and Evolution of Diatom-Associated Cyanobacteria through genetic analyses. PhD thesis, University of California, Santa Cruz, CA USA. Hilton J.A., Foster R.A., Tripp H.J., Carter B.J., Zehr J.P., and Villareal T.A. (2013) Genomic deletions disrupt nitrogen metabolism pathways of a cyanobacterial diatom symbiont. Nat Commun 4 (1767): 1-7. Hynes A.M., Chappell P.D., Dyhrman S.T., Doney S.C., and Webb E.A. (2009). Cross‐basin comparison of phosphorus stress and nitrogen fixation in Trichodesmium. Limnol Oceanogr 54 (5): 1438–1448. Ishaq S.L. and Wright A.D.G. (2014) Design and validation of four new primers for next-generation sequencing to target the 18S rRNA gene of gastrointestinal ciliate protozoa. Appl Environ Microbiol AEM 01644: 1–28. Jabir T., Dhanya V., Jesmi Y., Prabhakaran M.P., Saravanane N., Gupta G.V.M., and Hatha A.A.M. (2013) Occurrence and distribution of a diatom-diazotrophic cyanobacteria association during a Trichodesmium bloom in the southeastern Arabian Sea. Int J Oceanogr 2013: 1-6. Janson S., Rai A.N., and Bergman B. (1995) Intracellular Cyanobiont Richelia intracellularis - Ultrastructure and Immuno-Localization of Phycoerythrin, Nitrogenase, Rubisco and Glutamine-Synthetase. Mar Biol 124: 1–8. Janson S., Woulters J., Bergman B., and Carpenter E.J. (1999) Host specificity in the Richelia-diatom symbiosis revealed by hetR gene sequence analysis. Environ Microbiol 1: 431–438. Karl D.M., Church M.J., Dore J.E., Letelier R.M., Mahaffey C. (2012) Predictable and efficient carbon sequestration in the North Pacific Ocean supported by symbiotic nitrogen fixation. Pnas 109: 1842–1849. Karl D., Letelier R., Tupas L., Dore J., Christian J., and Hebel D. (1997) The role of nitrogen fixation in biogeochemical cycling in the subtropical North Pacific Ocean. Nature 388: 533–538. Karl D., Michaels A., Bergman B., Capone D., Carpenter E., Letelier R., et al. (2002) Dinitrogen fixation in the world ’s oceans. Biogeochemistry 57/58: 47–98. Karsten G. (1907) Das indische Phytoplankton nach dem Material der deutschen Tiefsee-Expedition 1898-I899. Wiss Ergebn dt Tiefsee-Exped "Valdivia" 2: 221– 548.

Kashiyama Y., Ogawa N.O., Kuroda J., and Shiro M. (2008). Diazotrophic cyanobacteria as the major photoautotrophs during mid-Cretaceous oceanic anoxic events: Nitrogen and carbon isotopic evidence from sedimentary porphyrin. Org Geochem 39: 532–549. Ke R., Mignardi M., Pacureanu A., Svedlund J., Botling J., Wählby C., et al. (2013) In situ sequencing for RNA analysis in preserved tissue and cells. Nat Methods 10: 1–6. Keeling P.J., Burki F., Wilcox H.M., Allam B., Allen E.E., Amaral-Zettler L.A., et al. (2014) The Marine Microbial Eukaryote Transcriptome Sequencing Project (MMETSP): illuminating the functional diversity of eukaryotic life in the oceans through transcriptome sequencing. PLoS biology 12(6): 1–6. Kemp A.E., Pearce R.B., Koizumi I., Pike J., and Rance S.J. (1999) The role of mat- forming diatoms in the formation of Mediterranean sapropels. Nature 398(6722): 57–61. Kemp A.E. and Villareal T.A. (2013) High diatom production and export in stratified waters–A potential negative feedback to global warming. Prog Oceanogr 119: 4– 23. Kermarrec L., Franc A., Rimet F., Chaumeil P., Humbert J.F., and Bouchez A. (2013). Next‐generation sequencing to inventory taxonomic diversity in eukaryotic communities: a test for freshwater diatoms. Mol Ecol Resour 13(4): 607–619. Kim J. and Rees D.C. (1992) Structural models for the metal centers in the nitrogenase molybdenum-iron protein. Science 257 (5077): 1677–1682. Kitajima S., Furuya K., Hashihama F., Takeda S., and Kanda J. (2009). Latitudinal distribution of diazotrophs and their nitrogen fixation in the tropical and subtropical western North Pacific. Limnol Oceanogr 54(2): 537–547. Knapke E.M. (2012) Influence of the Mississippi River Plume on Diazotroph Distributions in the Northern Gulf of Mexico During Summer 2011. Master Thesis (The University of Texas at Austin, USA). Kneip C., Voss C., Lockhart P.J., and Maier U.G. (2008) The cyanobacterial endosymbiont of the unicellular algae Rhopalodia gibba shows reductive genome evolution. BMC Evol Biol 8(30): 1–16. Krupke A., Lavik G., Halm H., Fuchs B.M., Amann R.I., and Kuypers M.M. (2014) Distribution of a consortium between unicellular algae and the N2 fixing cyanobacterium UCYN‐A in the North Atlantic Ocean. Environ Microbiol 16(10): 3153–3167. Krupke A., Musat N., LaRoche J., Mohr W., Fuchs B.M., Amann R.I., et al. (2013) In situ identification and N2 and C fixation rates of uncultivated cyanobacteria populations. Syst Appl Microbiol 36: 259–271. Kulkarni V.V., Chitari R.R., Narale D.D., Patil J.S., and Anil A.C. (2010) Occurrence of cyanobacteria-diatom symbiosis in the Bay of Bengal: implications in biogeochemistry. Curr Sci 99(6): 736-737. 74

Kumar K., Mella-Herrera R.A., and Golden J.W. (2010) Cyanobacterial heterocysts. Cold Spring Harb Perspect Biol 2: 1–20.

Kuypers M.M.M., Breugel Y Van, Schouten S., Burg D., and Erba E. (2004) N2-fixing cyanobacteria supplied nutrient N for Cretaceous oceanic anoxic events. Geology 32: 853–856. Laamanen M.J. (1997) Environmental factors affecting the occurrence of different morphological forms of cyanoprokaryotes in the northern Baltic Sea. J Plankton Res 19(10): 1385–1403. Lane D.J. (1991) 16S/23S rRNA sequencing. In: Stackebrandt E. and Goodfellow M. (eds). Nucleic acid techniques in bacterial systematics. New York: John Wiley and Sons, pp. 115-175. Langlois R.J., LaRoche J. and Raab P.A. (2005) Diazotrophic diversity and distribution in the tropical and subtropical Atlantic Ocean. Appl Environ Microbiol 71(12): 7910–7919. Larsson C., Grundberg I., Söderberg O., and Nilsson M. (2010) In situ detection and genotyping of individual mRNA molecules. Nat Methods 7: 395–397. Lemmermann E. (1905) Die algenflora der sandwich-inseln. ergebnisse einer reise nach dem Pacific. H. Schauinsland 1896/97. Bot Jahrbuch Syst Pflanzengesch Pflanzengeogr 34: 607–663. Lepage T., Bryant D., and Lartillot N. (2007) A General Comparison of Relaxed Molecular Clock Models. Mol Biol Evol 24: 2669–2680. Li N., Yu S.X., Wang Y.C., et al. (2016) Diversity of phototrophic phytoplankton in Northern South China Sea indicated by rbcL analysis. J Appl Phycol 28(2): 773– 781. Loescher C.R., Grosskopf T., Desai F.D., Gill D., Schunck H., Croot P.L., et al. (2014). Facets of diazotrophy in the oxygen minimum zone waters off Peru. ISME J 8: 2180–2192. Luddington I.A., Kaczmarska I., Lovejoy C. (2012) Distance and character based evaluation of the V4 region of the 18S rRNA gene for the identification of diatoms (Bacillariophyceae). PLoS ONE 7:e45664. Luo Y.W., Doney S.C., Anderson L.A., Benavides M., Berman-Frank I., Bode A., et al. (2012). Database of Diazotrophs in Global Ocean: Abundance, Biomass, and Nitrogen Fixation Rates. Earth Syst Sci Data 4: 47–73. Luo Y.W., Lima I.D., Karl D.M., Deutsch C.A., and Doney S.C. (2014) Data-based assessment of environmental controls on global marine nitrogen fixation. Biogeosciences 11 (3): 691–708. Madhu N.V., Paul M., Ullas N., Ashwini R., and Rehitha T.V. (2013) Occurrence of cyanobacteria (Richelia intracellularis)-diatom (Rhizosolenia hebetata) consortium in the Palk Bay, southeast coast of India. Indian J Mar Sci 42: 453– 457.

MacGillivary M.L., and Kaczmarska I. (2011) Survey of the efficacy of a short fragment of the rbc L gene as a supplemental DNA barcode for diatoms. J Eukaryot Microbiol 58(6): 529–536. Mague T.H., Weare N.M., and Holm-Hansen O. (1974) Nitrogen fixation in the North Pacific Ocean. Mar Biol 24: 109–119. Manjumol C.C., Libini C.L., Idu K. A., Mohamed K.S., and Kripa V. (2018) Occurrence of Diatom–Diazotrophic association in the coastal surface waters of south Andaman, India. Symbiosis: 1-10. Martínez-Pérez C., Mohr W., Löscher C.R., Dekaezemacker J., Littmann S., Yilmaz P., et al. (2016) The small unicellular diazotrophic symbiont, UCYN-A, is a key player in the marine nitrogen cycle. Nat Microbiol 1 (16163): 1–7. Marumo R. and Asaoka O. (1974) Distribution of pelagic blue-green algae in the North Pacific Ocean. J Oceanogr Soc Japan 30: 77–85. McCarthy J.J. and Carpenter E.J. (1983) Nitrogen cycling in near-surface waters of the open ocean In: Nitrogen in the Marine Environment, E.J. Carpenter and D.G. Capone (eds). Academic Press, New York, 487–512. Miller P.E. and Scholin C.A. (1996) Identification of cultured Pseudo‐nitzschia (Bacillariophyceae) using species‐specific LSU rRNA‐targeted fluorescent probes. J Phycol 32 (4): 646–655. Mills M.M., Ridame C., Davey M., La Roche J., and Geider R.J. (2004) Iron and phosphorus co-limit nitrogen fixation in the eastern tropical North Atlantic. Nature 429 (6989): 292–294. Moisander P.H., Beinart R.A., Hewson I., White A.E., Johnson K.S., Carlson C.A., et al. (2010) Unicellular cyanobacterial distributions broaden the oceanic N2 fixation domain. Science 327 (5972): 1512–1514. Moisander P.H., Benavides M., Bonnet S., Berman-Frank I., White A.E., and Riemann L. (2017) Chasing after non-cyanobacterial nitrogen fixation in marine pelagic environments. Frontiers Microbiol 8 (1736):1–8. Moisander P.H., Zhang R., Boyle E.A., Hewson I., Montoya J.P., and Zehr J.P. (2012) Analogous nutrient limitations in unicellular diazotrophs and Prochlorococcus in the South Pacific Ocean. ISME J 6: 733–744. Montoya J.P., Holl C.M., Zehr J.P., Hansen A., Villareal T., and Capone D.G. (2004) High rates of N2-fixation by unicellular diazotrophs in the oligotrophic Pacific. Nature 430: 1027–1031. Moore C.M., Mills M.M., Achterberg E.P., Geider R.J., LaRoche J., Lucas M., et al. (2009) Large-scale distribution of Atlantic nitrogen fixation controlled by iron availability. Nat Geosci 2(12): 867–871. Moran N.A. (2003) Tracing the evolution of gene loss in obligate bacterial symbionts. Curr Opin Microbiol 6: 512–518.

Mueller F., Senecal A., Tantale K., Marie-Nelly H., Ly N., Collin O., et al. (2013) FISH-quant: automatic counting of transcripts in 3D FISH images. Nat Methods 10: 277–278. Nakayama T., Ikegami Y., Nakayama T., Ishida K., Inagaki Y., and Inouye I. (2011) Spheroid bodies in rhopalodiacean diatoms were derived from a single endosymbiotic cyanobacterium. J Plant Res 124: 93–97. Nakayama T. and Inagaki Y. (2016) Cyanobacterial genes in the nuclear genome of a diatom bearing N2-fixing cyanobacterial endosymbionts: potential factors involved in the host-endosymbiont partnership. Protistology 10(2). Nakayama T. and Inagaki Y. (2017) Genomic divergence within non-photosynthetic cyanobacterial endosymbionts in rhopalodiacean diatoms. Sci Rep 7(1): 13075. Nakayama T., Kamikawa R., Tanifuji G., Kashiyama Y., Ohkouchi N., Archibald J.M. et al. (2014) Complete genome of a nonphotosynthetic cyanobacterium in a diatom reveals recent adaptations to an intracellular lifestyle. Proc Natl Acad Sci USA 111(31): 11407–11412. Needoba J.A., Foster R.A., Sakamoto C., Zehr J.P., and Johnson K.S. (2007). Nitrogen fixation by unicellular diazotrophic cyanobacteria in the temperate oligotrophic North Pacific Ocean. Limnol Oceanog 52(4): 1317–1327. Neefs J.M., Van de Peer Y., De Rijk P., Chapelle S., and De Wachter R. (1993) Compilation of small ribosomal subunit RNA structures. Nucleic Acids Res 21(13): 3025–3049. Nelson D.M., Tréguer P., Brzezinski M.A., Leynaert A., and Quéguiner B. (1995) Production and dissolution of biogenic silica in the ocean: revised global estimates, comparison with regional data and relationship to biogenic sedimentation. Global Biogeochem Cycles 9(3): 359–372. Norris R.E. (1961) Observations on phytoplankton organisms collected on the NZOI Pacific cruise, September, 1958. NZ J Sci 4: 162–188.

Olson J.B., Steppe T.F., Litaker R.W., et al. (1998) N2-fixing microbial consortia associated with the ice cover of Lake Bonney, Antarctica. Microb Ecol 36: 231– 238. Orcutt K.M., Lipschultz F., Gundersen K., Arimoto R., Michaels A.F., Knap A.H., et al. (2001). A seasonal study of the significance of N2 fixation by Trichodesmium spp. at the Bermuda Atlantic Time-series Study (BATS) site. Deep-Sea Res. Pt. II 48: 1583–1608. Ostenfeld C.H. and Schmidt J. (1901) Plankton fra det Rode Hav og Adenbugten (Plankton from the Red Sea and the Gulf of Aden). Copenhagen. Padmakumar K.B., Menon N.R., and Sanjeevan V.N. (2010) Occurrence of endosymbiont Richelia intracellularis (Cyanophyta) within the diatom Rhizosolenia hebetata in Northern Arabian Sea. Int J Biodivers Conserv 2: 70–74.

Papaefthimiou D., Van Hove C., Lejeune A., Rasmussen U., and Wilmotte A. (2008) Diversity and host specificity of Azolla cyanobionts. J Phycol 44(1): 60–70. Paul J.H., Alfreider A., and Wawrik B. (2000) Micro-and macrodiversity in rbcL sequences in ambient phytoplankton populations from the southeastern Gulf of Mexico. Mar Ecol Prog Ser: 9–18. Pernice M.C., Logares R., Guillou L., and Massana R. (2013) General patterns of diversity in major marine microeukaryote lineages. PLoS One 8(2): e57170. Pernthaler A and Amann R. (2004). Simultaneous fluorescence in situ hybridization of mRNA and rRNA in environmental bacteria. Appl Environ Microbiol 70(9): 5426–5433. Pernthaler A., Pernthaler J. and Amann R. (2004) Sensitive multi-color fluorescence in situ hybridization for the identification of environmental microorganisms. Mol Microb Ecol Man 3: 2613–2627. Postgate J.R. (1982) Biological nitrogen fixation: fundamentals. Phil Trans R Soc Lond B 296 (1082): 375–385. Prechtl J., Kneip C., Lockhart P., Wenderoth K., and Maier U.G. (2004) Intracellular spheroid bodies of Rhopalodia gibba have nitrogen-fixing apparatus of cyanobacterial origin. Mol Biol Evol 21: 1477–1481. Raj A., van den Bogaard P., Rifkin S.A., van Oudenaarden A., Tyagi S., Bogaard P. Van Den, et al. (2008) Imaging individual mRNA molecules using multiple singly labeled probes. Nat. Methods 5: 877–879. Raj A., Peskin C.S., Tranchina D., Vargas D.Y., and Tyagi S. (2006) Stochastic mRNA synthesis in mammalian cells. PLoS Biol 4: 1707–1719. Ratten J.M., LaRoche J., Desai D.K., Shelley R.U., Landing W.M., Boyle E. et al. (2015) Sources of iron and phosphate affect the distribution of diazotrophs in the North Atlantic. Deep Sea Res Part 2 Top Stud Oceanogr 11: 332–341. Rau G.H., Arthur M.A., and Dean W.E. (1987) 15N/14N Variations in Cretaceous Atlantic Sedimentary Sequences: Implication for Past Changes in Marine Nitrogen Biogeochemistry. Earth Planet Sci Lett 82: 269–279. Riemann L., Farnelid H., and Steward G.F. (2010) Nitrogenase genes in noncyanobacterial plankton: prevalence, diversity, and regulation in marine waters. Aquat Microb Ecol 61: 235–247. Rippka R., Deruelles J., Waterbury J.B., Herdman M., Stanier R.Y. (1979) Generic assignments, strain histories and properties of pure cultures of Cyanobacteria. J Gen Microbiol 111: 1–61. Ritchie R.J. amd Gibson J. (1987) Permeability of ammonia, methylamine and ethylamine in the cyanobacterium, Synechococcus R-2 (Anacystis nidulans) PCC 7942. J Membr Biol 95(2): 131–142.

Ronquist F., Teslenko M., van der Mark P., et al. (2012) MrBayes 3.2: efficient Bayesian phylogenetic inference and model choice across a large model space. Syst Biol 61: 539–542 Round F.E., Crawford R.M., and Mann D.G. (1990) The diatoms. Biology and morphology of the genera. Cambridge University Press, Cambridge, UK. 744pp. Sabina J. and Leamon J.H. (2015) Bias in whole genome amplification: causes and considerations. In Whole Genome Amplification. Humana Press, New York, NY, pp. 15-41. Sabir J.S.M., Yu M., Ashworth M.P., Baeshen N.A., Baeshen M.N., Bahieldin A., et al. (2014) Conserved gene order and expanded inverted repeats characterize plastid genomes of Thalassiosirales. PLoS One 9(9): 1–9. Sachs J.P. and Repeta D.J. (1999) Oligotrophy and nitrogen fixation during eastern Mediterranean sapropel events. Science 286: 2485–2488. Sánchez-Baracaldo P., Raven J.A., Pisani D., et al. (2017) Early photosynthetic eukaryotes inhabited low-salinity habitats. Proc Natl Acad Sci USA 114(37): 7737–7745. Sargent, E. C., Hitchcock, A., Johansson, S. A., Langlois, R., Moore, C. M., LaRoche, J., et al. (2016) Evidence for polyploidy in the globally important diazotroph Trichodesmium. FEMS Microbiol Lett 363 (21): 1–7. Satinsky B.M., Smith C.B., Sharma S., Ward N.D., Krusche A.V., Richey J. E. et al. (2017). Patterns of Bacterial and Archaeal gene expression through the lower Amazon River. Front Mar Sci 4 (253): 1–13. Schouten S., Villareal T.A., Hopmans E.C., Mets A., Swanson K.M., and Sinninghe Damsté J.S. (2013) Endosymbiotic heterocystous cyanobacteria synthesize different heterocyst glycolipids than free-living heterocystous cyanobacteria. Phytochemistry 85: 115–121 Sherman L.A., Meunier P., Colón-López M.S. (1998) Diurnal rhythms in metabolism: a day in the life of a unicellular diazotrophic cyanobacterium. Photosynth Res 58:25–42. Shiozaki T., Ijichi M., Kodama T., Takeda S., and Furuya K. (2014) Heterotrophic bacteria as major nitrogen fixers in the euphotic zone of the Indian Ocean. Global Biogeochem Cycles 28: 1096–1110. Siesser W.G., Bralower J., and De Carlo H. (1992) Mid-Tertiary Braarudosphaera- rich sediments on the Exmouth Plateau. In: Proceedings of the Ocean Drilling Program, Sc Res 122: 653–663. Snow J.T., Schlosser C., Woodward E.M.S., Mills M.M., Achterberg E.P., Mahaffey, C., et al. (2015) Environmental controls on the biogeography of diazotrophy and Trichodesmium in the Atlantic Ocean. Global Biogeochem Cycles 29(6): 865–884. Sohm J.A., Webb E.A., and Capone D.G. (2011) Emerging patterns of marine nitrogen fixation. Nat Rev Microbiol 9 (7): 499–508.

Sournia A. (1970). Les cyanophycees dans le plancton marin. Ann. Biol. 9, 63–76. Sukenik A., Kaplan-Levy R. N., Welch J. M., and Post A F. (2012) Massive multiplication of genome and ribosomes in dormant cells (akinetes) of Aphanizomenon ovalisporum (Cyanobacteria). ISME J 6: 670–679. Staal M., Meysman F.J.R., Stal L.J. (2003) Temperature excludes N 2 -fixing heterocystous cyanobacteria in the tropical oceans. Nature 425: 504–507. Stal L.J. (2009) Is the distribution of nitrogen‐fixing cyanobacteria in the oceans related to temperature?. Environ Microbiol 11 (7): 1632–1645. Stenegren M., Caputo A., Berg C., Bonnet S., and Foster R.A. (2018) Distribution and drivers of symbiotic and free-living diazotrophic cyanobacteria in the Western Tropical South Pacific. Biogeosciences 15 (5): 1559–1578. Subramaniam A., Yager P.L., Carpenter E.J., Mahaffey C., Björkman K., Cooley S., et al. (2008) Amazon River enhances diazotrophy and carbon sequestration in the tropical North Atlantic Ocean. Pnas 105: 10460–10465. Sundström B.G. (1984) Observations on Rhizosolenia clevei Ostenfeld (Bacillariophyceae) and Richelia intracellularis Schmidt (Cyanophyceae). Bot Mar 27: 345–355. Tesson B., Lerch S.J., and Hildebrand M. (2017) Characterization of a new protein family associated with the silica deposition vesicle membrane enables genetic manipulation of diatom silica. Sci Rep 7(1): 13457. Theriot E.C., Ashworth M., Ruck E., Nakov T., and Jansen R.K. (2010) A preliminary multigene phylogeny of the diatoms (Bacillariophyta): challenges for future research. Plant Ecol Evol 143(3): 278–296. Thompson A., Carter B.J., Turk-Kubo K., Malfatti F., Azam F., and Zehr J.P. (2014) Genetic diversity of the unicellular nitrogen fixing cyanobacteria UCYN-A and its prymnesiophyte host. Environ Microbiol 16: 3238–3249 Thompson A.W., Foster R.A., Krupke A., Carter B.J., Musat N., Vaulot D., et al. (2012) Unicellular Cyanobacterium Symbiotic with a Single-Celled Eukaryotic Alga. Science 337:1546–1550. Thompson A.W. and Zehr J.P. (2013) Cellular interactions: lessons from the nitrogen‐ fixing cyanobacteria. J Phycol 49 (6): 1024–1035. Tovar-Sanchez A., Sanudo-Wilhelmy S.A., Kustka A.B., Agustí S., Dachs J., Hutchins D.A., et al. (2006) Effects of dust deposition and river discharges on trace metal composition of Trichodesmium spp. in the tropical and subtropical North Atlantic Ocean. Limnol Oceanogr 51(4): 1755–1761. Tragin M., Zingone A., and Vaulot D. (2018) Comparison of coastal phytoplankton composition estimated from the V4 and V9 regions of the 18S rRNA gene with a focus on photosynthetic groups and especially Chlorophyta. Environ Microbiol 20(2): 506–520.

Trapp E.M., Adler S., Zau-ner S., and Maier U.G. (2012) Rhopalodia gibba and its endosymbionts as a model for early steps in a cyanobacterial primary endosymbiosis. Endocytobiosis Cell Res 23: 21–24. Tripp H.J., Bench S.R., Turk K.A., Foster R.A., Desany B.A., Niazi F., et al. (2010) Metabolic streamlining in an open-ocean nitrogen-fixing cyanobacterium. Nature 464: 90–94. Trobajo R., Mann D.G., Li C., Sato S., Rimet F., Rovira L., et al. (2017) A four-gene approach to the Bacillariaceae: establishing a framework for classifying a highly diverse and taxonomically difficult diatom group. Phycologia 56(4): 187. Turk-Kubo K.A., Farnelid H.M., Shilova I.N., Henke B., and Zehr, J.P. (2017) Distinct ecological niches of marine symbiotic N2-fixing cyanobacterium Candidatus Atelocyanobacterium thalassa sublineages. J Phycol 461: 451–461. Turk-Kubo K.A., Frank I.E., Hogan M.E., Desnues A., Bonnet S., and Zehr J.P. (2015) Diazotroph community succession during the VAHINE mesocosm experiment (New Caledonia lagoon). Biogeosciences 12: 7435–7452. Uyeda J.C., Harmon L.J., and Blank C.E. (2016) A Comprehensive Study of Cyanobacterial Morphological and Ecological Evolutionary Dynamics through Deep Geologic Time. PLoS One 11: 1–32. Vaillancourt R.D., Marra J., Seki M.P., Parsons M.L., and Bidigare R.R. (2003) Impact of a cyclonic eddy on phytoplankton community structure and photosynthetic competency in the subtropical North Pacific Ocean. Deep Sea Res Part 1 Oceanogr Res Pap 50(7): 829–847. Venrick E.L. (1974) The distribution and significance of Richelia intracellularis Schmidt in the North Pacific Central Gyre. Limnol Oceanogr 19: 437–445. Villareal T.A. (1989) Division cycles in the nitrogen-fixing Rhizosolenia (Bacillariophyceae)-Richelia (Nostocaceae) symbiosis. Eur J Phycol 24: 357–365. Villareal T.A. (1990) Laboratory culture and preliminary characterization of the nitrogen-fixing Rhizosolenia-Richelia symbiosis. Mar Ecol 11: 117–132. Villareal T.A. (1991) Nitrogen-fixation by the cyanobacterial symbiont of the diatom genus Hemiaulus. Mar Ecol Prog Ser 76(2): 201–204. Villareal T.A. (1992) Marine Nitrogen-Fixing Diatom-Cyanobacteria Symbioses. In Marine Pelagic Cyanobacteria: Trichodesmium and other Diazotrophs. Carpenter, E.J., Capone, D.G., Reuter, J.G. (eds). Dordecht: Kluwer pp. 163–75. Villareal T.A. (1994) Widespread occurrence of the Hemiaulus-cyanobacterial symbiosis in the southwest North Atlantic Ocean. Bull Mar Sci 54(1): 1–7. Villareal T.A., Brown C.G., Brzezinski M.A., Krause J.W., and Wilson C. (2012) Summer Diatom Blooms in the North Pacific Subtropical Gyre: 2008–2009 Peck M (ed). PLoS One 7: 1–15.

Wallner G., Amann R., and Beisker W. (1993) Optimizing fluorescent in situ hybridization with rRNA targeted oligonucleotide probes for flow cytometric identification of microorganisms. Cytometry 14: 136–143. Walworth N.G., Fu F.X., Lee M.D., Cai X., Saito M.A., Webb E.A., et al. (2018) Nutrient-colimited Trichodesmium as a nitrogen source or sink in a future ocean. Appl Environ Microbiol 84(3): e02137-17. Walworth, N.G., Fu F.X., Webb E.A., Saito M.A., Moran D., Mcllvin M.R., et al. (2016) Mechanisms of increased Trichodesmium fitness under iron and phosphorus co-limitation in the present and future ocean. Nat Comm 7(12081): 1– 11. Walworth N.G., Pfreundt U., Nelson W. C. Mincer T., Heidelberg J.F., Fu F.X., et al. (2015) Trichodesmium genome maintains abundant, widespread noncoding DNA in situ, despite oligotrophic lifestyle. Proc Natl Acad Sci (201422332): 1–6. Warshan D., Espinoza J.L., Stuart R.K., Richter R.A., Kim S.Y., Shapiro N., et al. (2017) Feathermoss and epiphytic Nostoc cooperate differently: expanding the spectrum of plant–cyanobacteria symbiosis. ISME J 11(12): 2821–2833. Waterbury J.B., Watson S.W., Valois F.W., and Franks D.G. (1986) Biological and ecological characterization of the marine unicellular cyanobacterium Synechococcus. In: Canadian Bulletin of Fisheries and Aquatic Sciences. Platt, T., and Li, W.K.W. (eds). Ottawa, Canada: Department of Fisheries and Oceans, pp. 71–120. Webb E.A., Ehrenreich I.M., Brown S.L., Valois F.W., and Waterbury J.B. (2009) Phenotypic and genotypic characterization of multiple strains of the diazotrophic cyanobacterium, Crocosphaera watsonii, isolated from the open ocean. Environ Microbiol 11: 338–348. Webb E.A., Moffett J.W., and Waterbury J.B. (2001) Iron stress in open-ocean cyanobacteria (Synechococcus, Trichodesmium, and Crocosphaera spp): identification of the IdiA protein. Appl Environ Microbiol 67: 5444–5452. Weber T. and Deutsch C. (2014) Local versus basin-scale limitation of marine nitrogen fixation. Proc. Natl. Acad. Sci. U.S.A. 111(24): 8741–8746. Weis V.M. (2008) Cellular mechanisms of Cnidarian bleaching: stress causes the collapse of symbiosis. J Exp Biol 211(19): 3059–3066. Welsh E.A., Liberton M., Stöckel J., Loh T., Elvitigala T., Wang C., et al. (2008). The genome of Cyanothece 51142, a unicellular diazotrophic cyanobacterium important in the marine nitrogen cycle. Proc Natl Acad Sci 105 (39): 15094– 15099. Wilson C., Villareal T.A., Maximenko N., Bograd S.J., Montoya J.P., and Schoenbaechler C.A. (2008) Biological and physical forcings of late summer chlorophyll blooms at 30 N in the oligotrophic Pacific. J Mar Syst 69(3-4): 164– 176.

White A.E., Prahl F.G., Letelier R.M., and Popp B.N. (2007) Summer surface waters in the Gulf of California: prime habitat for biological N2-fixation. Global Biogeochem Cycles 21: 1–11. Xu H.H. and Tabita F.R. (1996) Ribulose-1,5-bisphosphate carboxylase/oxygenase gene expression and diversity of Lake Erie planktonic microorganisms. Appl Environ Microbiol 62:1913–1921 Yu M., Ashworth M.P., Hajrah N.H., Khiyami M.A., Sabir M.J., Alhebshi A.M., et al. (2018) Evolution of the Plastid Genomes in Diatoms. In: Advances in Botanical Research (Vol. 85, pp. 129-155). Academic Press. Zeev E.B., Yogev T., Man-Aharonovich D., Kress N., Herut B., Béjà O., et al. (2008) Seasonal dynamics of the endosymbiotic, nitrogen-fixing cyanobacterium Richelia intracellularis in the eastern Mediterranean Sea. ISME J 2: 911–23. Zehr J.P. (2011) Nitrogen fixation by marine cyanobacteria. Trends Microbiol 19: 162–173. Zehr J.P., Bench S.R., Carter B.J., Hewson I., Niazi F., Shi T., et al. (2008) Globally distributed uncultivated oceanic N2-fixing cyanobacteria lack oxygenic photosystem II. Science 322, 1110–1112. Zehr J.P., Bench S.R., Mondragon E.A., McCarren J., and DeLong E.F. (2007) Low genomic diversity in tropical oceanic N2-fixing cyanobacteria. Proc Natl Acad Sci 104: 17807–17812. Zehr J.P., Jenkins B.J., Short S.M., and Steward G.F. (2003) Nitrogenase gene diversity and microbial community structure: a cross-system comparison. Environ Microbiol 5: 539–554. Zhang C.C., Huguenin S., and Friry, A. (1995) Analysis of genes encoding the cell division protein FtsZ and a glutathione synthetase homologue in the cyanobacterium Anabaena sp. PCC 7120. Res Microbial 146(6): 445–455. Zhang C., Stadler T., Klopfstein S., et al. (2016) Total-evidence dating under the fossilized birth–death process. Syst Appl Microbiol 65: 228–249. Zhang Y., Zhao Z., Sun J., and Jiao N. (2011) Diversity and distribution of diazotrophic communities in the South China Sea deep basin with mesoscale cyclonic eddy perturbations. FEMS Microbiol Ecol 78: 417–427. Zimmermann J., Jahn R., and Gemeinholzer B. (2011) Barcoding diatoms: Evaluation of the V4 subregion on the 18S rRNA gene, including new primers and protocols. Org Divers Evol 11: 173–192.