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# Response to light stimuli in the marine diatom Phaeodactylum tricornutum: Involvement of bHLH-PAS proteins in the circadian clock and plastid physiology

Alessandro Manzotti

► To cite this version:

Alessandro Manzotti. Response to light stimuli in the marine diatom Phaeodactylum tricornutum: Involvement of bHLH-PAS proteins in the circadian clock and plastid physiology. Biochemistry, Molecular Biology. Sorbonne Université, 2022. English. NNT: 2022SORUS232. tel-04562734

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Submitted on 29 Apr 2024

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Sorbonne Université

Ecole doctorale Complexité du Vivant - ED515

UMR7141 - Biologie du chloroplaste et perception de la lumière chez les micro-algues

Response to light stimuli in the marine diatom

*Phaeodactylum tricornutum :*

Involvement of bHLH-PAS proteins
in the circadian clock and plastid physiology

Par Alessandro Manzotti

Thèse de doctorat de Biologie

Dirigée par Angela Falciatore

Date de soutenance : 28 avril 2022

Devant un jury composé de :

Prof. Maria Mittag, rapportrice
Friedrich Schiller University, Jena

Prof. Wim Vyverman, rapporteur
Ghent University

Dr. François-Yves Bouget, examinateur
Laboratoire d'Océanographie Microbienne, CNRS, Sorbonne Université

Dr. Alessandra Rogato, examinatrice
Institute of Bioscience and BioResources, Naples, CNR

Dr. Angela Falciatore, directrice de recherche, directrice de thèse
Institut de Biologie Physico-Chimique, CNRS, Sorbonne Université

Dr. Jean-Pierre Bouly, maître de conférences, co-encadrant de thèse
Institut de Biologie Physico-Chimique, CNRS, Sorbonne Université

Membre invitée:

Prof. Eva Farré
Michigan State University

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SORBONNE
UNIVERSITÉ

CRÉATEURS DE FUTURS
DEPUIS 1257
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<figure><img src="image_4.png" /></figure>

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Except where otherwise noted, this work is licensed under

http://creativecommons.org/licenses/by-nc-nd/3.0/
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# Remerciements

Tant de personnes ont rendu cette thèse possible et je les remercie toutes du fond du cœur.

Merci à Angela, de m'avoir accueilli dans ton laboratoire il y a plus de quatre ans, alors que j'étais encore un étudiant en master perdu dans cette grande ville. Merci de m'avoir appris la valeur de la ténacité, de la persévérance et, pourquoi pas, même de l'entêtement. Merci pour toute l'énergie que tu mets dans ce que tu fais. En d'autres termes, merci de m'avoir appris l'importance d'avoir la capa tosta.

Merci à Jean-Pierre pour les longues discussions scientifiques et non scientifiques, au labo comme au pub. Merci pour ton ubiquité et ta capacité à toujours gérer tous les problèmes et imprévus avec la maîtrise d'un jongleur, merci pour tous les moments de défoulement et toutes les fois où tu as su me faire voir les difficultés sous le bon angle. Merci d'être un ami avant même d'être un collègue. Merci pour toutes les clopes, les bières, les whisky... bref, merci pour tout.

Thank of course to all the people that are part of my thesis committee: Maria Mittag, Wim Vyverman, François-Yves Bouget, Eva Farré and Alessandra Rogato, I hope you will enjoy the reading.

Merci à Marianne et Soizic, pour les innombrables fois où vous m'avez aidé et dépanné et pour m'avoir appris tant de choses sur la vie de laboratoire. Votre professionnalisme a été un exemple à suivre et m'a permis de grandir énormément. Merci à Carole, qui a commencé cette aventure du doctorat avec moi et qui la termine avec moi. Merci pour toutes les chansonnettes sifflées dans les couloirs et qui rentrent dans la tête pour ne plus en sortir, merci pour les memes, les rires et ta capacité à tout affronter avec une sérieuse légèreté.

Merci à tous les autres membres de l'UMR7141, un formidable groupe de chercheur.euse.s qui fait vivre ce laboratoire historique. Ce fut un plaisir et une joie de travailler avec vous.

Merci à Anaëlle, Anäis, Aurélian, Bashung, Camila, Coline, Dorian, Eleanor, Julie, Léa, Lorenzo, Noa, Tania... merci à tou.t.e.s les LUPAmi.e.s de m'avoir accueilli dans cette clique de bobos-babos-écolos parisien.ne.s. Merci d'avoir fait de Paris ma deuxième maison, merci d'être devenu.e.s ma deuxième famille (une famille un peu dysfonctionnelle, c'est vrai, mais sinon, où est le plaisir ?).

Grazie ai Rituals, ad Antonio, Arianna, Claudio, Eleonora, Flaminia, Matilde, Paolo, Seed e Stefano, gruppo matto e disperatissimo di italiano passato per Parigi ad un momento o ad un altro. Grazie per tutte le volte che abbiamo riso e sdrhamatizzato sulle nostre vite da dottorandi sull'orlo di una crisi di nervi. Vi aspetto dall'altra parte del varco, dai che manca poco.

Grazie a Marco e Luca, fratelli. Grazie per tutte le avventure, piccole e grandi. Grazie per le discussioni serissime sulle questioni più stupide e le discussioni stupidissime sulle questioni più serie. Grazie per avermi insegnato come elementi tanto diversi possono creare legami indissolubili. Grazie per aver dimostrato come si può cambiare ogni giorno senza cambiare mai davvero.

E in fine grazie a mia madre per tutti gli infiniti sacrifici che ha fatto e che continua a fare per me, anche se questo vuol dire avermi sempre lontano da casa. Sei la persona più forte e inarrestabile che conosca.

Enjoy the reading

Bonne lecture

Buona lettura
# Table of contents

## Chapter I: Introduction p.3

I.1 The life of phytoplankton p.4

I.2 The kaleidoscopic world of diatoms p.7

I.2.a General introduction on diatoms p.7

I.2.b Phylogenesis and evolution p.10

I.2.c Toolkit box for molecular biology in diatoms p.13

I.2.d *Phaeodactylum tricornutum* as a diatom model p.15

I.3 Light in the life of diatoms p.17

I.3.a Light as an energy source: the photosynthetic apparatus in diatoms p.17

I.3.b Photoacclimation and photoprotective mechanisms p.21

I.3.c Light as a source of information: photo-perception p.23

I.3.d Cell cycle progression and regulation in diatoms p.25

I.4 Biological clocks p.31

I.4.a Biological clocks: principles and characteristics p.31

I.4.b Biological clocks: diversity and commonalities p.34

I.4.c Translation Independent Oscillators: above and beyond TTFLs p.41

I.4.d Circadian rhythmicity in Stramenopila: an overview p.44

I.5 bHLH-PAS transcription factors in diatoms p.45

Aims of the thesis p.50

## Chapter II: The circadian clock of diatoms p.83

## Chapter II.a: Rhythms and clocks in marine organisms p.87

Abstract p.88

1. Cycles of the sun and moon cause a multitude of biological rhythms p.89

2. Other places, other rhythms: The oceans harbor multiple and diverse habitats with very different properties from land p.91

3. Rhythms in the life of marine algae p.94

Algal models for the elucidation of marine time-keeping mechanisms p.96

4. Animal models for marine chronobiology p.98

Vertical migrations in the pelagic zone p.99

Rhythms in the deep sea p.100

A plastic clock to time daily and tidal rhythms p.101

Timing reproduction with an inner calendar p.104

5. Marine rhythms and clocks under climate change p.107

Present-day climate effects on marine rhythmicity p.108

Temperature changes p.108

pH changes p.109

Oxygen concentration changes p.109

Adaptability of rhythms and clocks to future ocean conditions p.110

Phenotypic adaptive capacity p.111

Evolutionary adaptation of clock and rhythms p.111

Interspecies mismatches p.112

6. Conclusions and outlook p.113
<table><tr><td>Glossary: Rhythms and clocks in marine organisms</td><td>p.114</td></tr><tr><td>Acknowledgements</td><td>p.116</td></tr><tr><td>Bibliography</td><td>p.118</td></tr><tr><td>Supplementary Material</td><td>p.139</td></tr><tr><td>Supplementary Bibliography</td><td>p.140</td></tr></table>

## Chapter II.b: bHLH-PAS protein RITMO1 regulates diel biological rhythms in the marine diatom *Phaeodactylum tricornutum* p.145

<table><tr><td>Abstract and Significance</td><td>p.146</td></tr><tr><td>Results</td><td>p.147</td></tr><tr><td>Transcriptome Profiling Identifies Potential Regulators of Diurnal Rhythms in <i>P. tricornutum</i></td><td>p.147</td></tr><tr><td>Rhythmic <i>bHLH1a</i> and <i>bHLH1b</i> Expression Is Adjusted in a Photoperiod Dependent Manner and Persists in Continuous Light and Continuous Dark Conditions</td><td>p.147</td></tr><tr><td>Cellular Localization of <i>bHLH1a</i> Protein</td><td>p.147</td></tr><tr><td><i>bHLH1a</i> Regulates Pace of Diel Gene Expression</td><td>p.147</td></tr><tr><td>The Ectopic Overexpression of <i>bHLH1a</i> Results in Altered Circadian Rhythms</td><td>p.148</td></tr><tr><td>RITMO1-Like Proteins Are Widely Represented in the Genome of Marine Algae</td><td>p.148</td></tr><tr><td>Discussion</td><td>p.149</td></tr><tr><td>Methods</td><td>p.151</td></tr><tr><td>Gene Expression Analyses</td><td>p.151</td></tr><tr><td>Generation of the <i>bHLH1a</i> Transgenic Lines</td><td>p.151</td></tr><tr><td>Microscopic Analysis</td><td>p.151</td></tr><tr><td>Data Mining, Protein Sequence, and Phylogenetic Analysis</td><td>p.151</td></tr><tr><td>Acknowledgements</td><td>p.151</td></tr><tr><td>Bibliography</td><td>p.151</td></tr><tr><td>SI Appendix</td><td>p.152</td></tr><tr><td>Supplementary methods</td><td>p.152</td></tr><tr><td>Culture and irradiation conditions</td><td>p.152</td></tr><tr><td>Analysis of the rhythmic processes</td><td>p.152</td></tr><tr><td>RNA extraction and transcript analyses</td><td>p.153</td></tr><tr><td>Immunoblot analysis</td><td>p.153</td></tr><tr><td><i>bHLH1a</i> subcellular localization</td><td>p.154</td></tr><tr><td>Generation of the <i>bHLH1a</i> transgenic lines</td><td>p.154</td></tr><tr><td>Selection of rhythmic transcripts and clustering analysis</td><td>p.154</td></tr><tr><td>Data mining, protein sequence and phylogenetic analysis</td><td>p.155</td></tr><tr><td>Cell cycle analysis</td><td>p.155</td></tr><tr><td>References</td><td>p.177</td></tr></table>

## Chapter II.c: Physiological rhythms in the marine diatom *Phaeodactylum tricornutum* are regulated by the circadian clock transcriptional regulator RITMO1 p.181

<table><tr><td>Abstract</td><td>p.182</td></tr><tr><td>Introduction</td><td>p.183</td></tr><tr><td>Materials and Methods</td><td>p.185</td></tr><tr><td>Cultures and irradiation conditions</td><td>p.185</td></tr><tr><td>Generation of RITMO1 Knock Out (KO) strains</td><td>p.185</td></tr></table>
<table><tbody><tr><td>Western Blot Analysis</td><td>p.185</td></tr><tr><td>Cellular fluorescence analysis</td><td>p.186</td></tr><tr><td>Analysis of the rhythmic processes</td><td>p.186</td></tr><tr><td>Cell cycle analysis</td><td>p.186</td></tr><tr><td>Analysis of photosynthetic parameters</td><td>p.187</td></tr><tr><td>RNA extraction and RT-qPCR analysis</td><td>p.187</td></tr><tr><td>Results</td><td>p.189</td></tr><tr><td>Generation of knock-out mutants for <i>P. tricornutum</i> RITMO1 by CRISPR-Cas9</td><td>p.189</td></tr><tr><td>Diatoms cellular fluorescence rhythmicity is adjusted in a photoperiod-dependent manner and is not altered in knock-out mutants for RITMO1</td><td>p.190</td></tr><tr><td>RITMO1 obliteration affects fluorescence rhythmicity under free-running conditions</td><td>p.192</td></tr><tr><td>RITMO1 affects circadian fluorescence rhythmicity, without affecting growth and cell cycle progression</td><td>p.195</td></tr><tr><td>RITMO1 is implicated in the circadian regulation of photosynthesis in <i>P. tricornutum</i></td><td>p.196</td></tr><tr><td>RITMO1 plays a central role in the circadian control of gene expression in diatoms</td><td>p.198</td></tr><tr><td>Discussion</td><td>p.202</td></tr><tr><td>Acknowledgements</td><td>p.205</td></tr><tr><td>Bibliography</td><td>p.206</td></tr><tr><td>Supplementary Material</td><td>p.210</td></tr></tbody></table>

<table><tbody><tr><td>Chapter II.d: bHLH1b as a candidate partner of RITMO1 in diatom circadian clock</td><td>p.219</td></tr><tr><td>Abstract</td><td>p.220</td></tr><tr><td>Introduction</td><td>p.221</td></tr><tr><td>Materials and methods</td><td>p.223</td></tr><tr><td>Yeast two Hybrid (Y2H) Assay</td><td>p.223</td></tr><tr><td>Culture and irradiation conditions</td><td>p.223</td></tr><tr><td>CRISPR/Cas9 Mutagenesis</td><td>p.223</td></tr><tr><td>Flow cytometry and fluorescence rhythmicity analysis</td><td>p.225</td></tr><tr><td>Results</td><td>p.226</td></tr><tr><td>bHLH1b and RITMO1 physically interact via their bHLH fold</td><td>p.226</td></tr><tr><td>Generation of bHLH1b KO mutants</td><td>p.228</td></tr><tr><td>bHLH1b mutations affect free-run fluorescence oscillations</td><td>p.230</td></tr><tr><td>Conclusions</td><td>p.232</td></tr><tr><td>Bibliography</td><td>p.234</td></tr></tbody></table>

<table><tbody><tr><td>Chapter III: bHLH2 is a light-intensity dependent modulator of photosynthesis and growth in the marine diatom <i>Phaeodactylum tricornutum</i></td><td>p.239</td></tr><tr><td>Abstract</td><td>p.241</td></tr><tr><td>Introduction</td><td>p.242</td></tr><tr><td>Analysis of photosynthesis via 2 chlorophyll fluorescence</td><td>p.243</td></tr><tr><td>Methods</td><td>p.247</td></tr><tr><td>Generation of bHLH RNAi lines</td><td>p.247</td></tr><tr><td>On plate colony screening</td><td>p.247</td></tr><tr><td>Liquid culture and irradiation conditions</td><td>p.248</td></tr><tr><td>ΦPSII analysis via chlorophyll fluorescence</td><td>p.248</td></tr></tbody></table>
<table><tr><td>relative Electron Transfer Rate</td><td>p.249</td></tr><tr><td>Evaluation of PSI redox state</td><td>p.249</td></tr><tr><td>Lincomycin treatment</td><td>p.249</td></tr><tr><td>RNA extraction and RT-qPCR analysis</td><td>p.249</td></tr><tr><td>Subcellular localization prediction</td><td>p.250</td></tr><tr><td>Generation of <i>bHLH2</i> Knock-Out (KO) strains</td><td>p.250</td></tr></table>

<table><tr><td>Results and Discussion</td><td>p.251</td></tr><tr><td>Generation of RNAi transgenic lines targeting <i>bHLH</i> genes</td><td>p.251</td></tr><tr><td><i>bHLH2</i> RNAi lines are affected in photosynthetic capacity in a light intensity dependent way</td><td>p.252</td></tr><tr><td>Effects of antisense RNAi on <i>bHLH2</i> transcription</td><td>p.255</td></tr><tr><td><i>bHLH2</i> RNAi lines show a deficit in electron transfer at the thylakoid chain level</td><td>p.256</td></tr><tr><td>Response of photosynthesis upon transition to high light is affected in <i>bHLH2</i> RNAi lines</td><td>p.256</td></tr><tr><td><i>bHLH2</i> RNAi lines are not affected in response to photoinhibition</td><td>p.259</td></tr><tr><td>Photosynthetic activity represses <i>bHLH2</i> expression</td><td>p.260</td></tr><tr><td>Preliminary characterization of <i>bHLH2</i> KO mutants</td><td>p.263</td></tr><tr><td>Conclusion</td><td>p.266</td></tr><tr><td>Bibliography</td><td>p.268</td></tr></table>

<table><tr><td>Chapter IV: Conclusions and perspectives</td><td>p.273</td></tr><tr><td>Bibliography</td><td>p.282</td></tr></table>
# List of Figures and Tables

## Chapter I

<table><tr><td>Figure 1. Chlorophyll distribution in the world ocean</td><td>p.5</td></tr><tr><td>Figure 2. Diatom characteristics and morphological diversity</td><td>p.8</td></tr><tr><td>Figure 3. The eukaryotic Tree of Life</td><td>p.10</td></tr><tr><td>Figure 4. The evolution of diatoms</td><td>p.12</td></tr><tr><td>Figure 5. The photosynthetic Linear Electron Transfer in diatoms</td><td>p.18</td></tr><tr><td>Figure 6. Diatom cell cycle and cyclins</td><td>p.27</td></tr><tr><td>Figure 7. Main characteristics of biological rhythms</td><td>p.32</td></tr><tr><td>Figure 8. Circadian TTFL networks in model organisms</td><td>p.36</td></tr><tr><td>Figure 9. Architecture of circadian clock systems</td><td>p.43</td></tr></table>

## Chapter II.a

<table><tr><td>Figure 1. Complex rhythms emerge from solar and lunar cycles in marine ecosystems</td><td>p.90</td></tr><tr><td>Figure 2. Overview of the diverse marine habitats and discussed exemplary organisms</td><td>p.92</td></tr><tr><td>Figure 3 Established and emerging marine microalgal model systems for chronobiology</td><td>p.94</td></tr><tr><td>Figure 4. Climate change effects on timing and how they could impact on species interactions and ecosystems</td><td>p.110</td></tr><tr><td>Table 1. Characterized regulators, central clock system components and rhythmic outputs of marine algal models</td><td>p.98</td></tr><tr><td>Table 2. Overview of a comprehensive selection of different marine animals studied for their rhythms</td><td>p.104</td></tr><tr><td>Table S1. Examples of biological rhythms characterized in marine algae</td><td>p.139</td></tr></table>

## Chapter II.b

<table><tr><td>Figure 1. Diurnal expression of rhythmic <i>P. tricornutum</i> genes</td><td>p.148</td></tr><tr><td>Figure 2. bHLH1a localizes in the nucleus, and its overexpression alters rhythmic diel gene expression</td><td>p.149</td></tr><tr><td>Figure 3. bHLH1a overexpression alters circadian rhythms of cellular fluorescence</td><td>p.150</td></tr><tr><td>Figure 4. bHLH-PAS protein family structure and phylogeny</td><td>p.150</td></tr></table>
<table>
  <tbody>
    <tr>
      <td>Figure S1. nCounter expression analysis of genes maintaining rhythmic expression in D:D condition and 16L:8D condition in Wt cells</td>
      <td>p.157</td>
    </tr>
    <tr>
      <td>Figure S2. nCounter expression analysis of representative genes with altered rhythmic expression in Wt cells in D:D condition compared to 16L:8D condition</td>
      <td>p.158</td>
    </tr>
    <tr>
      <td>Figure S3. Diel expression patterns of <i>bHLH1a</i> and <i>bHLH1b</i> genes under Fe-depletion conditions in 12L:12D photoperiods</td>
      <td>p.159</td>
    </tr>
    <tr>
      <td>Figure S4. Analysis of selected rhythmic gene expression profiles in L:D and D:D in Wt and <i>bHLH1a</i> OE1 cells</td>
      <td>p.160</td>
    </tr>
    <tr>
      <td>Figure S5. Cell cycle progression dynamics of Wt <i>P. tricornutum</i></td>
      <td>p.161</td>
    </tr>
    <tr>
      <td>Figure S6. Diurnal oscillations of chlorophyll fluorescence (FL3-A parameter) in Wt and <i>bHLH1a</i> OE lines</td>
      <td>p.162</td>
    </tr>
    <tr>
      <td>Figure S7. Analysis of <i>bHLH1a</i> gene expression in continuous blue light</td>
      <td>p.163</td>
    </tr>
    <tr>
      <td>Figure S8. Circadian oscillation of chlorophyll fluorescence in Wt and <i>bHLH1a</i> OE lines</td>
      <td>p.164</td>
    </tr>
    <tr>
      <td>Figure S9. Analysis of rhythmicity in Wt and <i>bHLH1a</i> OE lines under blue L:L</td>
      <td>p.165</td>
    </tr>
    <tr>
      <td>Figure S10. Growth rate of Wt and <i>bHLH1a</i> OE lines grown in blue L:L</td>
      <td>p.166</td>
    </tr>
    <tr>
      <td>Figure S11. Circadian oscillation of chlorophyll fluorescence in Wt and a transgenic control line (pNAT) under blue (L:L)</td>
      <td>p.167</td>
    </tr>
    <tr>
      <td>Table S1. Accession codes and sequence probes used in the nCounter analysis</td>
      <td>p.168</td>
    </tr>
    <tr>
      <td>Table S2. Calculated periods, phases, amplitudes and relative amplitude errors of the Wt and <i>bHLH1a</i> OE lines growing in 16L:8D</td>
      <td>p.171</td>
    </tr>
    <tr>
      <td>Table S3. Calculated periods, phases, amplitudes and relative amplitude errors of the Wt and <i>bHLH1a</i> OE lines growing in L:L</td>
      <td>p.172</td>
    </tr>
    <tr>
      <td>Table S4. Accession numbers of the proteins utilized in the <i>bHLH-PAS</i> phylogenetic analysis</td>
      <td>p.173</td>
    </tr>
    <tr>
      <td>Table S5. List of the oligonucleotides used in this work</td>
      <td>p.176</td>
    </tr>
  </tbody>
</table>

## Chapter II.c

<table>
  <tbody>
    <tr>
      <td>Figure 1. Generation of knock-out (KO) mutants for <i>P. tricornutum</i> RITMO1</td>
      <td>p.189</td>
    </tr>
    <tr>
      <td>Figure 2. Fluorescence rhythmicity of wild-type and ritmo1 knock-out mutants under different light:dark cycles</td>
      <td>p.191</td>
    </tr>
    <tr>
      <td>Figure 3. Characterization of fluorescence rhythmicity of wild-type <i>P. tricornutum</i> cells after the transition from L:D cycles to free running conditions</td>
      <td>p.192</td>
    </tr>
    <tr>
      <td>Figure 4. Effects of RITMO1 mutation on cellular fluorescence in free running</td>
      <td>p.194</td>
    </tr>
  </tbody>
</table>
<table>
  <tbody>
    <tr>
      <td>Figure 5. Re-entrainment of Wt, KO1, KO3 and OE1 cultures adapted to continuous light (L:L) to L:D cycles</td>
      <td>p.195</td>
    </tr>
    <tr>
      <td>Figure 6. Analysis of the effect of RITMO1 obliteration and overexpression on growth capacity and cell cycle progression</td>
      <td>p.196</td>
    </tr>
    <tr>
      <td>Figure 7. Analysis of the effect of RITMO1 mutation and overexpression on different photosynthetic parameters under periodic L:D cycles and free running conditions</td>
      <td>p.197</td>
    </tr>
    <tr>
      <td>Figure 8. Analysis of the expression of selected genes by qRT-PCR in Wt, OE1 and KO3 lines under periodic L:D cycles and free running conditions</td>
      <td>p.201</td>
    </tr>
    <tr>
      <td>Figure S1. Sequence analysis of the KO mutants in the RITMO1 gene</td>
      <td>pp.210-211</td>
    </tr>
    <tr>
      <td>Figure S2. Characterisation of growth profile in P. tricornutum grown under 16L:8D cycles</td>
      <td>p.212</td>
    </tr>
    <tr>
      <td>Figure S3. Effects of RITMO1 mutation on cellular fluorescence in L:L 25 µmol photons m-2 s-1</td>
      <td>p.213</td>
    </tr>
    <tr>
      <td>Figure S4. Time-resolved analysis of growth in free-run cultures</td>
      <td>p.214</td>
    </tr>
    <tr>
      <td>Table S1. List of the sgRNAs used for CRISPR-Cas9 mutagenesis of the RITMO1</td>
      <td>p.215</td>
    </tr>
    <tr>
      <td>Table S2. Growth capacity of P. tricornutum Wt cells in the different conditions used throughout this study</td>
      <td>p.216</td>
    </tr>
    <tr>
      <td>Table S3. Characterisation of cellular fluorescence rhythmicity in Wt, CTR, KO1, KO3 and OE1 lines in L:D</td>
      <td>p.217</td>
    </tr>
    <tr>
      <td>Table S4. Characterisation of cellular fluorescence rhythmicity in Wt, CTR, KO1, KO3 and OE1 lines in L:L</td>
      <td>p.218</td>
    </tr>
    <tr>
      <td>Table S5. List of the oligonucleotides used in this work</td>
      <td>p.219</td>
    </tr>
  </tbody>
</table>

## Chapter II.d

<table>
  <tbody>
    <tr>
      <td>Figure 1. Genomic arrangement of <i>RITMO1</i> and <i>bHLH1b</i> loci</td>
      <td>p.221</td>
    </tr>
    <tr>
      <td>Figure 2. Results of the Yeast two Hybrid assay (Y2H)</td>
      <td>p.227</td>
    </tr>
    <tr>
      <td>Figure 3. Generation of <i>bHLH1b</i> KO mutants</td>
      <td>p.228</td>
    </tr>
    <tr>
      <td>Figure 4. Characterisation of <i>bHLH1b</i> KO mutants</td>
      <td>p.230</td>
    </tr>
    <tr>
      <td>Figure 5. Effects of <i>bHLH1b</i> mutation on cellular fluorescence</td>
      <td>p.231</td>
    </tr>
    <tr>
      <td>Table 1. List of the oligonucleotides used in this work</td>
      <td>p.224</td>
    </tr>
    <tr>
      <td>Table 2. sgRNA used for <i>bHLH1b</i> KOs generation</td>
      <td>p.224</td>
    </tr>
    <tr>
      <td>Table 3. Characterisation of cellular fluorescence rhythmicity of <i>bHLH1b</i> mutants in L:L</td>
      <td>p.230</td>
    </tr>
  </tbody>
</table>
Chapter III

<table>
  <tbody>
    <tr>
      <td>Figure 1. Principles of chlorophyll fluorescence analysis of PSII activity</td>
      <td>p.243</td>
    </tr>
    <tr>
      <td>Figure 2. Use of P700 redox state to localize perturbations in the plastid electron transfer chain</td>
      <td>p.245</td>
    </tr>
    <tr>
      <td>Figure 3. Screening on plate based on chlorophyll fluorescence</td>
      <td>p.251</td>
    </tr>
    <tr>
      <td>Figure 4. On plate screening of phleomycin resistant control</td>
      <td>p.252</td>
    </tr>
    <tr>
      <td>Figure 5. bHLH2 RNAi lines are affected in photosynthesis and growth</td>
      <td>p.253</td>
    </tr>
    <tr>
      <td>Figure 7. Gene expression of <i>bHLH2</i> under the different continuous light irradiations</td>
      <td>p.255</td>
    </tr>
    <tr>
      <td>Figure 8. <i>bHLH2</i> RNAi lines show a defect in electron transfer located upstream of PSI</td>
      <td>p.257</td>
    </tr>
    <tr>
      <td>Figure 9. Response to light intensity transition is affected in <i>bHLH2</i> RNAi lines</td>
      <td>p.258</td>
    </tr>
    <tr>
      <td>Figure 10. <i>bHLH2</i> RNAi lines are not affected in photoinhibition</td>
      <td>p.259</td>
    </tr>
    <tr>
      <td>Figure 11. Expression profiles of <i>bHLH2</i> from publicly available transcriptomic data</td>
      <td>p.261</td>
    </tr>
    <tr>
      <td>Figure 12. <i>bHLH2</i> expression in the dark</td>
      <td>p.262</td>
    </tr>
    <tr>
      <td>Figure 13. Mutants on <i>bHLH2</i> gene induced by CRISPR/Cas9</td>
      <td>p.263</td>
    </tr>
    <tr>
      <td>Figure 14. Sequence alignment of the KO mutants for <i>bHLH2</i> gene</td>
      <td>p.264</td>
    </tr>
    <tr>
      <td>Figure 15. Response to light intensity transition in <i>bHLH2</i> KO lines</td>
      <td>p.265</td>
    </tr>
    <tr>
      <td>Table. 1 Primers used for the construction of the RNAi cassettes</td>
      <td>p.247</td>
    </tr>
    <tr>
      <td>Table 2. List of the oligonucleotides used in this work</td>
      <td>p.250</td>
    </tr>
    <tr>
      <td>Table 3. gRNAs used for <i>bHLH2</i> KOs generation</td>
      <td>p.250</td>
    </tr>
    <tr>
      <td>Table 4. Summary results of on plate screening</td>
      <td>p.252</td>
    </tr>
    <tr>
      <td>Table 5. Summary results of the RLC analysis</td>
      <td>p.258</td>
    </tr>
  </tbody>
</table>
Table of abbreviations

ALAN: Artificial Light At Night

BCP: Biological Carbon Pump

bHLH: basic Helix Loop Helix domain

BP: Before Present

CBBC: Calvin Benson Bassham Cycle

CCM: Carbon Concentrating Mechanism

CET: Cyclic Electron Transfer

D:D: continuous dark

DVM: Diel Vertical Migration

EET: Electron Energy Transfer

EGT: Endosymbiont Gene Transfer

eToL: eukaryote Tree of Life

ETR: Electron Transfer Rate

FCP: Fucoxanthin Chlorophyll binding Protein

FD: Ferredoxin

FFC: FRQ-FRH Complex

FLD: Flavodoxin

Fx: Fucoxanthin

Gt C: Giga Ton of carbon

HGT: Horizontal Gene Transfer

KD: Knock-Down transgenic line

KO: Knock-Out mutant

L:D: light dark cycles

L:L: continuous light

LET: Linear Electron Transfer

LGM: Last Glacial Maximum

LHC: Light Harvesting Complex

LOV: Light Oxygen Voltage domain

MC: alternative of MTOC

MT: Micro Tubules

MTOC: Micro Tubules Organising Centre

Mya: Million years ago

NPP: Net Primary Production

NPQ: Non Photochemical Quenching

OE: Over-Expressing transgenic line

OMZ: Oxygen Minimum Zones

OUC: Ornithine-Urea Cycle

PAS: *Per Arnt Sim* domain

PQ: PlastoQuinone

PS: Photo System

PTO: Post-Translational Oscillator

qE: energy (heat) dependent component of NPQ

qI: photoinhibition component of NPQ

qT: state transition component of NPQ

RC: Reaction Center

rETR: relative Electron Transfer Rate

RNAi: RNA interference

ROS: Reactive Oxygen Species

SAR: Stramenopila Alveolata Rhizaria

SDV: Silica Deposition Vesicles

TCA: TriCarboxylic Acid cycle

TF: Transcription Factor

TIO: Transcription-Independent Oscillator

TTFL: Translational Feedback Loops

XC: Xanthophyll Cycle

Y2H: Yeast two Hybrid assay

ZT: *Zeitgeber* Time
<figure><img src="image_8.png" /></figure>
*The winds, the sea, and the moving tides are what they are. If there is wonder and beauty and majesty in them, science will discover these qualities. If they are not there, science cannot create them. If there is poetry [...] about the sea, it is not because I deliberately put it there, but because no one could write truthfully about the sea and leave out the poetry.*

Rachel Carson<fnref n="1" />

<footnote n="1">¹ From Rachel Carson's acceptance of the National Book Award for Nonfiction, 1952 (“Rachel Carson Excerpts Continued - Rachel Carson - U.S. Fish and Wildlife Service,”)</footnote>
# Chapter I:
## Introduction

This introduction chapter is organized in five main sub-parts: in the first one, the life of phytoplankton in the oceans and their role in the biogeochemical cycles at planetary level are briefly presented.

The second part focuses on the general presentation of diatoms, the group of algae on which this thesis work focuses. Their cell biology, their physiology and their role in the global ecological context are described. Particular attention is given to the description of the model species *Phaeodactylum tricornutum* that has been the subject of study during my thesis.

The third section gives an overview on light-dependent or light-regulated biological processes in diatoms. These processes include photosynthetic activity, photoacclimation, photoperception and cell division.

An in-depth description of biological rhythmicity and biological clocks is presented in the fourth section. The molecular architecture and function of biological timekeepers is given, with a comparative depiction of their molecular components in different taxa.

Finally, the fifth part represents an overview of transcription factors in diatoms. Particular space is dedicated to the transcription factors of the bHLH-PAS family, on which I have focused my research in *P. tricornutum*.
I.1 The life of phytoplankton

"How inappropriate to call this planet
'Earth' when it is quite clearly 'Ocean'"<fnref n="2" />. With
skill and poetry, the renowned science fiction
author and undersea explorer Arthur Charles
Clarke perfectly synthesized the vastness of the
ocean and its importance for all lifeforms. The
world ocean covers approximately 70% of the
surface of our planet and has a variety of
habitats and environmental conditions
comparable to, if not greater than, that of dry
land. Life has emerged in the oceans, which
are today home to an endless variety of living
organisms of all shapes and sizes ranging from
the smallest bacteria to the blue whale. Still,
the vast majority of life underwater is
represented by microbes, with around 70% of
the marine biomass being represented by
bacteria and protists (Bar-On et al., 2018).
These microorganisms, along with
microscopic free floating animals, are
collectively called plankton and represent the
beating heart of the trophic web of the oceans
(Pierella Karlusich et al., 2020). Meta-genomic
analysis of samples extracted in the wild reveal
that the number of eukaryotic planktonic
species ranges between the hundreds of
thousands to the tens of millions (de Vargas et
al., 2015). Of this teeming and kaleidoscopic
microscopic biodiversity, only a few thousand
species have been classified to date, leaving to
our imagination the wonders yet to be
discovered below the surface of water.

The photosynthetic fraction of these
organisms is called phytoplankton, also
commonly referred to as microalgae, a term
that encompasses a variety of unicellular
organisms that are extremely diverse and also
phylogenetically very distant from each other
(Blaby-Haas and Merchant, 2019; Keeling,
2013; Nowack and Weber, 2018).
Phytoplankton proliferation is inextricably
reliant on light irradiation, which is the source

of energy for photosynthesis and the
metabolism of these organisms. Because of
this, phytoplankton is fundamentally found
only in the photic zone, the most superficial
fraction of the water column where the sun's
rays can penetrate (approximately up to 200 m
depth). Yet, by converting solar energy and
atmospheric CO₂ into organic matter,
phytoplankton is of critical importance for the
bio-geo-physical processes that regulate the
functioning of the biosphere (Pierella
Karlusich et al., 2020). Eukaryotic marine
algae and prokaryotic cyanobacteria are
indeed accounted for circa 45% of the global
Net Primary Production (NPP) or, in other
words, for half of the total flux of carbon
fixation from the atmosphere to organic
compounds (Field et al., 1998). If terrestrial
plants are commonly called the lung of the
earth, phytoplankton can rightfully be
honored with the same title.

Remarkably, while contributing so
much to NPP on a planetary scale,
phytoplankton contribute with only one
Gigatonne of carbon (Gt C, where 1 Gt C =
10¹⁵ g of carbon) to the global living biomass, a
tiny fraction in comparison to the estimated
470 Gt of higher terrestrial plants (Bar-On et
al., 2018). This seemingly paradoxical
disproportion is possibly explained by the high
reproduction turnover of these organisms.
Despite representing a relatively small fraction
of the global biomass at any given moment,
many more generations follow in a short space
of time in comparison to slow-growing, slowly
reproducing multicellular organisms as plants.
As these organisms die, they generate organic
matter which precipitates on the seabed. Dead
organic matter transported to the deep ocean
by gravitational sinking is sequestered on
timescales of thousands of years before being
recirculated to the surface via the slow natural

<footnote n="2">² See (Rozwadowski, 2010)</footnote>
<figure><img src="image_9.png" /><figcaption>Figure 1. Chlorophyll distribution in the world ocean</figcaption></figure>

Global distribution of chlorophyll in the oceans detected via Moderate Resolution Imaging Spectroradiometer by the Aqua MODIS Satellite (NASA). The image represents the composite of Chlorophyll concentration over a period ranging from July 4, 2002 and December 31, 2018. Image retrieved from the OceanColor Web tool, NASA (https://oceancolor.gsfc.nasa.gov/)

movements of deep-water masses. A fraction of this sedimented primary production (~0.3% of marine NPP) will eventually form the major long-lasting reservoirs of carbon that will persist for millions of years in the form of sedimentary rocks (Zhang et al., 2018). This myriad of photosynthetic organisms inhabiting the photic zone also becomes the prey of heterotrophic grazers, viruses and microbial decomposers, thus representing the base that supports the entire marine trophic web. (Bar-On et al., 2018; Zhang et al., 2018). The rapid rate of reproduction of planktonic phototrophs also means that, in marine ecosystems, primary producers can sustain a population of consumers five times higher in terms of biomass, a ratio between trophic levels opposite to that found on dry lands (Bar-On et al., 2018). This flux of CO₂ from the atmosphere to the oceans through photosynthetic fixation, trophic web functioning and sinking of dead matter is named Biological Carbon Pump (BCP). Its ecological relevance is evident when we consider that, thanks to BCP, oceans are held

responsible for the reabsorption of about 26% of anthropogenic CO₂ emitted since the industrial revolution (while land vegetation stocked 34% of it and the remaining fraction represents the net atmospheric increase of CO₂) (Le Quéré et al., 2018).

In order to thrive to this extent, phytoplankton necessarily had to adapt to the light environment of the oceans, an environment that is anything but constant and uniform. Due to the spectral properties of water, light intensity quickly decreases in the first meters of the water column, so that direct light irradiation is orders of magnitude lower in the uppermost layers of seawater than on land (Depauw et al., 2012). In addition, water absorbs light extremely unevenly depending on its wavelength, with red light being completely extinguished in the very first meters, while blue light manages to penetrate much deeper. In this way, the light profile of the water column is very different from that which can be found on land, where the air absorbs the different wavelengths more
uniformly. Moreover, because of this phenomenon, the chromatic profile of light changes extremely rapidly as a function of depth, making the marine environment highly inhomogeneous and mainly dominated by green/blue light after the first few meters below the surface (Depauw et al., 2012; Stomp et al., 2007). In addition to the physical properties of water, other factors intervene in making the oceans an ever-changing environment. Atmospheric conditions, water movements (e.g. tides, and streams) and light scattering caused by coloured inorganic and organic suspended matter are all factors that play a role in characterizing the underwater light field (Kirk, 2011; Stomp et al., 2007). Even the scattering and absorption of light by the living phytoplankton itself can contribute substantially to the attenuation of irradiation. In nutrient rich waters (like estuaries) or during seasonal algal blooms, the algae can reach concentrations sufficient to cause self-shading to limit their own growth (Kirk, 2011).

Light quality and intensity perceived by marine organisms also dynamically change in function of time. The rotation of the Earth imposes the well-known constant ~24 hours cycles of light and dark since the beginning of life. Day length (photoperiod) also varies along the year as a function of latitude as a consequence of the revolution of the Earth around the sun and the inclination of its rotation axis, generating additional seasonal 365 days cycles. Finally, it should not be forgotten that reflected light from the moon, although not a relevant energy source for photosynthetic activity, is nevertheless a highly important source of information on the

environment. The ~24.8 hours periodicity of moonrise and moonset in combination with the ~29.5 days period of moon phases create a complex, but cyclical and predictable, pattern of illumination of the earth's surface intertwined with solar rhythms (Andreatta and Tessmar-Raible, 2020). To note, lunar cycles also play a crucial role in the environmental dynamics of coastal environments, as they are responsible for tidal rhythms and associated processes (e.g. the cyclic immersion of intertidal zones), profoundly shaping life in these ecosystems which are among the most productive and biodiverse in a marine context (Cahonn, 1999).

Light is not the only fundamental factor for the life of phytoplankton. Equally crucial is the availability of nutrients such as carbon, nitrogen, phosphorus, silicon or iron, (Speight and Henderson, 2010). The scarcity of these elements is one of the main limiting factors for the proliferation of phytoplankton, especially far from the coasts, where instead the runoff of elements from the mainland or upwelling phenomena of deep waters can lead to a local increase in the availability of nutrients (Lassiter et al., 2006; Speight and Henderson, 2010, Fig.1). If we add to this the fact that other factors such as salinity or temperature are part of the dynamic landscape of biotic and abiotic elements that regulate life in the oceans (Mann and Lazier, 2009) it is clear that, in order to proliferate, phytoplankton must be able to respond quickly and efficiently to all these stimuli in order to maximize their fitness. Among the various groups of phytoplankton, diatoms clearly emerge as one of the most successful.
# I.2 The kaleidoscopic world of diatoms

## I.2.a General introduction on diatoms

Diatoms are unicellular photosynthetic eukaryotes representing one of the most abundant, diversified and widely distributed groups of phytoplankton. Far from being relegated to oceanic environments, these cosmopolitan algae are distributed at all latitudes and in all types of habitats and can be found in any body of water, whether saltwater or freshwater, as well in soil moist (Falciatore et al., 2020). Diatoms are particularly relevant in polar regions (both arctic and antarctic), where they represent the most abundant group of photosynthetic eukaryotes (Malviya et al., 2016). In these regions, diatoms have also adapted to thrive and prosper in the apparently lifeless ice cap, where they reach such high concentrations to discolor the ice brown (Thomas and Dieckmann, 2002). Diatoms account for around 40% of marine carbon fixation, or 25% of the global NPP (Nelson et al., 1995; Tréguer et al., 2018), which is comparable to that of all terrestrial rainforests combined (Field et al., 1998). Estimations of diatom biodiversity are still uncertain, with the number of possible different species projected to be as high as 200,000 (Mann and Droop, 1996), although more recent studies consider this figure to be closer to 100,000 different taxonomic units (Malviya et al., 2016).

The life modes of diatoms are extremely varied. Depending on the species, they can live on the seabed (benthonic) or in suspension near the surface (pelagic or planktonic), they can appear as single isolated cells or grouped in colonies and in some cases they can establish close relationships of symbiotic interdependence with other microorganisms (Foster et al., 2011).

From a metabolic point of view, diatoms can be exclusive autotrophs (i.e.

totally dependent on photosynthetic activity for the production of carbohydrates) or mixotrophs (i.e. able to combine photosynthesis with the uptake of organic carbon sources dissolved in the environment) as in the case of *Phaeodactylum tricornutum* (Villanova et al., 2017) and also facultative heterotrophs as in the case of *Cyclotella cryptica* (Pahl et al., 2010). Although almost all diatoms are as mentioned photosynthetic, there are also known cases of species in which the plastid has lost its photosynthetic ability. These algae belong to the genera *Nitzschia* and *Hantzschia* and since most of the phylogenetically close species are photosynthetic, the loss of photosynthesis has clearly happened recently (Kamikawa et al., 2017).

Diatoms exhibit a rich variety of cellular and metabolic processes due in part to their complex evolutionary history and their composite genome, an assemblage of genes derived from different sources (the heterotrophic ancestor, photosynthetic symbionts, and prokaryotic donors, see Paragraph I.2.b). This genetic mosaic gives diatoms a variety of biological traits and processes that contribute in large part to their evolutionary success.

One of the most noteworthy characteristics of diatoms is their chiseled cell wall, called frustule (Fig.2), impregnated with hydrated glass (SiO2.nH2O, Kröger and Poulsen, 2008). Silicon is dissolved in marine waters mainly in the form of monomeric orthosilicic acid Si(OH)₄ which is converted to organic silica mainly by diatoms and, to a minor extent, by other protists like radiolarians and silicoflagellates (Tréguer et al., 1995). Due to these features, diatoms play a prominent role in oceanic biogeochemical
cycles of silica (Nelson et al., 1995; Tréguer et al., 1995). An estimate of 200-280 Tmol or biogenic silica is fixed each year by diatoms via their cell wall, of which approximately 50% precipitates to the seabed by gravitational sinking (Nelson et al., 1995). The importance of diatoms in the biogeochemical cycle of silica becomes glaring when we consider that,

according to projections, Si dissolved in the
ocean passes through diatom
biomineralization an average of about 39 times
before being deposited to the seafloor
(Tréguer et al., 1995). Once precipitated, the
glassy cell walls of diatoms can generate vast
deposits of siliceous sands, also called
diatomaceous earth or diatomite.

<figure><img src="image_10.png" /><figcaption>Figure 2. Diatom characteristics and morphological diversity</figcaption></figure>

Examples of different diatoms showing their morphological diversity. (A) Scanning Electron Microscopy (SEM) micrograph of *Skeletonema marinoi*. (B) Transmission Electron Microscopy micrograph of *Cyclotella atomus*. (C) SEM micrograph of *Fragilariopsis cylindrus*. (D) SEM micrograph of a mix of diatom species of the genus *Thalassiosira*. Light microscopy micrographs of (E) *Fragilariopsis kerguelensis*, (F) *Chaetoceros sp.*, (G) *Hemiaulus haukii* and (H-K) *Phaeodactylum tricornutum* in its three different morphotypes: tridadiate (H), fusiform (I and K) and oval (J). Images were kindly provided by Diana Sarno and Marina Montresor (Stazione Zoologica Anton Dohrn, Napoli, Italy)
Diatom frustule is composed of two parts, the smaller *hypotheca* and the slightly bigger *epitheca*, capable of overlapping in a manner often compared to the two halves of a Petri dish. Each one of these two halves will become the *epitheca* of the daughter cells after cell division, causing a progressive reduction in cell size generation after generation, in a phenomenon called MacDonald-Pfitzer rule (Macdonald, 1869; Pfitzer, 1869). In many diatoms, restoration of the original cell size involves sexual reproduction and the generation of an enlarged zygote named auxospore. The generation of this auxospore is induced by the reaching of a lower bound cell size threshold. It has also been shown that the artificial, abrupt reduction of cell size induces sexual reproduction, demonstrating that this process is not dependent on the age of the population (Chepurnov et al., 2004). Still, the molecular mechanisms involved in the perception of cell volume remain non clarified (for a general review on sexual reproduction in diatoms, see (Mann, 2011; Montresor et al., 2016). To note, centric and pennate diatoms show different mating strategies, the former being mostly homothallic (hence, self-fertile and not showing mating types) and the latter being heterothallic (hence requiring the coupling of cells of different mating-types for inducing meiosis, (Chepurnov et al., 2004)

The tri-dimensional structure of the *thecae* (Fig.2) is characterized by regular patterns of pores of variable dimension, whose geometry and spatial distribution is species-specific and under strict genetic control. Indeed, the biomineralization of silica and its regular deposition in diatom cell wall involve a variety of well characterized proteins including silaffins, frustulins, silacidins, pleurialins and cingulins (for a detailed overview on frustule structure, composition and morphogenesis see (Babenko et al., 2022). The biomineralization of silica occurs in large intracellular vesicles also named Silica

Deposition Vesicles (SDV) (Heintze et al., 2020).

The diatom glass armor performs several fundamental functions for the cell, including protection against mechanical shocks and against grazers like copepods (Hamm et al., 2003; Pančić et al., 2019). In addition, the porous frustule can also function as a natural filter possibly capable of blocking the passage of viruses and bacteria potentially dangerous to the alga (Losic et al., 2006). The periodic nanostructure of silica may also perform the function of a photonic crystal, capable of redirecting and redistributing light incident on the cell wall towards the interior of the cell (Goessling et al., 2018). The optical coupling between the frustule and the plastid could consequently allow an increase in photosynthetic efficiency, although to which extent remains an open question.

Another of the metabolic characteristics of diatoms is the fact that they possess a fully functional ornithine-urea cycle (OUC). This metabolic pathway was initially considered as a peculiarity of metazoans (animals) where it performs the task of ammonium disposal in urea, less toxic and more easily eliminated by the body (Mommsen and Walsh, 1989). The fact that genes involved in this metabolic cycle are found in brown algae and *Haptophyta* (another group of protists phylogenetically related to *Stramenopila*) suggests that OUC was inherited from their common ancestor (A. E. Allen et al., 2011). Unlike in *Metazoa*, the OUC does not perform a function of ammonium detoxification and amino acid catabolism. On the contrary, its main function is to capture the ammonium dissolved in the environment and to fix it in organic compounds, allowing the storage of the nitrogen in the form of arginine, ornithine and proline, the main osmolytes of diatoms (accounting for 70% of carbon and nitrogen accumulated in metabolites (A. E. Allen et al.,
2011). The OUC is therefore functional, in diatoms, to the rapid response to episodic cases of high availability of nitrogen (A. E. Allen et al., 2011), normally a limiting element in oceanic habitats (Falkowski et al., 2008, 1998). Since proline represents the main

precursor of polyamines (essentials for biomineralization of silica (Lin and Lin, 2018; Sumper and Kröger, 2004), OUC may also play a function in the production of the frustule (A. E. Allen et al., 2011).

<figure><img src="image_11.png" /><figcaption>Figure 3. The eukaryotic Tree of Life</figcaption></figure>

Phylogenetic tree representing the eukaryotic Tree of Life including all major supergroups. Red color highlights supergroups and taxa containing known examples of photosynthetic organisms with a plastid derived from a secondary endosymbiotic event with a red algal host. The *Archaeplastida* group, containing primary endosymbiotic phototrophs (including red and green algae and land plants) is highlighted in green. The *Stramenopila* *Alveolata* *Rhizaria* (SAR) supergroup is represented in detail to show the major taxa included in this branch and their relative phylogenetic position. Light red area highlights the position of diatoms within the *Stramenopila*. The tree is based on Dorrell and Smith 2011 and Burki *et al.* 2020.

## I.2.b Phylogenesis and evolution

In order to fully understand the biology of diatoms and the reasons for their ecological success, it is essential to know their evolutionary history and their position on the eukaryote Tree of Life (eToL, Fig.3). Diatoms are photosynthetic protists belonging to the Stramenopiles group, also referred to as Heterokonts. Stramenopiles fall in the larger SAR supergroup, a branch of the eToL which takes its name from the three subgroups that compose: *Stramenopila* themselves, *Alveolata* and *Rhizaria*, which derive from a single

common ancestor (Burki et al., 2020). Besides diatoms, several other organisms of utmost importance for marine ecosystems belong to SARs, such as *foraminifera*, *radiozoa* (both belonging to *Rhizaria*) and dinoflagellates (a mixed group of photosynthetic and heterotrophic unicellular organisms belonging to Alveolates). Of interest, dinoflagellates are the only family of microorganisms capable of competing with diatoms in terms of biodiversity abundance in oceanic ecosystems (Pierella Karlusich et al., 2020).
Stramenopiles encompass both unicellular heterotrophs (such as the water molds oomycetes) and a variety of photosynthetic organisms, including the multicellular brown algae *Phaeophyceae*, the unicellular *Eustigmatophyceae* and, obviously, diatoms (*Bacillariophyta*, Blaby-Haas and Merchant, 2019; Burki et al., 2020; Keeling, 2013). Diatoms first appeared ~190.4 millions of years ago (Mya, Fig.3 and Fig.4), close to the transition between the Triassic and Jurassic periods (Nakov et al., 2018). Diatoms are commonly divided into two main groups, clearly distinguishable by their shape: centric diatoms (*Coscinodiscophyceae*) with radial symmetry, the most ancient, and the fusiform, bilaterally symmetrical pennate diatoms (evolutionarily more recent and derived from centric, Kooistra et al., 2007). The separation of these two groups can be traced back to approximately 150 Mya by means of comparative taxon phylogeny (Nakov et al., 2018). Pennate diatoms can in turn be subdivided in raphid (*Bacillariophyceae*) or araphid (*Fragilariophyceae*) according to the presence or absence of a raphe, a longitudinal slit through the cell wall that is involved in motility and first appeared ~120 Mya (Nakov et al., 2018). The raphe is aligned with actin-myosin complexes responsible for the secretion of mucilaginous secretions used by these microalgae to actively slide on their substrate (Poulsen et al., 1999). To date, raphid diatoms are the most abundant, diversified and cosmopolitan. This ecological success is believed to be due in part to the evolutionary advantage given by the ability of active movement (Nakov et al., 2018).

Undoubtedly, one of the most extraordinary aspects of diatom evolutionary history is the complex origin of their plastid. Photosynthesis arose at least 3.5 billion years ago in bacteria as anoxygenic photosynthesis. The subsequent evolution of oxygenic photosynthesis in cyanobacteria promoted the generation of O₂ in the atmosphere (Nowack and Weber, 2018). Subsequently, a

photosynthetic cyanobacterium (endosymbiont) was engulfed by a heterotrophic eukaryote host, giving rise to the progenitor of eukaryotic plastids and to the evolution of the *Archaeplastida* group, which includes *Rhodophyta*, *Glaucophyta* green algae (*Chlorophyta*) and *Viridiplantae* (land plants, Fig.4, Keeling, 2013). This event of primary endosymbiosis is considered to be monophyletic and dates back to almost one billion years ago (Baldauf, 2008; Keeling, 2013). The only known example of independent primary endosymbiosis of a photosynthetic prokaryote is given by the amoeboids of the genus *Paulinella*, that engulfed a cyanobacterium between 65 and 90 Mya (Delaye et al., 2016). Hence, the entire diversity of plastids found in the eToL can be considered to have a single common origin. However, diatoms are not directly related to *Archaeplastida* and belong, as seen, to a distinct branch of the eToL (Fig.3). Phylogenetic and comparative genomics studies suggest that diatoms' ancestors acquired their photosynthetic organelle by means of secondary endosymbiosis, the process of engulfment and retention of a primary eukaryotic algae. In detail, the analysis of host genes transferred into the diatom genome suggests that they underwent at least two serial secondary endosymbiotic events of this kind with, in order, a green and then a red algae (Fig.4, Dorrell et al., 2017; Dorrell and Smith, 2011; Moustafa et al., 2009). The details and timing of this process and the real contribution of the green and red algal symbiont to diatom genomes remain debated (e.g. see Dorrell et al., 2017; Moustafa et al., 2009; Strassert et al., 2021). Still, it is important to keep in mind that secondary (and tertiary) endosymbiosis processes are responsible for plastid acquisition by all other non-*Archaeplastida* phototrophs, including other members of the SAR (e.g. dinoflagellates), but also photosynthetic haptophytes and excavata (for a detailed state of the art on the subject, see
<figure><img src="image_12.png" /><figcaption>Figure 4. The evolution of diatoms</figcaption></figure>

Schematic representation of the main evolutionary events that led to the emergence of diatoms. The monophyletic primary endosymbiotic engulfment of a cyanobacterium (in blue) by a heterotrophic eukaryotic host led to the apparition of the *Archaeoplastida*. The blue arrow shows the transfer of part of the cyanobacterium genes to the nucleus of the host. This supergroup contains *Glaucophyta* (light blue), red algae (Rhodophyta, light red) and the photosynthetic organisms of the green lineage that include green algae as well as land plants. Secondary endosymbiotic event of a red algal led to the acquisition of plastids in the photosynthetic *Stramenopila* ancestors of diatoms, as well as in variety of organisms including *Chryptopyta*, *Haptophyta* and part of the photosynthetic *Alveolata* (which altogether form the *Chromalveolata* group). The cryptic secondary endosymbiosis with a green alga predating the engulfment of the *Rhodophyta* endosymbiont is also represented. The evolutionary details of these endosymbiotic event are still disputed (see Stiller et al 2014). Endosymbiotic gene transfer of the red and green algae to the nucleus of the eukaryotic host is represented by the red and green arrows. The horizontal gene transfer from bacteria (in purple) is also represented. The time of emergence of centric diatoms from the *Stamenopila* ancestor, as well as the time of separation of araphid pennate diatoms from centric ones is reported accordingly to Nakov 2018. The time of evolution of the raphe and consequent emergence of raphid pennate diatoms is equally reported. Mya: millions of years ago.
(Burki et al., 2020; Dorrell et al., 2017; Nowack and Weber, 2018; Stiller et al., 2014)). The secondary endosymbiosis origin of diatom plastid is further supported by the fact that diatom chloroplasts are bounded by a total of four membranes, the two innermost deriving from the envelope of the plastid of the primary alga and the outer ones descending from the plasma membrane of the endosymbiont plus the phagotrophic membrane of the host cell (Kroth and Strotmann, 1999). In addition, the phenomenon of endosymbiosis left obvious marks in the genetic makeup of diatoms, as a portion of the endosymbiont genes were transferred into the nucleus of the acceptor host (Moustafa et al., 2009). Such a process is termed Endosymbiont Gene Transfer (EGT) and is estimated to be responsible for a significant proportion of the protein-coding genes in diatoms (about 16% according to (Moustafa et al., 2009), of which a portion is derived from the endosymbiont red alga and a portion from the green alga cryptic symbiont (Dorrell et al., 2017; Moustafa et al., 2009).

Horizontal Gene Transfer (HGT) from prokaryotes donors is another process that has largely contributed to the complexity of the genomes of diatoms (Vancaester et al., 2020) and of other SAR protists (Chan et al., 2012; Nosenko and Bhattacharya, 2007). Also named lateral gene transfer, HGT is the transfer of genetic information between different species by a process of direct passage and integration of genetic material. In diatoms, bacterial HGT contributed to an average of 3-5% of the genetic pool through multiple transfer events spanned over time and with different prokaryotic donors (Vancaester et al., 2020). The genes obtained by these

processes contribute to processes such as cobalamin uptake, cold tolerance and metabolism of carbon and nitrogen (Vancaester et al., 2020). Notably, several enzymes and metabolic shunts associated with ornithine-urea cycle in diatoms are supposed to derive from HGT from bacterial donors (A. E. Allen et al., 2011). The integration of viral genomic material, although observed, has occurred to a lesser extent in diatom genomes than what has been observed in other eukaryotes (Hongo et al., 2021).

Regarding the more general characteristics of diatom genomes, they show variable ploidy. In fact, some of them are characterized by a diploid genome with limited divergence between the two haplotypes (e.g. *P. tricornutum* (Bowler et al., 2008) and *Thalassiosira pseudonana* (Armbrust, 2004)), in others the two haplotypes show a strong divergence (e.g. *Fistulifera solaris* (T. Tanaka et al., 2015)) while still others are allopolyploid, as in the case of *Fragilariopsis cylindrus* (Mock et al., 2017). Diatom genes are usually short (<2,500 bp) and with one intron, only 20% of them possessing two or more introns (Basu et al., 2017). Despite the low complexity of gene structure, alternative splicing remains possible, with for example 15% of the genes of *P. tricornutum* undergoing this process (Rastogi et al., 2018). Diatoms have also been shown to transcribe non-protein-coding RNAs such as long intergenic non-coding RNAs (lincRNAs (Basu et al., 2017)), small non-coding RNA (sRNAs, (Rogato et al., 2014) and RNAs transcribed from the antisense strand of protein coding genes (NATs: Natural Antisense Transcripts (Cruz de Carvalho and Bowler, 2020)).

## I.2.c Toolkit box for molecular biology in diatoms

Our ability to understand the complexity of the processes that govern life in the ocean depends irrevocably on the

availability of model organisms capable of representing the enormous variety of taxa that inhabit it and of genetic manipulation
techniques essential for their study. For this
reason, the scientific community has made
great efforts to fill these gaps, leading to the
increasing availability of manipulation
techniques in an ever-increasing number of
organisms, most notably of protists (Faktorová
et al., 2020). Concerning diatoms, the
characterization of their cellular physiology
and molecular biology has seen a major
propulsion in the past two decades thanks to
the development of molecular and genomic
resources, as well as genetic manipulation
techniques. Over time, the community of
diatomists has therefore chosen and adopted
a variety of different species to characterize
according to the different focus of interest of
the research. Recently, this variegation of
study models has been reflected in the
genomic and transcriptomic sequencing of
several reference species, each chosen for its
physiological and metabolic peculiarities or
ecological relevance: e.g. *T. pseudonana*
(Armbrust, 2004) and *P. tricornutum* (Bowler
et al., 2008), chosen as first reference models
for centric and pennate diatoms respectively;
*T. oceanica* (Lommer et al., 2012) as a model
for the adaptation to iron-limited
environments; *Pseudo-nitzschia multistriata*
(Basu et al., 2017) chosen for its well
characterised sexual reproduction and
meiosis; *F. cylindrus* (Mock et al., 2017)
chosen as an ecological model for its
adaptation to the cold water and distribution
in the antarctic ocean; *S. robusta* as a reference
for the adaptation to a benthic life-style and its
rhythmic physiology (Bilcke et al., 2021a,
2021b); *Pseudo-nitzschia multiseries* (Basu et
al., 2017) and *Skeletonema costatum* (Ogura
et al., 2018) for their health relevance as toxic
algae; *F. solaris* (T. Tanaka et al., 2015) and
*Cyclotella cryptica* (Traller et al., 2016) for
their mixotrophic metabolism and for their
interest in biotechnological applications.

In addition to genomic and
transcriptomic information, different nuclear
transformation strategies are available for
different species (Falciatore et al., 2020). P.

*tricornutum*, for example, can be transformed
by biolistic bombardment (Apt et al., 1996;
Falciatore et al., 1999; Siaut et al., 2007), and
nuclear electroporation (Huang and
Daboussi, 2017; Niu et al., 2012; Zhang and
Hu, 2014). Similarly, nuclear transformation is
also available for the centric diatom model *T.
pseudonana* (Poulsen et al., 2006) and for less
developed models such as *Cylindrotheca
fusiformis* (Fischer et al., 1999; Poulsen and
Kröger, 2005), *Chaetoceros spp.* (Miyagawa-
Yamaguchi et al., 2011), *Fistulifera spp.* (Muto
et al., 2013), *Navicula saprophila* and *C.
cryptica* (Dunahay et al., 1995). Biolistic
bombardment and electroporation can lead to
the integration of multiple copies of
exogenous DNA into the genome, adding an
additional layer of variability between
transgenic lines (Karas et al., 2015; Sharma et
al., 2018).

Bacterial conjugation (the passage of
plasmids from an *Escherichia coli* bacterial
donor to the target cells via a conjugation
pilum) is another method that has been
proven to be efficient in both *P. tricornutum*
and *T. pseudonana* (Karas et al., 2015). The
advantage of this technique is that it allows the
introduction of an episomal transgene vector
in the form of a plasmid into the cell without
integration into the genome (episomal vector),
differently from what happens by
microparticle bombardment or
electroporation. This avoids the random
integration of exogenous DNA that could
consequently interrupt endogenous sequences
either in coding or regulatory regions (e.g.
promoters) and ensures greater genetic
uniformity among mutants (George et al.,
2020). Furthermore, since the vector is
maintained inside the cells by antibiotic
resistance, the conjugation system also allows
the reversibility of the transformation, since
the culture in the absence of antibiotics leads
to the elimination of the episome. A final
advantage of this technique is that it allows one
to control the number of copies of the vector,
since the episome usually occurs in a single
copy for each cell.

It is possible to modulate the
expression of genes of interest by coupling the
transgene to a large variety of different
promoters. These promoters can be
constitutive (constantly expressed at a stable
level, as in the case of the Histone 4
promoter), light responsive (as in the case of
the promoter of the Fucoxanthin-Chlorophyll
binding Protein FCPb) or even inducible by
nutrient availability cues (as in the case of the
Nitrate Reductase NR promoter, activated by
the presence of NO₃⁻ in the culture medium)
(Apt et al., 1996; Falciatore et al., 1999;
Hempel et al., 2009; Poulsen et al., 2006; Siaut
et al., 2007). Transformation also allows the
expression of proteins coupled with Human
influenza hemagglutinin (HA), fluorescent
protein moieties or other tags (Siaut et al.,
2007) and the expression of reporter genes
like aequorin and fluorescent proteins
(Falciatore et al., 2000; Karas et al., 2015;
Poulsen et al., 2006; Siaut et al., 2007;
Zaslavskaja et al., 2001b, 2001a). Gene
expression can also be manipulated via gene
knockdown silencing through RNAi (De Riso
et al., 2009) and artificial miRNAs (Kaur and
Spillane, 2015).

Targeted gene editing for gene
function obliteration is also established in
diatoms, using domesticated Transcription
activator-like effector nucleases TALENs
(Daboussi et al., 2014) and CRISPR/Cas9
(Nymark et al., 2016). Recent works have also
implemented the direct biolistic
bombardment of the Cas9 ribonucleoprotein
in *P. tricornutum* cells. This innovation allows
the transient and well regulated mutagenic
activity of the CRISPR/Cas9 method, avoiding
all the possible side effects of the random
genomic insertion of the Cas9 vector and of
the persistent activity of the protein after the
transformation (Serif et al., 2018).

To conclude, the possibility of
expressing exogenous genes in the chloroplast
after transformation by electroporation has
been demonstrated (Xie et al., 2014), as well
as targeted mutagenesis of plastid genes by
homologous recombination (Materna et al.,
2009). Nevertheless, these techniques of
chloroplast transformation remain unrefined
and poorly used by the community of
diatomists because of the limited
reproducibility of the available protocols
(Kroth et al., 2018).

I.2.d *Phaeodactylum tricornutum* as a diatom model

Given the very high diversity of
species, life strategies, and complex
evolutionary history, the selection of models is
particularly challenging in diatoms.
Historically, the earliest and best characterized
models in molecular biology were
*Thalassiosira pseudonana* (the first diatom
whose genome has ever been sequenced
(Armbrust, 2004)) and *Phaeodactylum
tricornutum* (Bowler et al., 2008), chosen as
reference species for centric/planktonic
diatoms and pennate diatoms, respectively. *P.
tricornutum* can occur in three distinct
morphotypes: oval, fusiform, and triradiate.
Although the three-horned form is the one

that gives the species its Linnean name, the
most common morphotype under laboratory
conditions in liquid cultures is the fusiform
one (de Martino et al., 2007). Even though the
genus *Phaeodactylum* is not of ecological
prominence (Armbrust, 2004; Malviya et al.,
2016), this has been isolated in a variety of
environments. In fact, this species can be
found at tropical latitudes as well as near the
Arctic Circle and at all longitudes (de Martino
et al., 2007). This characteristic, as well as the
availability in the laboratory of different
ecotypes with various origins, make it of
particular interest for the study of adaptation
to the environment. Recent sequencing of the
genomes of the ten *P. tricornutum* ecotypes was made available (Rastogi et al., 2020) and showed relatively low genetic diversity among different accessions. All my thesis work has been carried out on *P. tricornutum*, chosen for being the diatom species for which the characterisation of molecular and cellular biology and physiology is most advanced. This makes it an ideal point of access for the acquisition of knowledge that can be translated to other diatom species in the future. In addition *P. tricornutum* possess a small genome (27.4 Mb) and its gene pool (in particular of the set of transcription factors) has been studied in detail (Ait-Mohamed et al., 2020; Bowler et al., 2008; Rastogi et al., 2018; Rayko et al., 2010). Finally, a variety of techniques of genetic manipulation is available for this alga (see previous paragraph for details). The wild type strain used as reference for all my work is *Pt1*, an ecotype from

temperate regions (Blackpool, UK, 54°N). *Pt1* was also the ecotype chosen for the first *P. tricornutum* genome sequencing project (Bowler et al., 2008).

Also, *P. tricornutum* has the peculiarity of being the only known diatom species which does not peremptorily require silicon uptake nor the synthesis of the frustule (de Martino et al., 2007). The analysis of *P. tricornutum* genome perfectly reflects the mixed and highly complex origin of diatom genetic heritage. Genome annotation predicts a total of ~12,200 genes, ~9% of which show no homologous sequences in any other organism, 12% are shared only with diatoms, 4% genes originated from independent HGT events with prokaryotes, 3% are derived from the red algal ancestor of the plastid and 9% are related to genes specific of the green lineage of the *Archaeplastida* (Bowler and Falciatore, 2019; Rastogi et al., 2018).
I.3 Light in the life of diatoms

I.3.a Light as an energy source: the photosynthetic apparatus in diatoms

For phototrophic organisms such as
diatoms, light is the main energy source for
sustaining cellular metabolism. The
conversion of light energy into electrochemical
potential occurs, as known, at the level of the
thylakoid membrane of the chloroplast. Given
the peculiar evolutionary history of the plastid
of *Stramenopila* (presented in Paragraph
I.2.b), a description of this organelle is
indispensable for understanding the
characteristics of photosynthesis in diatoms.
As already aforementioned, diatom
chloroplasts are bounded by four membranes
deriving from the plastid and the plasma
membrane of the endosymbiont plus the
phagotrophic membrane of the host cell.
These four membranes are at the center of a
well-regulated traffic of proteins synthesized in
the cytoplasm and targeted to the chloroplast.
This protein traffic is necessary since only a
fraction of the proteins active in the
chloroplast are coded in the genome of the
organelle, while the large part has been
transferred to the nuclear genome. The
nuclear-encoded proteins targeted to the
plastid are characterized by a bipartite N
terminal signal sequence constituted of a
signaling peptide followed by a chloroplast
transit peptide (Kilian and Kroth, 2004). The
presence of a conserved amino acid sequence
(ASAFAP) at the level of this targeting
sequence (Kilian and Kroth, 2004) allows a
bona fide identification of the plastid targeted
proteins in diatoms (Gschloessl et al., 2008).
Furthermore, unlike the better characterized
green line plastids, diatom thylakoids are not
divisible into grana and thylakoid plastids, but
are stacked in groups of three thylakoids
spanning the entire length of the organelle
(Flori et al., 2016; A. Tanaka et al., 2015).

Another characteristic of diatoms is
the fact that they lack lutein and α-carotene,
the carotenoids typically present in
*Archaeplastida* (Grossman et al., 2004). Still,
diatoms are able to synthesize other
compounds of the carotenoid family,
specifically β-carotene, diatoxanthin,
diadinoxanthin and fucoxanthin (Fx). The
latter represents one of the most abundant
pigments in these algae and is a major
harvester of light (Bertrand, 2010; Dambek et
al., 2012; Kuczynska et al., 2015). Notably, the
characteristic brownish/golden color of
diatoms and related algae (e.g. *Phaeophyceae*)
is caused by the accumulation of large
quantities of Fx. Beside carotenoids, diatoms
also produce chlorophylls a (chl a) and c (chl
c) which, together with Fx, are responsible for
the capture of photons and their funneling to
the photosynthetic apparatus (Kuczynska et
al., 2015). The proper and efficient
functioning of these molecules is ensured by
their binding with special proteins of the
thylakoid membrane called LHC (Light
Harvesting Complexes) or, alternatively, FCP
(Fucoxanthin Chlorophyll binding Proteins)
(Lang and Kroth, 2001; Lepetit et al., 2007).
The precise spatial arrangement of pigments
in the LHCs ensures efficient and rapid energy
transfer from the Fx to the chl (Wang et al.,
2019). In addition, Fx has a broader
absorption spectrum than chl which allows it
to efficiently capture light for photosynthesis
even at depths where blue and green light are
predominant (Wang et al., 2019).

In the next paragraphs I will give an
overview of the photosynthetic apparatus in
diatoms. Due to the vastness of the subject and
the fact that this process has been historically
studied mainly in plants and green algae,
general information on photosynthesis valid
<figure><img src="image_13.png" /><figcaption>Figure 5. The photosynthetic Linear Electron Transfer in diatoms</figcaption></figure>

Graphical representation of the photosynthetic Linear Electron Transfer (LET) at the level of the thylakoid and of the main protein complexes involved in the process. The red arrow highlights the path followed by electrons along the LET from a molecule of water to the NADPH molecule. Black arrows highlight proton release due to water oxidation and their movements across the thylakoid membrane. The representation is based on Allen et al. 2011 and adapted so to include specificities of the photosynthetic apparatus of diatoms. Abbreviations: Photosystem (PS), oxidised mobile plastoquinone (PQ), reduced mobile plastoquinone (PQH2), cytochrome b6f (Cyt b6f), soluble cytochrome c6 (Cyt c6) ferredoxin (FD), flavodoxin (FLD), flavodoxin(ferredoxin)-NADPH reductase (FNR), ATP synthase (ATPase), Calvin Benson Bassham Cycle (CBBC). P680 and P700 indicate the chlorophyll a special pair of PSII and PSI respectively, while Mn₄CaO₆ indicate the Oxygen Evolving Complex at the luminal side of PSII.

also for diatoms are all derived from a small number of generalist publications (J. F. Allen et al., 2011; Eberhard et al., 2008; Taiz et al., 2015). Where the photosynthetic machinery of diatoms diverges from that of the *Archaeplastida*, bibliographic references will be punctually provided. A schematic representation of the photosynthetic machinery in diatoms is reported in Fig.5.

The capture of light by the LHC is only the first step of the photosynthetic process. These antenna complexes are located at the periphery of two protein systems (photosystem I, PSI, and photosystem II, PSII) where the actual conversion of light energy into oxidoreductive potential occurs

through the excitation of an electron. In short, photosynthesis can be described as the transport of electrons along a succession of molecular actors at the level of the thylakoid and against the oxidoreductive gradient in a process powered by light energy. The journey of these electrons begins at the level of the PSII. The peripheral antenna complexes capture the energy of a photon through their pigment and then funnel it via excitation energy transfer (EET) towards a pair of chl a molecules placed at the heart of PSII (the so called special pair or Reaction Center RC). The special pair of chl a is associated to two homologous proteins, D1 and D2, that form the core of PSII. Also, the special pair presents a peak of excitability at 680 nm, the reason
why the PSII is also called P680. Once excited
directly by incident light or via EET from the
peripheral antennae, the RC becomes a good
reducer and passes an electron to a chain of
electron carriers (pheophytin and a non-
mobile plastoquinone QA). The final
consequence of the excited electron transfer
within the PSII is the reduction of a mobile
plastoquinone (QB or PQ). In order to
recover the lost electron and to be excitable
again, PSII is able to oxidize water molecules
thanks to the Oxygen Evolving Complex
(OEC), located on the lumenas side of the
thylakoid membrane. By oxidizing two
molecules of H₂O the OEC is able to reduce
four times the P680⁺ cation, generating a
molecule of O₂ as a byproduct and releasing
four protons (H⁺) in the lumen of the
thylakoid. This process is also called
photolysis of water or water splitting, and is
made possible by a metalloenzyme core
containing four atoms of Mn and one of Ca,
coordinated with oxygen atoms.

After the uptake of two electrons plus
two protons coming from the stromal side of
the membrane, the reduced PQ (PQH₂) is
released within the thylakoid membrane
where it is able to move freely due to its
hydrophobic nature. The electrons
transported by PQH₂ are picked up by the
second protein complex of the photosynthetic
chain, cytochrome b6f (Cyt b6f), which
transfers them to a new, mobile transporter,
this time located at the level of the thylakoid
lumen. In green algae, as well as in plants, this
soluble electron transporter is predominantly
a plastocyanin, a molecule containing copper.
In case this element is scarce in the
environment, plastocyanin can be replaced by
an alternative transporter, cytochrome c6,
which binds electrons at the level of an iron
atom coordinated to a heme group. However,
in most diatoms cytochrome c6 is the only
soluble transporter used. To date, only a
restricted number of species such as
*Thalassiosira oceanica*, adapted to live in
marine environments particularly poor in

iron, produces a plastocyanin in addition to
the cytochrome c6 (Peers and Price, 2006). A
second consequence of the oxidation of PQH₂
at the level of Cyt b6f, in addition to the
reduction of the soluble transporter, is the
release of a proton in the thylakoid lumen
which is added to the four released during the
photolysis of water.

The soluble electron carrier brings
the electrons to PSI. Exactly like PSII, also
PSI has a special pair of chl a which can be
excited directly by light or by EET from
peripheral antennas. In this case, however, the
excitability peak of the RC is at 700 nm and
not at 680 nm. Also in this case, once excited,
the chlorophylls of the special pair release an
energized electron and the P700+ is
"recharged" via reduction by the soluble
carrier. Once energized, the electron again
passes through a chain of carriers associated
with PSI protein complex (in order: chl A, chl
A⁺, phylloquinone and Iron-Sulfur complex
Fx). The final electron acceptor in PSI is, in
*Archaeplastida*, an iron-containing molecule
called ferredoxin (FD). In diatoms, FD can be
replaced by an alternative electron acceptor,
flavodoxin (FLD), in case of iron deficiency in
the habitat. Indeed, FLD binds electrons via a
flavin group that does not require any metal
moiety, thus reducing iron dependence in
these algae (Groussman et al., 2015; La Roche
et al., 1996; Raven and Raven, 2019). Through
the enzyme flavodoxin(ferredoxin)-NADP(H)
reductase (FNR), the electron is shifted from
flavodoxin/ferredoxin to its final acceptor, a
molecule of Nicotinamide Adenine
Dinucleotide Phosphate (NADP) which is
reduced to NADPH after the reception of two
electrons and a proton sequestered from the
stroma. NADPH is the principal reducing
agent of the cell, used for a plethora of
biochemical processes including lipid and
nucleic acid biosynthesis and the Calvin
Benson Bassham cycle described later in this
section. Globally, the entire photosynthetic
process just described can be summarized as a
passage of electrons from water to NADPH according to the formula:

$$2H_2O + 2NADP^+ \rightarrow O_2 + 2H^- + NADPH$$

This transport of electrons through PSII, PQ, Cyt b6f, cytochrome c6 and PSI is also called Linear Electron Transfer (LET, Fig.5).

Importantly, in diatoms PSII and PSI turn out to be structurally isolated. PSII, in particular, is localized at the level of the central thylakoid, while PSI is placed at the level of the two outer thylakoids of the stack of three, together with the Cyt b6f (Flori et al., 2017). Since the thylakoids are in continuity, this configuration does not prevent the transfer of electrons from PSII to PSI via the mobile carrier PQH₂ (Flori et al., 2017). The advantage of the segregation between PSII and PSI is that it avoids the direct thermodynamic energy transfer (spillover) from PSII to PSI bypassing the intermediate complexes (Flori et al., 2017). Energy spillover is observable in red algae, where it can cause a relevant drop in the efficiency of photosynthesis (Biggins and Bruce, 1989).

Anyhow, the generation of reducing potential in the form of NADPH is only the first result of photosynthesis. Indeed, the accumulation of positive charges in the lumen of the thylakoid thanks to the photolysis of water at the level of the OEC, the activity of the Cyt b6f complex and the reduction of PQ and of NADP⁺ generates an electrochemical transmembrane potential. By exploiting this energetic potential, letting the protons move from the thylakoid lumen to the stroma, the ATPase complex phosphorylates ADP to obtain ATP. ATP will be then used as the energetic token used to power the metabolic processes of the cell, primarily the fixation of carbon in the form of sugars.

An important feature of the diatom chloroplast is its connection with the mitochondrion, so that the lumen of the two organelles, although functionally different, is in continuity (Bailleul et al., 2015). As in all

photosynthetic organisms, the ratio of NADPH to ATP generated by the operation of the LET in the thylakoid is unbalanced with respect to the energy requirements of the cell with, in particular, a surplus of reducing potential with respect to ATP produced (Bailleul et al., 2015). In order to balance the NADPH/ATP ratio, *Viridiplantae* have developed a series of redox processes ancillary to LET such as cyclic electron transport (CET) around the PSI or water-water cycles, which are however of minor importance in diatoms (Bailleul et al., 2015). In these algae this metabolic imbalance is resolved by the direct exchange of ATP, NADP(H) and O₂ between mitochondria and chloroplasts. This mechanism of ATP/NADPH ratio increase is also called meta-cyclic electron flow given the substitutive role with respect to CET (Bailleul et al., 2015; Zhou et al., 2020).

One of the main metabolic processes using the ATP and NADPH generated by the electron transfer machinery is the Calvin Benson Bassham Cycle (CBBC), sometimes called the dark phase of photosynthesis. This pathway is realized at the level of the stroma and consists, in short, in the fixation of CO₂ in the form of carbohydrates. Once the carbon accumulates as CO₂ or HCO₃⁻, it is moved to the stroma at the level of the pyrenoid. Pyrenoids are large aggregates of Ribulose-1,5-bisphosphate carboxylase-oxygenase (RuBisCO). The RuBisCO combines CO₂ molecules and a five-carbon sugar (ribulose 1,5 diphosphate) to form two molecules of a three-carbon product (3-phosphoglycerate, PGA), a process also called carboxylation. The reduction of PGA, involving NADPH and ATP (reduction phase of the CBBC) leads to the formation of glyceraldehyde 3-phosphate (G3P, two molecules generated per each CO₂ molecule entering the CBBC). For each 3 CO₂ entering in the CBBC, one excess molecule of G3P is produced and later used for the biosynthesis of sucrose, while the five remaining G3P molecules are on the contrary
reconverted to ribulose 1,5 diphosphate to
restart the process.

To conclude, it is useful to consider
how the concentration of dissolved CO₂ in the
oceans in the form of HCO₃⁻ (10-30 µM) is
insufficient to saturate the enzymatic activity of
RuBisCO at the pyrenoid level (Matsuda et al.,
2017). For this reason, diatoms, like other
photosynthetic organisms, have evolved
carbon-concentrating mechanisms (CCM) to
maximize CO₂ concentration at the level of the
thylakoid stroma to maximize CBBC
efficiency (Tsuji et al., 2017). The CCM
includes the use of a variety of bicarbonate
active transporters (at least ten have been
identified in model species) phylogenetically
related to the ones found in *Metazoa*. In
addition, CCM relies on the presence of

multiple Carbonic Anhydrases (CA) enzymes
able to convert CO₂ to HCO₃⁻ and vice versa.
Diatoms use CAs to favor the diffusive uptake
of CO₂ at the cell surface by converting HCO₃⁻
to CO₂ in the extracellular environment and
the sequestration of carbon by transforming
CO₂ back to HCO₃⁻ once it has entered the
cytosol. Finally, dedicated CA enzymes in the
pyrenoid convert HCO₃⁻ to increase the local
CO₂ concentration improving RuBisCO
activity (Matsuda et al., 2017; Tsuji et al., 2017;
Young and Hopkinson, 2017). Finally, a C4-
like photosynthesis, a pathway described in
plants (Sage, 2004) has been proposed to be
present in diatoms (Kroth et al., 2008). Still,
the existence of C4-like photosynthesis in
diatoms is controversial and has yet to be
clarified in diatoms.

I.3.b Photoacclimation and photoprotective mechanisms

If on the one hand light is
indispensable for the activation of
photosynthesis, on the other hand excessive
irradiation can be harmful to the chloroplast.
In strong light, in fact, the excitation of the
chlorophylls of the special pair can be too fast
to allow a proper uptake of electrons by the
acceptors downstream of the chain. The
inability of the RC chlorophylls to yield the
excited electron leads to the formation of the
triplet excited state of the chlorophyll (³Chl*),
which can in turn interact with O₂ generating
the highly reactive singlet oxygen form (⁴O₂)
(Müller et al., 2001). ¹O₂ in turn generates
additional reactive oxygen species (ROS) such
as superoxide oxygen (O₂⁻), hydrogen
peroxide (H₂O₂) and hydroxyl radical (OH⁻),
which can damage membrane lipids, hence
representing a relevant threat for the cell.
Given that the uptake processes downstream
of PSI are extremely rapid and difficult to
saturate, the generation of ³Chl* and
subsequent production of ROS are primarily
attributable to overexcitation of PSII.

In order to optimize the use of light to
fuel photosynthesis without it becoming
dangerous, diatoms, like all other
photosynthetic organisms, put into practice a
series of strategies to adapt their
photosynthetic apparatus according to the
intensity of radiation. These phenomena are
called photoacclimation (Dubinsky and
Stambler, 2009). In case of weak light, plastids
accumulate a greater amount of pigments so as
to maximize the optical density of the cell and
increase the capacity of photon absorption.
On the contrary, in case of high light, the
content per cell of pigments functional to the
absorption of photons such as chl and Fx
decreases so as to reduce the tendency to over
excite the photosystems (Anning et al., 2000;
Dubinsky and Stambler, 2009; Pniewski and
Piasecka-Jędrzejak, 2020). The net amount of
photosystems present on thylakoids also
decreases (Dubinsky and Stambler, 2009;
Taddei et al., 2018), but in return the
expression of associated proteins remains high
to compensate for the higher turnover of
photosystem degradation in high light
(Dubinsky and Stambler, 2009). To
counteract the increased production of ROS, cells also produce more antioxidant enzymes such as catalases, peroxidases, or superoxide dismutases (Dubinsky and Stambler, 2009). At high photon fluxes the cells also increase the amount of RuBisCO subunits per photosystem (Dubinsky and Stambler, 2009), which causes the photosynthetic activity and carbon fixation rate to saturate at higher light intensities (Anning et al., 2000; Schellenberger Costa et al., 2013a; Taddei et al., 2018). The increased energy availability at high light normally results in increased growth rates compared to what is observed at low light (Anning et al., 2000; Brand and Guillard, 1981, 1981; Pniewski and Piasecka-Jędrzejak, 2020), at least until the light stress becomes excessive (Brand and Guillard, 1981).

To avoid the occurrence of ³Chl* and its subsequent risks, photosynthetic organisms possess a variety of defensive processes that avoid overexcitation of the RC in PSII. Because these phenomena lead to a de-excitation of chlorophyll (which also occurs alternatively as a result of normal activation of LET in the photochemical process) these mechanisms take the name Non-Photochemical Quenching (NPQ) of chlorophyll fluorescence. The mechanisms of NPQ can be distinguished by the timing of activation and relaxation following the end of excess light irradiation (Eberhard et al., 2008).

A first, slow component of NPQ is the so-called photoinhibition (or qI), which consists in the degradation of the protein D1 of PSII that, as seen in the previous paragraph, interacts with the special pair of chlorophylls. The reversion of qI occurs through the resynthesis of functional forms of the D1 protein and the replacement of the damaged copy with the new one, a process that can take several minutes or even hours. In *Archaeplastida*, another form of slow NPQ is the state transition (qT), which involves the displacement of a portion of the peripheral antennas of PSII toward PSI, so as to balance

the excitation between the two photosystems and reduce the pressure on PSII. In diatoms, however, no form of qT has ever been observed, a fact that can be explained given the spatial segregation between the two photosystems in different thylakoid regions (Flori et al., 2017).

As for the fast components of NPQ, this photoprotective system is related to the dissipation of excess energy in the form of heat, it responds rapidly to changes in light intensity and is commonly called qE. In qE, a main role is played by the de-epoxidation of xanthophylls (i.e. oxygen containing carotenoids) in a reversible process named Xanthophyll Cycle (XC). In plants and green algae, the XC consists of the pH-dependent conversion from violaxanthin, with two epoxide groups, first to antheraxanthin (one epoxide group) and then to zeaxanthin (no epoxide). The de-epoxy xanthophylls are believed to favor the relaxation of chlorophyll by both directly absorbing the excess energy (that will be then released as heat) or by acting as allosteric regulators of the LHC (Müller et al., 2001). In particular, they might cause a conformational switch of the LHC that eases the autonomous de-excitation of chl via heat dissipation. In diatoms, the violaxanthin-zeaxanthin cycle is present, but an additional XC involving the de-epoxidation of diadinoxanthin (one epoxide) to diatoxanthin (no epoxide) (Lohr and Wilhelm, 1999) is the main contributor to the high photoprotective capacity of these algae (Blommaert et al., 2021). Interestingly, in *Archaeplastida*, the synthesis of de-epoxidized xanthophylls by the enzyme violaxanthin de-epoxidase (VDE) is strictly dependent on excessive acidification of the thylakoid lumen, one of the consequences of overloading the photosynthetic chain. In contrast, in diatoms, diatoxanthin epoxidation is repressed upon acidification of the thylakoid, but once the diatoxanthin concentration rises up NPQ capacity becomes independent on proton gradient (Goss et al., 2006).
Also, diatom fast NPQ relies on specific members of the LHC family specialized in photoprotection, the LHCXs, which act as qE effectors. This family of proteins has undergone a major expansion in diatoms. *P. tricornutum* genome for example contains 4 genes encoding for LHCX antennae (Taddei et al., 2016), a number that increases to 11 in the polar diatom *Fragilariopsis cylindrus* (Mock et al., 2017). LHCX genes are regulated in response to a variety of environmental signals, including light and nutrient deficiency (Taddei et al., 2016). In *P. tricornutum* a down regulation or obliteration of one of these proteins, LHCX1, affects significantly NPQ photoprotective capacity (Bailleul et al., 2010; Buck et al., 2019; Giovagnetti et al., 2022). LHCX proteins are phylogenetically related to LHCSR proteins from green algae, which are also involved in NPQ induction (Bailleul et al., 2010). The photoprotective effect of LHCSRs is activated following the protonation of some of their residues on the lumenal side and the consequent conformational change of the protein (Ballottari et al., 2016). The photoprotection mediated by LHCSR proteins is induced by the excessive acidification of the thylakoid lumen. However, diatom LHCX proteins do not present all the amino acids responsible for this process in the green lineage and lost a large portion of the C-terminal end immersed in the thylakoid

lumen, suggesting a different mode of activation (Bailleul et al., 2010; Giovagnetti et al., 2022). Still, it has been proved in some diatoms that the acidification of the thylakoid is necessary to induce the activation of the xanthophyll cycle and, therefore, of a component of NPQ (Chukhutsina et al., 2014; Goss et al., 2006). This suggests the existence of a proton concentration sensing system in the thylakoid. Although the details of LHCX protein activation remain to be elucidated, recent investigations have identified an amino acid domain containing a tryptophan near the site of interaction with xanthophylls that is involved in qE generation (Buck et al., 2021).

As a concluding remark, it is useful to note that the oxidation status of the PQ pool is involved in the regulation of LHCX gene expression (encoded in the nucleus) and in xanthophyll neosynthesis (Lepetit et al., 2013). The control of nuclear gene expression according to the level of PQ oxidation (also called retrograde signaling) is a phenomenon already observed in the past in *Archaeplastida* (Kleine et al., 2009), but the details of how this process occurs are still unclear in diatoms. In any case, this phenomenon makes evident the presence of a tight crosstalk between the plastid and the nucleus as well as their interdependence in phenomena of regulation of physiology.

### I.3.c Light as a source of information: photo-perception

Given the highly variability and dynamic of the photic environment in oceans, light represents a fundamental source of information for aquatic life. Light quality underwater changes in function of multiple parameters, such as water turbidity, depth, daytime and season among others. Organisms evolved a variety of proteins able to perceive light, the photoreceptors, of which most are composed of a non-protein moiety (the chromophore) associated with an amino acid

backbone. The activation of the photoreceptor is induced by the conformational changes in the tridimensional structure of the protein after the chromophore interacts with a photon of the correct wavelength. These proteins can therefore be categorized depending on the wavelength they are sensitive to, the nature of their chromophore and the protein domain composition (Jaubert et al., 2017).
Diatoms, in particular, possess a rich set of photoreceptors capable of detecting light throughout the whole visible spectrum (Jaubert et al., 2017). Red and Far-Red radiation is perceived by phytochromes which, in diatoms, show a structure similar to that of their bacterial counterpart and are commonly named DPH. Moreover, DPHs absorb at longer wavelengths of red and far-red light, compared to plant phytochrome (PHY). In *P. tricornutum*, it has been also shown that far-red light is the wavelength inducing signal transduction propagation (Fortunato et al., 2016). Photosynthetic organisms absorb light predominantly in red and blue, while reflecting most of the far-red. This means that the greater the density of phototrophs in the vicinity, the smaller the R:FR ratio. In terrestrial plants it has been thoroughly characterized how the ratio between irradiation in the red and far-red (R:FR ratio) sensed through phytochromes plays an essential role in the perception of surrounding phototrophs. Specifically, phytochromes play a central role in shade avoidance processes (Pierik and de Wit, 2014). Phytochromes of marine algae, including diatoms, could play a similar role in determining the presence of neighbor phototrophs and therefore the perception of cell density in the surrounding environment. However, the density of microalgae per unit volume remains very low overall, even during reproductive blooms, making this an unlikely possibility. It is important to recall that long wavelength light is quickly absorbed in the water column, making it tricky to hypothesize the possible biological role of these receptors in diatom biology. It is not to be excluded that red light perception might play a role in the regulation of biological rhythmicity, as it has already been largely demonstrated in plants (Devlin and Kay, 2000; Somers et al., 1998). Still, no direct evidence for such a function has been found yet.

Diatom genomes also show the presence of putative photoreceptors of the rhodopsin (RHO) family. Although no

characterisation of their function and absorption properties has been carried out, diatom RHOs could act as detectors of intermediate wavelengths (centered in the blue-green) or as photoreceptors that function as transmembrane ion pumps as in other organisms (Jaubert et al., 2017). Notably, diatom RHOs show high similarity to bacterial xanthorhodopsin, suggesting that their genes were acquired by HGT (Strauss, 2012).

Since blue light dominates below the sea surface and is the most penetrating in the water column, it is not surprising that diatoms (like other groups of phytoplankton) have multiple receptors for this wavelength (Jaubert et al., 2017). Photoreceptors of the Cryptochrome Photolyase Family (CPF) belong to this class of blue detectors. CPF proteins non covalently bind flavin adenine dinucleotide (FAD) as chromophore. FAD is reduced upon photon absorption, causing a conformational modification of the CPF protein and determining its capacity to interact with other protein partners. As their name suggests, CPFs include Photolyase proteins (PL), enzymes involved in blue light-activated DNA repair upon UV damage like cyclobutane pyrimidine dimer (CPD PLs) or (6-4) pyrimidine-pyrimidone photoproducts (6-4 PLs). Still, not all CPFs conserved this DNA repair activity, as in the case of plant cryptochromes which serve as blue light photoreceptors. Concerning diatoms, these algae possess animal like CPF 6-4 photolyases (Coesel et al., 2009), plant-like CRYs (CryP, only distantly related to canonical plant cryptochromes (Juhas et al., 2014) and several CRY-DASH (Drosophila Arabidopsis Synechocystis Human) proteins (Coesel et al., 2009; Fortunato et al., 2015). The latter are CPF proteins found in eukaryotes as well as in prokaryotes, whose function has not been fully characterized yet (Fortunato et al., 2015; Jaubert et al., 2022).

Contrary to green algae and plants, blue-detecting phototropins, containing LOV
and kinase domains, cannot be found in diatoms. On the contrary, diatoms possess other blue light photoreceptors with LOV domains known as aureochromes. Originally identified in the *Xanthophyta Vaucheria frigida* (Takahashi et al., 2007), photoreceptors of the aureochrome family are found only in *Stramenopila* and in the *Raphidophyta Heterosigma akashiwo* (Ji et al., 2017). Aureochromes consist of a light-sensing LOV domain associated with a bZIP domain involved in DNA binding and regulation of gene expression (Kroth et al., 2017; Mann et al., 2020). Specifically, it has been shown in *P. tricornutum* that approximately 75% of the genes respond to the transition from red to blue light by rapid (few minutes) down-regulation or up-regulation (Mann et al., 2020). This response is almost completely suppressed in KO mutants of *aureochrome 1a*, indicating the broad-spectrum impact that these blue receptors may have on gene transcription, possibly inducing a cascade of reactions mediated by other transcription factors. The role of aureochromes in gene regulation has recently been characterized in the *Eustigmatophyceae Nannochloropsis oceanica*, in which these photoreceptors appear to play an important role in fatty acid metabolism (Poliner et al., 2022).

Blue light photoreceptors can regulate the physiology of diatoms in response to environmental stimuli. First of all, blue light

plays a pivotal role in the regulation of photoacclimation and photosynthesis (Schellenberger Costa et al., 2013a; Valle et al., 2014) which is mediated by the above-mentioned photoreceptors. Indeed, in *P. tricornutum* CryP, CPF1 and aureochromes 1a and 1b are all involved in the expression of the protein complexes responsible of light harvesting and photoprotection (Coesel et al., 2009; Mann et al., 2020, 2017; Schellenberger Costa et al., 2013a, 2013b), see Paragraph I.3.a). Moreover, blue light is also central in the modulation of cell cycle progression and circadian rhythmicity. Cryptochromes are known to play a role in the circadian clock in animals (Kume et al., 1999; Putker et al., 2021), as well as in plants (Somers et al., 1998) and (potentially) in other secondary endosymbiont algae (Brunelle et al., 2007). In mammalian cells, the cryptochromes mCRY1 and mCRY2 are involved in the repression of CLOCK and BMAL1, positive elements of the transcriptional feedback loop of the circadian oscillator (Kume et al., 1999). Notably, the animal-like CPF1 of *P. tricornutum* is able to repress CLOCK:BMAL in simian *mCRY1* knockout cells, thus complementing the function of the lost endogenous cryptochrome (Coesel et al., 2009). This result suggests that CPF1 might play a similar role in the circadian oscillator in diatoms, although no evidence in this direction has been collected so far.

### I.3.d Cell cycle progression and regulation in diatoms

Diatoms are characterized by a high growth rate both under laboratory conditions and in natural environments (Sarthou et al., 2005; Spilling and Markager, 2008). Such a growth capacity plays an important role in the ecological success of these algae both at low latitudes and in polar environments (e.g. (Lafond et al., 2019; Smayda, 1980; Spilling and Markager, 2008). Understanding how diatoms regulate their cell division and how it

responds to environmental stimuli is essential to understanding their ecological success. The first studies on diatom cell division long precede the advent of molecular biology and date back to the end of the 19th century, with the pioneering studies of Robert Lauterborn (Lauterborn, 1896; Pickett-Heaps, 1991). The use of molecular analysis and advanced microscopy techniques (in particular transmission electron microscopy) have more
recently allowed us to characterize in more
detail the fascinating complexity of cell
division in these microalgae.

As in all other eukaryotes, the mitotic
cycle of diatoms includes a precise succession
of phases. The S phase (for DNA Synthesis) is
the one in which takes place the replication of
genomic material to be redistributed to
daughter cells; the M phase (for Mitosis),
where the actual duplication of the nucleus in
two copies, the redistribution in them of the
genome copies and, to end, the division of the
mother cell in two daughters (cytokinesis).
The S phase and M phase are separated by
two G phases (for Gap) in which the cell
performs its normal metabolic functions,
grows and prepares for division. These two G
phases are called G1 (which is between M and
S) and G2 (between S and M), respectively.

Such a complex process as the cell
cycle has to be regulated by a refined
molecular machinery. In eukaryotes, this
machinery involves proteins called cyclins
(CYC) and the CYC dependent protein
kinases (CDK), proteins that are highly
conserved throughout the eukaryotic kingdom
(De Martino et al., 2009; Huysman et al.,
2014b). The function of CDKs is to regulate
the activity and stability of target proteins
involved in the cell cycle by means of their
phosphorylation. The cyclins, for their part,
determine the substrate specificity of CDKs
and make them active by inducing their
conformational change. A classic example of a
process regulated by cyclin-CDK complexes is
the entry into S-phase associated with the Rb
(retinoblastoma) protein and E2F-DP
transcription factors, a mechanism conserved
in all eukaryotes except yeast (Giacinti and
Giordano, 2006). G1-phase-specific cyclin-
CDK complexes induce phosphorylation of
Rb, thereby interfering with its interaction with
E2F-DP. Freed from the inhibitory interaction

of Rb, the E2F-DP complex can initiate
transcription of genes involved in genome
duplication. In *P. tricornutum* it is
hypothesized that the complex formed by the
CDKA1 and dsCYC2 proteins is involved in
this G1-S transition (Huysman et al., 2013,
2014b). It is also important to note that the
proper activation of CDK proteins does not
only involve cyclins. Phosphorylation of
CDKs by CDK Associated Kinases (CAKs)
also plays a central role (Kaldis, 1999).
Equally, the timely proteolysis of CYCs,
CDKs and their targets is regulated by
appropriate E3 ubiquitin ligase complexes, the
Skp1/Cullin/F-box complex (SCF) and the
anaphase-promoting complex (APC). *P.
tricornutum* has been shown to have
conserved the genes coding for these
complexes (Huysman et al., 2014a).

Remarkably, comparative genomic
studies demonstrated that canonical cyclins
underwent an important process of
duplication in diatoms with a consecutive
expansion of this gene family, with a total of 13
canonical cyclin coding genes found in *P.
tricornutum* (Bowler et al., 2008; Huysman et
al., 2010, 2014b). In addition to the canonical
cyclins, diatom also evolved a complex set of
unique cyclins called diatom specific cyclins
(dsCYC) (Huysman et al., 2010). Different
CYCs and CDKs are expressed during well
determined moments of the cell cycle and
regulate the transition through the
aforementioned checkpoints, so that each
phase of the cell cycle can be associated to a
defined set of CYCs and CDKs (De Martino
et al., 2009; Huysman et al., 2010, 2014b).
The follow up of the expression of these
proteins (e.g. via reverse transcription
quantitative PCR) can be used as a proxy to
follow the progression of cells through the
different phases of cell cycle (Huysman et al.,
2010, Fig.6).
<figure><img src="image_14.png" /><figcaption>Figure 6. Diatom cell cycle and cyclins</figcaption></figure>

Schematic representation of the cell cycle. Arrows indicate the different phases (G1: Gap1, S: Synthesis of genome copy, G2: Gap2, M: Mitosis). The different cyclins possibly involved in the regulation of diatom cell cycle advancement are loosely arranged to correspond to the time of the cycle in which they are expressed, following (Huysman et al., 2010). Nutrients responsive cyclins are highlighted in red. Blue light dependent expression of dsCYC2, mediated by the photoreceptor aureochrome 1a in *Phaeodactylum tricornutum* is highlighted in blue. Light dependent segment of G1 phase is equally shown. Image based on (Bilcke et al., 2021; Huysman et al., 2010, 2013).

It has been proposed that this expansion allows diatoms to respond promptly to environmental conditions and adapt cell division accordingly (Huysman et al., 2014b). Indeed, the progress of the cell cycle is strictly regulated by so-called check points that ensure the correctness of the genetic material before its distribution among the daughter cells as well as the proper accumulation of energy resources essential for the completion of mitosis. These checkpoints can be located in the G1 phase (before the G1-S transition) or during the G2 phase (for the G2-M transition). In diatoms, light exposure plays a major role in controlling the transition from one phase of the cycle to another (Huysman et al., 2013; Vaulot et al., 1986). We can distinguish between light-dependent

and light-independent phases of the cell cycle, which can be studied by inducing a forced arrest of cell division via a prolonged dark treatment, after which the cells will tend to synchronize in the same phase (Gillard et al., 2008; Huysman et al., 2010; Vaulot et al., 1986). In some diatom species, the exposure to prolonged dark induces only part of the cultures to stop cell cycle progression in G1, while the remaining fraction arrests in G2 phase (Brzezinski et al., 1990). The centric diatom *Thalassiosira weissflogii* is an example of species showing similar responses to dark (Ashworth et al., 2013; Vaulot et al., 1986). These results suggest the existence of two light-dependent checkpoints in G1 and G2 respectively. In other species the dark treatment causes an almost complete
synchronization of cultures only in G1 phase, as it happens in *P. tricornutum* (Huysman et al., 2010) or *S. robusta* (Gillard et al., 2008). The G1 light-dependent progression has been dissected in *P. tricornutum* (Huysman et al., 2013), while the G2 light-dependent checkpoint observed in other species remains uncharacterised. Specifically, the diatom specific cyclin dsCYC2 has been demonstrated to be involved in the gating of cell division in response to light exposure in *P. tricornutum*. dsCYC2 ensures the proper transition from G1 to S phase to the extent that downregulation of dsCYC2 causes reduced growth capacity in *P. tricornutum* (Huysman et al., 2013). In *P. tricornutum*, light dependency of dsCYC2 is ensured by the blue light-sensitive photoreceptor aureochrome 1a (Fig.6). As reported, aureochromes act both as photoreceptors via their LOV domain and as DNA binding transcription factors via their bZIP fold. Upon light irradiation, *P. tricornutum* aureochrome 1a dimers bind ds CYC2 promoter along with homodimers of bZIP10 transcription factor and induce its expression (Huysman et al., 2013). A similar blue-light dependent regulation of dsCYC2 expression and G1 progression has been recently described in the benthic pennate diatom *S. robusta*, suggesting a possible conservation of this mechanism (Bilcke et al., 2021b).

Nutrient availability is another factor affecting diatom cell cycle and population dynamics, a principle valid for other marine microorganisms. Various dsCYC genes have been found to be differentially expressed under conditions including phosphate repletion (dsCYC5, dsCYC7, and dsCYC10), and the presence of silica (dsCYC9, Fig.6, Bowler et al., 2008; Huysman et al., 2010, 2013; Sapriel et al., 2009). In oceanic ecosystems, the elements that represent the main limiting factors for planktonic proliferation are nitrogen, phosphorus, iron and, in part, silicon (Falkowski et al., 2008, 1998). The depletion of these nutrients causes

a slowdown if not an arrest of cell division in diatoms. A prime example is given by nitrogen, an element fundamental for the biosynthesis of amino acids as well as pigments and other biomolecules and for overall diatom growth (Valenzuela et al., 2012). Nitrogen limitation and starvation have been shown to stop cell cycle progression at the G1 phase in diatoms (Huysman et al., 2014b; Vaulot et al., 1987).

The last step of cell division is, as already mentioned, cytokinesis. The cytokinesis patterns of diatoms exhibit features intermediate to those seen in other organisms. They possess a unique microtubule (MT)-organizing center (MTOC), also called the MT center, similar to those that can be found in animals and yeasts but absent in plants (De Martino et al., 2009; Pickett-Heaps and Tippit, 1978). Pennate diatoms lack animal-like centrioles, which are however present in microgametes of centric diatoms (De Martino et al., 2009). Also, diatom MTOC differs from its counterpart in brown algae (*Stramenopila*) that have an animal-like centrosome containing two centrioles. This diversity highlights the extreme evolutionary malleability of cell division mechanisms and organization even in phylogenetically closely related organisms (De Martino et al., 2009). During cytokinesis, the division of the mother cell in the two daughters occurs thanks to a longitudinal cleavage furrow that spans across the cell. The new Golgi apparatus, endoplasmic reticulum, mitochondria and plastids all align to this furrow ready to split to the two new cells, implying a major synchronization of the duplication of these organelles with the cell cycle and an important intracellular membrane trafficking (A. Tanaka et al., 2015). Of particular relevance, is the timing and regulation of chloroplast(s) division along the cell cycle. Diatoms display a considerable variability in the number of chloroplasts depending on the species. They can be indeed classified in five different groups: monoplastidic, diplastidic,
tetraplastidic, oligoplastidic, and polyplastidic
species (Mann and Droop, 1996). In diatoms,
with the exception of polyplastidic species, the
development and division of the chloroplast is
tightly coordinated with cell cycle progression
(Gillard et al., 2008; Huysman et al., 2010).
This synchronization is clearly of cardinal
importance to ensure an equitable
redistribution of plastids among daughter cells
and has been extensively studied in pennate
diatoms, in particular in the model species
*Seminavis robusta* (Gillard et al., 2008) and
*Phaeodactylum tricornutum* (Huysman et al.,
2010). When synchronized under prolonged
dark, the cells begin DNA synthesis 4 hours
upon re-illumination and cytokinesis starts
after 9-12 hours when first cells begin to
display the longitudinal division furrow
(Gillard et al., 2008). Cytological observations
and analysis of the transcriptional profile of
these algae also showed that chloroplast
division precedes nuclear and cellular
division, thus ensuring the correct and orderly
succession of division processes (Gillard et al.,
2008; Huysman et al., 2010). Given this fine
synchronization between cell cycle and
chloroplast division, the optical properties of
the plastid can be used as a proxy for the
progress of cell division. One of the methods
that can be used for this purpose is the analysis
of chlorophyll fluorescence per cell by flow-
cytometry (Annunziata et al., 2019;
Hunsperger et al., 2016; Ragni and d'Alcalà,
2007) (described in more detail throughout
this thesis). Such a coordination of plastid
biogenesis and cell division obviously requires
fine-tuning of gene expression and a dedicated
set of proteins. Some of these are conserved
and shared with plants and other algae, being
a relic of the cell division mechanism of the
host prokaryote that gave rise to the plastid.
Among these, the FtsZ protein is the best
known in plants (TerBush et al., 2013) and
several FtsZ copies have been found in
diatoms (Ait-Mohamed et al., 2020; Gillard et
al., 2008). Analysis of FtsZ genes expression
can be used as a tool to follow the temporal

progression of chloroplast division (Gillard et
al., 2008).

In the natural environment diatoms
(like other groups of phytoplankton) can
display a “bloom and bust” life cycle. When
they encounter favorable environmental
conditions of light, temperature, nutrient
availability (often linked to fluvial runoff of
onshore deposits and upwelling of nutrient-
rich deep water in proximity of the coast
(Lassiter et al., 2006)) they undergo a rapid
acceleration of the mitotic rate and a
consequent exponential increase in cell
concentration, to such a degree that these
"flowering" phenomena are easily observed
from space (Anderson et al., 2011; Gross,
2012; Raitsos et al., 2008). These blooms are
followed by a population collapse likely due to
a combination of abiotic and biotic factors,
including predation by grazers, depletion of
nutrient resources and the action of viral
pathogens (Gross, 2012; Smetacek, 2012;
Vincent et al., 2021). These algal bloom events
are not only of utmost importance for the
functioning of marine trophic networks, but
also represent a phenomenon of major
importance for public health issues. In fact,
toxic microalgae blooms are well known and
extensively studied and can represent a
significant danger to humans and marine life,
including those of the diatom genus *Pseudo-
nitzschia* (Walsh, 2008).

Notably, structure and composition of
oceanic microbial communities show seasonal
patterns, also due to the response to seasonal
cyclic phenomena such as the availability of
nutrients and other physical and chemical
parameters of the water (Gilbert et al., 2012;
Lambert et al., 2019; Marquardt et al., 2016)
and, possibly, day length. Photoperiod
perception and photoperiodism have already
been long studied in different terrestrial
organisms (Pittendrigh and Minis, 1964) and
the primary role of endogenous molecular
clocks (see below) in regulating these
phenomena is well known in terrestrial
organisms (Farré and Weise, 2012). For this,
one of the dominant hypotheses is that also the
molecular clocks of microalgae have a leading
role in the control of blooming phenomena in
response to the correct photoperiod (Lambert

et al., 2019). Still, how and to what extent this
is true remain unanswered questions.
# I.4 Biological clocks

## I.4.a Biological clocks: principles and characteristics

For every living organism, the ability to adapt to its environment and its changes is the basis of evolutionary success. However, some of the environmental stimuli to which they have to respond are not unpredictable and follow a precise rhythmicity linked to precise geophysical processes: the day/night cycles due to the rotation of the earth on its axis (with the consequent rhythmic oscillations of temperature), the succession of seasons due to the revolution of our planet around the sun, the tides and the cycles of illumination due to the light reflected from the moon linked to the rhythms of our satellite. Being able to anticipate these changes and prepare metabolism and physiology in advance to best respond to these changes is therefore a huge fitness advantage (Johnson and Golden, 1999; Ouyang et al., 1998; Sanchez et al., 2011). For this reason, a large variety of prokaryotic and eukaryotic organisms have developed molecular timekeeping machinery that allows them to generate endogenous rhythms that oscillate with the same frequency of the environmental rhythms and thus regulate cellular as well as organism-wide activity. Given their timekeeping function, these molecular mechanisms are commonly referred to as biological clocks<fnref n="3" />

We can describe such an oscillator as a system that tends, in a regular manner, to move away from equilibrium before moving back to it (Dunlap, 1999). The rhythmic activity of such a mechanism (and of the processes it regulates downstream) can be assimilated to a sinusoidal function (Fig.7) and is characterized by a series of parameters such as: the amplitude of oscillation, i.e. the difference between the value of the system at

the equilibrium and the value assumed at its maximum deviation from the equilibrium itself; the phase, which for simplicity can be described here as the relative position of the peak of oscillation relative to the reference system (in the case of a biological clock, for example, it can be the moment of maximum activity of a daily cyclic process with respect to the time of the day); and the period, i.e. the time interval between two peaks of oscillation.

Depending on the length of the oscillations (and therefore, the environmental cycles they are synchronized with) biological clocks can be divided in: circadian (“approximately daily”) when they follow solar 24hrs cycles; tidal (24,8hrs) and semitidal (12,4hrs) when they follow the course of tides; lunar (29,5 days) or semilunar (14,8 days) in the case they are synchronized with cycles of moon illumination phases; circannual when they oscillate following the 365 days earth revolution. The discipline that studies biological rhythmicity and the cellular and molecular processes that govern them is also called chronobiology. Although biological clocks are composed of different molecular "gears" depending on the phylum considered, they all share common features. Before entering in the description of the different molecular clocks known today, it is useful to describe which are these common characteristics. A first important feature of biological clocks is their “three-level” structure. They are in fact composed of the proper molecular oscillator called the core clock. At a second level, a series of mechanisms of perception of environmental stimuli (input pathways). Finally the output pathways are the set of biological processes controlled by the core clock and that are

<footnote n="3">³The use of the term biological clocks has become mainstream during the 1960 Cold Spring Harbor</footnote>

<footnote>Symposium of Quantitative Biology, titled “Biological Clocks.” (Cold Spring Harbour, 1960)</footnote>
<figure><img src="image_15.png" /><figcaption>Figure 7. Main characteristics of biological rhythms</figcaption></figure>

Schematic representation of a generic rhythmic biological process. Oscillations can be modelled to a cosine function with a period equivalent to that of the environmental cycles in Zeitgeber time (i.e. in presence of rhythmic external cues). Two headed arrows highlight the main parameters that can be used to describe rhythmic oscillations: amplitude, phase (in this case, the acrophase, i.e. the peak of the oscillation), and period (the time separating two points with the same phase). Following release to free run, the oscillations persist if the process is regulated by a circadian clock, but can show a progressive damping of the oscillation amplitude over time (e.g. Ould et al., 2018). The period in free run, although close to that observed in Zeitgeber Time, can shorten or lengthen (as in the example shown in the figure). In the figure, light has been chosen as an example of rhythmic cue: white boxes represent light periods (days) in the *Zeitgeber* time and in free run; black boxes represent dark periods (nights) under *Zeitgeber* time; gray boxes represent subjective dark periods after release to continuous light. Other environmental cues (e.g. temperature cycles) can exert a similar synchronization of the biological activity.

rhythmic in response to the cyclic activity of the latter (Dunlap, 1999; Wijnen and Young, 2006). Focusing on the core clock, such a mechanism can be achieved by negative feedback loops in which a set of positive elements induces the synthesis of negative elements which, in turn, acts as a brake by blocking the synthesis or activity of the positive ones (Dunlap, 1999). The core clock is taken to mean the minimal set of molecular causes (hence proteins and regulatory DNA sequences) sufficient to operate circadian cycles (Dunlap, 1999).

The input pathways are involved in the second common feature of biological clocks, that is their ability to synchronize with the external environment through the perception of periodic cues detected by specific receptors (principle commonly named “entrainment”). The primary function of entrainment is to ensure that endogenous

biological oscillations constantly adapt to external conditions, ensuring for example that cellular physiology synchronizes to light/dark transitions and vice versa even when these vary over the course of the year with the passing of the seasons (Webb et al., 2019). A classic example of entrainment in humans is given by the readaptation of biological rhythms following sudden geographical displacements (jet lag recovery) (Leloup and Goldbeter, 2013; Roenneberg et al., 2007). The factors capable of synchronizing the endogenous clock are commonly called *Zeitgebers* (from the German "timegivers") a term coined by Jürgen Aschoff, one of the founders of chronobiology (Aschoff, 1954). The most obvious *Zeitgeber* involved in the synchronization of the clock is undoubtedly light, an indicator of the passage of the day-night cycle. However, the temperature cycles associated with the alternation of day and night represent a time giver of primary importance
in the synchronization of biological activity (Eckardt, 2005). Small temperature oscillations (less than 2°C) have been shown to be sufficient to affect the entrainment of a variety of organisms as different as fungi, plants and animals (Rensing and Ruoff, 2002). Still, the molecular actors involved in the sensing of these cues and in the downstream synchronization of the endogenous oscillator have to be clarified. *Zeitgebers* can also be represented by factors other than simple physical stimuli from the environment. For example, metabolic inputs (food in the case of animals or sugars produced by the cyclic activity of photosynthesis in autotrophs) also play a relevant role in synchronizing the rhythmicity of the organism (Carneiro and Araujo, 2012; Haydon et al., 2017, 2013). Significantly, in the case of social animals, the interaction with peers can also promote a synchronization of activity and behavior at the level of the group in a mechanism that can even override the synchronization mediated by physical environmental stimuli (Fuchikawa et al., 2016; Levine et al., 2002). Importantly, the synchronization of biological activity with the environment also allows for the temporal separation of conflicting physiological processes. A classic example of this temporal separation is found, for example, in nitrogen-fixing cyanobacteria. In these organisms the clock regulates the differential expression of the enzyme nitrogenase (inhibited by oxygen) during the night, when oxygenic photosynthesis is not active (Kondo and Ishiura, 2000; Mitsui et al., 1986).

The ability to synchronize the oscillations of the circadian clock with the external cues via the input pathway is also at the basis of the third common characteristic of circadian clocks, namely the aforementioned anticipation of environmental cycles. In other words, the predictability of environmental cycles and the fact that the endogenous timekeeper oscillates on their same frequency allows the clock to prepare the physiological response to a phenomenon before it occurs

(for example the preparation of metabolism to the arrival of sunlight in phototrophic organisms (Dodd et al., 2014, 2005)). Typically, the appearance of a biological response before an environmental stimulus and not in direct response to it is strong evidence of its regulation by an endogenous clock (Harmer et al., 2000). However, it is important to remember that when *Zeitgebers* act on input pathways outside the expected time window and/or if they are too intense (e.g. high light irradiation) they can affect the overt rhythms in a positive or negative way, overriding the activity of the clock and inducing/repressing a specific rhythmic process out of time. In this case, the organism uncouples its activity from the endogenous oscillator and responds directly to the stimulus, in a phenomenon called clock masking (Aschoff, 1999; Redlin, 2001). On one side, masking effects show that clocks do not control physiology in an inflexible manner and that some sort of flexible and adaptive response to the environment is possible. On the other side, small "disturbances" in the rhythmic environment will not have major effects on the timing of an activity, ensuring some degree of rigidity in the system.

The fourth characteristic of endogenous clocks is persistence, probably the most relevant for the study of biological rhythms. Persistence consists in the ability of oscillators to maintain their rhythmic activity even after the organism is moved to a "free run" study condition, i.e. in which the environment is constant and all the rhythmicity of *Zeitgebers* is eliminated (e.g. continuous light or continuous darkness, Fig7). Crucially, free-run oscillations should have a frequency close to that observed before the transition (e.g., approximately 24-hour rhythms in the case of circadian clocks). This phenomenon is due to the nature of the feedback loop that composes the core clock, self-sustained by the activity of its components only, with no need for an external factor to activate the oscillation. The presence of external rhythmic *Zeitgebers* and
their consequent action on the input pathways
are involved only in the synchronization of the
core mechanism with the environment,
without being indispensable to its operation.
Since the dawn of chronobiology, free-run
persistence has been considered an ideal study
condition to dissect the characteristics of
biological clocks (Sweeney and Hastings,
1958). In addition, persistence allows one to
discriminate between processes that are
properly endogenously regulated (which
persist in free run) from those that under
normal conditions oscillate only in direct
response to environmental inputs (which
immediately lose all oscillations in a constant
environment). In the case of the rhythms of 24
hours, all processes that show oscillations
under photoperiodic conditions are defined as
diel (or, interchangeably, daily), but only those
which persist upon release to continuous
environment can be properly defined as
circadian (Leming et al., 2014; Rund et al.,
2011). In particular, the seminal studies that
led to the identification of genes of
endogenous clocks were based on the
characterization of mutants exhibiting altered
persistence of rhythmicity under free-running
conditions (Konopka and Benzer, 1971).

To conclude, a last characteristic
common to biological oscillators is
temperature compensation, or the
maintenance of a constant periodicity of
oscillation even when the environmental
temperature varies. Temperature
compensation is a characteristic of
endogenous clocks that has been known for a
long time and already characterized in the first

half of the 20th century in animal (Brown and
Webb, 1948; Pittendrigh, 1954; Wahl, 1932)
as well as algal models (Bruce and Pittendrigh,
1956; Hastings and Sweeney, 1957).
Biochemical processes dependent on protein
activity speed up as the temperature rises as a
matter of thermodynamic efficiency of
enzymatic reactions (Copeland, 2000). If this
were to happen in timekeeping systems it
would result in their total loss of functionality
outside of an optimal temperature range, and
circadian clocks have evolved to avoid this
problem (Eckardt, 2006).

To date, biological processes with
tidal, lunar and seasonal/circumannual
rhythmicity regulated by endogenous clocks
have been observed (Andreatta and Tessmar-
Raible, 2020; Gwinner, 2012). However, the
best known and best characterized
endogenous clocks are undoubtedly those of
circadian nature, with an oscillation period of
about 24 hours (Wijnen and Young, 2006).
The greater knowledge accumulated on the
oscillators with this frequency is obviously due
to the possibility to more easily follow
biological activity in free run for several cycles
of clock activity in controlled conditions. This
is obviously more prohibitive in the case of
clocks that oscillate on frequencies of several
weeks or even months. For this reason and for
the fact that my thesis work has focused mainly
on the characterization of circadian
rhythmicity, in the next paragraph I will give an
overview only on this type of clocks. A more
extensive description of biological rhythms in
the marine environment is given in the review
presented in Chapter II.a.

I.4.b Biological clocks: diversity and commonalities

The field of chronobiology was
recently brought to the attention of the general
public thanks to the award of the 2017 Nobel
Prize in Physiology and Medicine to Jeffrey C.
Hall, Michael Rosbash and Michael W.
Young for their discoveries of molecular

mechanisms controlling the circadian rhythm
(The Nobel Assembly at Karolinska Institutet,
2017). In the 1980s, the three Nobel laureates
launched the first pioneering studies on the
molecular players of the endogenous clock of
the fruit fly *Drosophila melanogaster*
(Bargiello et al., 1984; Zehring et al., 1984), working on the mutants generated a decade before by Seymour Benzer and Ronald Konopka (Konopka and Benzer, 1971). These studies laid the foundation for the future characterization of circadian clocks in metazoans and other organisms, as well as the methodological framework on which modern molecular chronobiology is based (Peschel and Helfrich-Förster, 2011; Wijnen and Young, 2006).

In eukaryotes, biological oscillators are so-called Transcriptional Translational Feedback Loops (TTFL), in which the positive and negative elements are proteins (notably transcription factors, TFs) capable of regulating their own gene expression and that of target genes (representing the outputs). The structure of the central oscillator of animals has been largely characterized and described thanks to studies in fruit flies and murine models (Fig.8A and Fig.8B). In *D. melanogaster*, the bHLH-PAS Clock (CLK) and Cycle (CYC) proteins interact to form a heterodimer that represents the positive element of TTFL, capable of inducing the expression of a number of target genes that also include the negative elements of the clock, in this case the Timeless (TIM) and Period (PER) nuclear localized proteins (Peschel and Helfrich-Förster, 2011). Light obviously plays a crucial role in fruit fly entrainment, in particular through the photoreceptor cryptochrome that, activated by blue light, induces the degradation of the negative elements TIM and PER, thus releasing the brake on the expression of CLK and CYC (Peschel and Helfrich-Förster, 2011; Stanewsky et al., 1998; Yang et al., 1998). In mammals, only some of the components found in *Drosophila* retain their function in the circadian clock. Also in this case, the positive element is given by heterodimers of bHLH-PAS transcription factors (no more CLK/CYC, but their homologues CLOCK/BMAL1 and NPAS2/BMAL1) (Hirayama et al., 2007; Huang et al., 2012;

Takahashi, 2017a). Negative elements induced by bHLH-PAS proteins again include a homolog of PER, which however in this case associates not with a homolog of TIM, but with mammalian cryptochromes which lose their function as direct light sensors (Takahashi, 2017a; Tamaru et al., 2015; Wijnen and Young, 2006). Light entrainment in mammal clocks is more dependent on visual perception and activation of cAMP-dependent signal transduction pathways induced by neural signaling (Gau et al., 2002; Shearman et al., 1997; Wijnen and Young, 2006). This divergence makes it clear that even organisms that are (relatively) evolutionarily close may exhibit substantial differences in the molecular players involved in the central oscillator, while retaining its basic structure. Lest we forget, the central feedback loop acts in close association with other peripheral TTFLs that refine the mechanisms of expression and repression of positive and negative core elements (loops composed of the VRI and PDP1 proteins in *D. melanogaster* and RORα and REV-ERBα in mammals). A complete overview of the functioning of animal clocks can be found in Peschel and Helfrich-Förster, 2011 (*D. melanogaster*) and Takahashi, 2017b (mammals), but what described above shows how the core clock, although reducible to a TTFL, can actually have a much more complex and baroque architecture.

Fungi are organisms relatively close to animals in phylogenetic terms, being part of the same phylum, *Opisthokonta* and are classically organisms widely used in molecular biology. This is why some of the earliest and most important studies of the characterization of circadian clocks were developed in fungi. In particular, the model organism most applied in chronobiology is the filamentous fungus *Neurospora crassa*, belonging to the *Ascomycota* (Davis, 2000). *N. crassa* has a long history as a model organism in genetics (Beadle and Tatum, 1941) and shows a rhythmic daily production of spores for
<figure><img src="image_16.png" /><figcaption>Figure 8. Circadian TTFL networks in model organisms</figcaption></figure>

Simplified schematic representation of the TTFL networks at the base of circadian regulation in different organisms: insects (Drosophila melanogaster, A), mammals (Mus musculus, B), fungi (Neurospora crassa, C) and plants (Arabidopsis thaliana, D). Circles represent the different regulatory proteins reported in the text. Black arrows represent positive regulation (induction of expression or activation of protein function). Red blunt lines represent negative regulation (repression of induction, induction of protein degradation or repression of protein function). Rainbow arrow highlight light dependent activity of photoreceptors. Images are based on Saini et al 2019 (panels A-C) and Creux 2019 (panel D). Protein names abbreviations are reported in the main text.
asexual reproduction (macroconidia) (Baker et al., 2012). This rhythmicity, in particular, has an endogenous regulation being able to persist even in free run in a temperature-independent manner (Pittendrigh et al., 1959). As in animals, the endogenous timekeeper of *N. crassa* is based on a TTFL, although composed of completely different proteins (Fig.8C). In particular, the positive element of *N.crassa* TTFL are two GATA-family transcription factors, *white-collar* (WC-1 and WC-2), that heterodimerize thanks to the physical interaction of their LOV domains (of the PAS domain family) forming the so called WCC (White-Collar Complex) (Cheng et al., 2002). The WCC induces the expression, among others, of the main negative element of the clock, the *frequency* (FRQ) protein (Crosthwaite et al., 1997). Upon expression, FRQ proteins form homodimers (Cheng, 2001) that in turn interact with FRH (FRQ interacting RNA helicase) forming the FRQ-FRH Complex (FFC) (Cheng et al., 2005). FFC interacts with WCC, favouring the phosphorylation of WC proteins and the consequent destabilization of the WCC and its dissociation from the *frq* promoter (Hong et al., 2008; Schafmeier et al., 2005) hence closing the loop. Concerning the light entrainment of *N.crassa* timekeeper, this occurs primarily through the WC-1 protein. In fact, it does not only play the role of a transcription factor, but is also a photoreceptor activated by blue light. This sensitivity to light is related to its LOV domain, which is capable of binding the chromophore flavin adenine dinucleotide (Chen et al., 2010). In particular, the stability of WCC (and the consequent functioning of the positive arm of TTFL) is implemented by irradiation in the blue light of WC-1 (Crosthwaite et al., 1997). Overall, this dual function of WC-1 demonstrates how the division of the clock into different components (input, core, and output) is not rigid and that the same molecular actor can intervene on multiple levels of clock regulation (in this case photo perception and core TTFL).

Despite the interest and scientific efforts expended in characterizing the animal and fungal clock, the history of chronobiology dates back to far before the introduction of the fruit fly and *N.crassa* as model organisms. From the beginning, chronobiology has been a discipline inextricably linked to the study of photosynthetic organisms. The first scientific reports of rhythmic biological phenomena date back to the 18th century, with the famous observations made by Jean-Jacques d'Ortus de Mairan, who noted that cyclic daily leaf movements in heliotrope plants continue after transferring the plants to constant darkness (D'Ortus de Mairan, 1729). Similarly, the first studies of genetics applied to chronobiology were carried out on plant models, with the work of Erwin Bünning, first to demonstrate that period length is heritable in bean plants (Bünning, 1936, 1935; Kuhlman et al., 2018).

Similar to the animal clock, the clock of higher plants is controlled by a central mechanism based on feedback loops using transcription factors (Fig.8D). The plant central circadian oscillator has a high level of complexity, being composed of multiple loops that follow one another and intertwine over the 24 hours and depend on the expression of specific proteins at different times of the day. At the beginning of light phase MYB-like transcription factors CIRCADIAN CLOCK ASSOCIATED1 (CCA1) and LATE ELOGATED HYPOCOTYL (LHY) are expressed and repress the afternoon PSEUDO-RESPONSE REGULATOR (PRR) genes, including TIMING OF CAB EXPRESSION1 (PRR1/TOC1), PRR5, PRR7, and PRR9 (Alabadí et al., 2001; Farré et al., 2005; Kamioka et al., 2016). PRR TFs in turn repress the expression of the morning genes creating a double repressor loop (Creux and Harmer, 2019). The correctly timed expression of PRR proteins in the afternoon (and the consequent override of the inhibitory action of CCA1 and LHY) is guaranteed by the mid-day expression of other MYB-like proteins, REVEILLE (RVE) RVE4, RVE6
and RVE8 that interact with NIGHT LIGHT INDUCIBLE AND CLOCK REGULATED (LNK1 and LNK2) to induce the expression of PRR TFs (Farinas and Mas, 2011; Rawat et al., 2011). The 24-hour regulatory cycle is closed by the late-day expression of the evening regulatory transcriptional complex proteins, EARLY FLOWERING ELF3 and ELF4, and LUX ARRHYTHMO (LUX). These proteins repress genes expressed in the morning and afternoon to complete the feedback loop in the network (Huang and Nusinow, 2016).

As in *Drosophila* circadian systems, blue light and its perception through photoreceptors of the cryptochrome family play a central function in entrainment processes also in plants. This is evidenced by alterations in the persistence of free-running oscillations in CRY1 cryptochrome mutants (Gould et al., 2013; Somers et al., 1998). In addition to cryptochromes, plants have also co-opted other photoreceptors in clock regulation, as in the case of the red sensing phytochrome (Somers et al., 1998) or the blue light receptor of the LOV ZEITLUPE family (ZTP) that when activated contributes to the stabilization of TOC1 via the interaction with GIGANTEA (GI) (Kim et al., 2007; Más et al., 2003)

It is important to note that many terrestrial plants are able to perceive the length of the day (photoperiod) and exploit it as a seasonal indicator. Through the perception of day length, plants are able to correctly time processes such as flowering, dormancy, and bud break so as to induce them in the appropriate moment of the year (a phenomenon called photoperiodism). Links between the circadian clock and photoperiodism have been widely studied, especially in correlation to flowering in response to short and long days (Creux and Harmer, 2019). FLOWERING LOCUS T (FT) family proteins, which are highly conserved in angiosperms, play a central role

in regulating flowering (Andrés and Coupland, 2012). FT expression is induced by the transcription factor CONSTANS (CO), which in turn is induced by a protein complex formed by GI and FKF1 (FLAVIN-BINDING KELCH REPEAT, F-box 1, a protein related to ZTL). The GI-FKF1 complex is stabilized by blue light and thus represents a central link in the mechanism that regulates the interaction between light input, central clock and flowering (Imaizumi et al., 2005; Song et al., 2015). Also, the obliteration of central genes of the plant circadian clock can lead to relevant alterations in photoperiodic flowering, as in the case of the evening complex component ELF3 (Zagotta et al., 1996) or PRR genes (Nakamichi et al., 2012). This close interdependence between day length perception and the circadian clock in plants suggests that similar links may play out in other organisms in which these mechanisms are less well characterized. It is also relevant that, in contrast to plants, animal photoperiodic phenomena (which include, for example, hibernation and reproductive behaviors) do not appear to be altered by central clock mutations, suggesting that photoperiodism and circadian rhythmicity may undergo independent evolutionary pathways (Bradshaw and Holzapfel, 2007).

To conclude this comparative analysis of known TTFIs, it is important to consider what is known to date about circadian clocks in unicellular algae. Like plants, algae have been the subject of study long before the development of modern molecular chronobiology (Noordally and Millar, 2015). Historically, some of the earliest algal models to be used in chronobiology are evolutionarily very distant from land plants and are indeed secondary endosymbionts. The marine dinoflagellate *Lingulodinium polyedrum* (formerly *Gonyaulax polyedra*, belonging to the *Alveolata* family) has been extensively studied since the 1950s because of its highly rhythmic bioluminescence able to persist for weeks in free run condition (Hastings, 2007).
This has made it a perfect model for classical
chronobiology and many concepts now
established in the field, such as temperature
compensation of circadian clocks, have been
developed on this organism (Hastings and
Sweeney, 1957). Another secondary
endosymbiont alga that has been widely
studied in chronobiology is *Euglena gracilis*, a
freshwater flagellated protist belonging to the
phylum *Discoba*. In *E. gracilis* the
endogenous circadian control of cell division,
as well as the involvement of cAMP in the
regulation of cellular rhythmicity, have been
well characterized (Bolige et al., 2005; Carré
and Edmunds, 1993). However, due to the
poor advancement of genomic sequencing of
these algae and the few gene manipulation
techniques available, knowledge about the
molecular players involved in TTFLs of
secondary endosymbionts is basically null
(Noordally and Millar, 2015). Also for this
reason, the only microalgae in which the
molecular characterization of the circadian
clock is advanced are those belonging to the
green lineage, evolutionary closer to the
already characterized land plants and for
which more molecular and genomic resources
are available.

One of the most prominent among
them is *Chlamydomonas reinhardtii*. The
endogenous control of its circadian rhythms of
phototaxis and cell division are in fact already
known since the early 70's (Bruce, 1970). The
first studies on mutants with alterations of this
rhythmicity in free run date from the same
period (Bruce, 1974, 1972). Subsequent
studies have shown that chloroplast gene
expression is also under the control of the
same circadian clock (Hwang et al., 1996).
The creation of transgenic lines expressing a
luciferase under the control of the elongation
factor promoter Tu (*tufA*), a strongly rhythmic
plastid gene, later allowed the high throughput
identification of the genes involved in its
endogenous timekeeper. This analysis was
also made possible by the sequencing of *C.
reinhardtii* genome (Merchant et al., 2007).

The screening of insertional mutants led to the
identification of genes, named *rhythm of
chloroplast* (ROC), whose mutations induce
major deregulation of persistent circadian
rhythms of the chloroplast luminescent *tufA*
reporter (Matsuo et al., 2008). Among others,
the four best known are the nuclear genes
ROC15, ROC40, ROC66 and ROC75.
Although they cannot be considered
homologs in the strict sense, these proteins
share some functional domains of several core
clock components of *A. thaliana* such as
CCA1 or LHY (for the Myb TF ROC40),
LUX (in the case of ROC15 and ROC75) and
Constans/TOC1 (as in the case of the CCT-
containing ROC66) (Matsuo et al., 2008;
Schulze et al., 2010). Still, this similarity does
not go beyond the single domain level, plus
other of the ROC proteins show no similarity
for plant clock regulators (Matsuo et al., 2008).
This suggests a clear differentiation of
timekeeping mechanisms even between
relatively close organisms is possible. This
differentiation is made even clearer by the role
played by the protein complex CHLAMY1.
CHLAMY1 is a RNA-binding protein able to
bind in a circadian fashion an UG repeated
sequence located in the 3'-UTR of target
mRNA (Mittag, 1996; Mittag et al., 1994;
Waltenberger et al., 2001). This complex,
composed of two subunits C1 and C3 (Zhao
et al., 2004), has been shown to repress the
activity of the target genes, most probably
acting as a repressor of translation (Iliev et al.,
2006; Schulze et al., 2010). Moreover, the
deregulation of the expression of its two
subunits has major effects on cellular
rhythmicity (Iliev et al., 2006). Despite its
crucial function in the circadian rhythmicity of
*C. reinhardtii*, this protein complex has not
been found in plant genomes, further stressing
how timekeeping mechanisms can diverge
very rapidly and co-opt different players in
relatively short evolutionary timescales
(Schulze et al., 2010). However, one of the
factors that has remained conserved with
respect to plants is the role of blue light in
regulating endogenous rhythmicity. Indeed, *C*.
*reinhardtii* possesses 4 cryptochromes and the mutation of one of them (pCRY) has major effects on free-run phototaxis rhythmicity, indicating a role in input pathways (Müller et al., 2017).

A second green alga of major relevance in chronobiology is *Ostreococcus tauri*, the smallest free-living (non-symbiont) eukaryote described to date (Courties et al., 1994). *O. tauri* is part of the taxon *Prasinophyceae*, one of the first classes to differentiate into green algae (Leliaert et al., 2012), a fact that makes it a relevant study subject for characterizing elements of the circadian timekeeper common to other *Viridiplantae*. This unicellular alga exhibits strong rhythmicity of gene expression when exposed to cycles of light and dark (Monnier et al., 2010; Moulager et al., 2007). Equally, *O. tauri* cell division is rhythmic under photoperiod and maintains a rhythmicity of approximately 24 hours following transition into continuous light, demonstrating that this process is under the control of an endogenous clock (Moulager et al., 2007). *O. tauri* has an extremely compact and low redundant genome, with only 8000 genes (Derelle et al., 2006). This reduced genetic redundancy is also reflected in the structure of its endogenous clock. In fact, *O. tauri* has only one homologous gene of the TOC1 protein found in higher plants and one TF of the Myb family homologous of CCA1. The variety of PRR genes or a second Myb gene comparable to LHY that can be found in plants are not present in the genome of this alga (Corellou et al., 2009). In *O. tauri*, TOC1 sees a peak in expression during the end of the light period, followed by a decrease in expression level during the dark hours, when it is CCA1 that is transcribed instead (Corellou et al., 2009). The expression patterns of these two genes adapt dynamically to different photoperiods (Thommen et al., 2010) and the rhythmicity of expression persists following release into continuous light (Corellou et al., 2009). The misexpression by overexpression or

downregulation of each of these two genes induces a complete disruption of this persistence, demonstrating their cardinal role in the core oscillator (Corellou et al., 2009). The small number of components of the *O. tauri* oscillator has made this alga an excellent subject of study for the development of mathematical models to describe the clock regulation circuit (Morant et al., 2010; Ocone et al., 2013; Thommen et al., 2010; Troein et al., 2011). Some of these models predict that TOC1 acts as the positive arm of the feedback loop, while CCA1 (induced by TOC1), would represent the negative arm of the circuit and would be responsible for the repression of TOC1 (Morant et al., 2010; Thommen et al., 2010; Troein et al., 2011). However, it has also been proposed that the clock circuit of *O. tauri* is actually based on a triple repressor circuit (repressilator), in which CCA1 represses TOC1, responsible for the inhibition of a third component, not yet identified, which in turn represses CCA1 (Ocone et al., 2013). In this three-loop model, TOC1 would assume a repressor role similar to that observed in the circadian clock of higher plants. Concerning light perception, *O. tauri* possesses five proteins of the cryptochrome/photolyase family, none of which turn out to be plant-like (Heijde et al., 2010). Downregulation of one of these proteins, CPF1, causes a perturbation of the persistence of CCA1 expression rhythmicity in continuous light, suggesting its involvement in the circadian clock (Heijde et al., 2010). Furthermore, the CPF1 from *O. tauri* is capable of complementing the function of mammalian cryptochrome in repressing the activity on CLOCK and BMAL1 proteins in a cryptochrome knockout simian cell model (Heijde et al., 2010). In addition, *O. tauri* also possesses a Histidine Kinase-LOV and a Histidine Kinase-Rhodopsin photoreceptors. Downregulation as well as overexpression of LOV-HK protein induces an important deregulation of *O. tauri* free-run rhythmicity (Djouani-Tahri et al., 2011), indicating its involvement in clock inputs, whereas a
function of HK-Rhodopsin has not yet been
fully characterized.

I.4.c Translation Independent Oscillators: above and beyond TTFLs

The mechanisms described in the
previous paragraph only represent the heart of
the core clocks. Although TTFLs are the best
characterized timekeeping mechanism, other
oscillatory systems that are not directly
dependent on gene expression have been
described. TTFLs appear to be
interconnected to a more primitive metabolic
oscillator, able to work even when
transcription and translation are inhibited or
inactive (Wong and O'Neill, 2018). Such
oscillators are also referred to as TIO
(Transcription Independent Oscillators) or
PTO (Post-Translational Oscillators).

Post-translational oscillators are already present in cyanobacteria, the prokaryotes in which a circadian timekeeper has been first identified and characterized in detail (Iwasaki and Kondo, 2004). This mechanism is based on the activity of the Kai periodosome formed by the proteins KaiA, KaiB and KaiC. KaiC is capable of autophosphorylation, with KaiA favoring the process and KaiB repressing it. The Kai periodosome regulates gene expression by rhythmically modifying the supercoiling of the bacterial chromosome (Iwasaki and Kondo, 2004). This circadian clock follows autonomous cycles of phosphorylation and dephosphorylation even without gene expression. In addition, only ATP is required to assemble a working periodosome clock *in vitro* (Nakajima et al., 2005).

In eukaryotes, the ability to maintain
a circadian rhythm in the absence of
transcription has been known since the '60s,
when it was evidenced that the giant unicellular
green alga *Acetabularia* maintains rhythms in
photosynthetic activity even after enucleation
(Sweeney and Haxo, 1961). In addition, it has

been recently demonstrated the wide
preservation of a metabolic rhythmicity in the
form of cycles of cellular redox potential and
ROS content, which can be tracked by
following the oxidative state of the enzyme
peroxiredoxin (PRX, involved in the
neutralization of H₂O₂). Circadian cycles of the
redox state of PRX were originally observed in
erythrocytes (cells naturally lacking the
nucleus and without transcriptional activity)
(O'Neill and Reddy, 2011) and in the
unicellular green alga *O. tauri* specifically
treated to inhibit transcription (O'Neill et al.,
2011). Subsequently, transcription-
independent rhythms of PRX redox state were
similarly observed in organisms spanning the
entire tree of life, including prokaryotes and
archaea, plus a variety of eukaryotes belonging
to different phyla (Edgar et al., 2012). On the
one hand, redox potential and ROS content
are modulated by the circadian clock through
its regulation of cellular metabolism (Putker
and O'Neill, 2016). On the other hand, these
factors act on timekeeping mechanisms
(TTFL and TIO) through modulation of
protein activity (including that of TFs) and a
perturbation of these parameters can have
repercussions on cellular rhythmicity (Putker
and O'Neill, 2016). Therefore, it cannot be
excluded that also the cellular redox potential
has played an ancestral role in biological
timekeeping. However, to date the functioning
of PTO oscillators and their interaction with
cellular redox potential and TTFLs remain
largely unresolved. A simplified, schematic
representation of the structure of the
endogenous timekeeper including both TTFL
and TIO is reported in Fig.9.

To conclude, post-translational
regulators of the circadian clock, such as
Casein Kinases (Lin et al., 2002; Poliner et al.,
2019; Tang et al., 2009; Uehara et al., 2019; Yang et al., 2003), protein phosphatases, protein methyltransferases and ubiquitination systems (Wong and O'Neill, 2018) are conserved in evolutionarily distant eukaryotic organisms and their inhibition or mutations causes an alteration of cellular rhythmicity (Wong and O'Neill, 2018). This led to the hypothesis that an ancestral timekeeper originated as a post-translational oscillator based on the mechanisms of phosphorylation,

stabilization, and degradation of clock proteins and of their molecular targets (Wong and O'Neill, 2018). The diversity of transcriptional oscillators found in eukaryotes can be explained by a later evolutionary divergence of the gene regulatory mechanisms controlled by the TIO, yet maintaining a common mechanism (the negative feedback loop) via the co-option of different actors depending on the phylum (Wong and O'Neill, 2018).
<figure><img src="image_17.png" /><figcaption>Figure 9. Architecture of circadian clock systems</figcaption></figure>

Schematic and simplified architecture of a circadian clocks system. Cellular rhythmicity is synchronized with the environment by the input pathways, able to perceive external stimuli. Light (represented as a rainbow arrow) plays a major role in this entertainment process. In eukaryotes, the core oscillator is composed by a transcriptional translational feedback loop (TTFL). Positive (+) elements induce the expression of negative (-) elements that in turns repress the activity of the formers, closing the loop. The core clock regulates the timing of a variety of downstream processes, the outputs, by controlling the expression of clock controlled genes. These include cell cycle (in particular in unicellular organisms), metabolic activity and other functions such as behaviour. Light inputs can also act directly on the outputs, independently of the central oscillator, potentially causing masking effects (see main text). The persistence of circadian rhythms in absence of transcriptional/translational activity, demonstrate the possible existence of an additional TIO (Transcription Independent Oscillator) acting mainly on the post translational modification of proteins. The structure and components of the TIO remain largely unknown. Two-way arrows underlie the reciprocal influence between different components of this system (e.g. the central TTFL regulating the rhythmicity of the metabolism, whose products, like sugars, act in turn as entrainers of the timekeeper). Figure based on Wong and O'Neill, 2018.
### I.4.d Circadian rhythmicity in *Stramenopila*: an overview

Since my thesis work has largely focused on the characterization of the circadian clock of diatoms, it is useful to introduce here a quick overview of what is already known about these algae and, more generally, about the phylum *Stramenopila* to which they belong.

As in the other taxa described above, the characterization of biological rhythmicity in *Stramenopila* dates back well before the era of molecular biology. Notably, several species of diatoms exposed to light-dark cycles show a synchronization of cell division with light and dark inputs, both under controlled laboratory conditions (Chisholm and Brand, 1981; Gaidarenko et al., 2019; Nelson and Brand, 1979; Ostgaard and Jensen, 1982, 1982; Paasche, 1968; Ragni and d'Alcalà, 2007; Williamson, 1980) and in the case of samples from natural environment (e.g. (Chang and Carpenter, 1991; Owens et al., 1980; Smayda, 1975; Williamson, 1980). In detail, cytokinesis appears to be limited to specific times of the day, which in the case of diatoms normally corresponds to the end of the light period (e.g. Hunsperger et al., 2016; Nelson and Brand, 1979; Ragni and d'Alcalà, 2007). Such a phasing of cell division emphasizes the interdependence between cell cycle advancement and light irradiation described in Paragraph I.3.d. In some diatoms, cell division oscillations observed under light and dark cycles show persistence after release in continuous light (Ostgaard and Jensen, 1982), indicating a clock influence in regulating the timing of cell division. Still, in literature, the persistence of rhythmic divisions was not unequivocally found in all studies and all species, including models like *T. pseudonana* or *P. tricornutum* (Chisholm and Brand, 1981; Ragni and d'Alcalà, 2007). It is important to note that the synchronization of cell division with external light and dark cycles may result from a metabolic response of the cells, which need to accumulate sufficient

energy to sustain cell division. Consequently, the regulation of the rhythmicity of cell division can be either under the direct control of the clock, either an indirect consequence of the clock's control of metabolism or a simple direct response to environmental rhythmic stimuli.

To date, studies on this class of organisms in molecular chronobiology are still limited and have focused mainly (but not only) on some models of diatoms (*P. tricornutum* mainly) and of *Eustigmatophyceae*, a group of algae that includes some species of high biotechnological interest such as the unicellular algae *Nannochloropsis spp.* Both diatoms and *Eustigmatophyceae* algae show rhythmic gene expression under light dark photoperiod (Chauton et al., 2013; Poliner et al., 2015; Smith et al., 2016) and persistent oscillations in transcription in free run have also been observed (Oeltjen et al., 2004; Poliner et al., 2019). This regulation of expression may, potentially, underlie the multiple circadian processes observed in *Stramenopila*. These include: rhythms in cell division, observed in *Nannochloropsis* (Poliner et al., 2019), but also in the diatom *Skeletonema costatum* (Ostgaard and Jensen, 1982) and multicellular brown algae (Makarov et al., 1995); diel oscillations in cellular fluorescence associated with pigment content (Ostgaard and Jensen, 1982; Owens et al., 1980; Ragni and d'Alcalà, 2007); photosynthetic rate and associated metabolic processes like carbon fixation (Brand, 1982; Nütsch et al., 1984; Palmer et al., 1964; Schmid and Dring, 1992); migration of benthic diatoms in sediments (Fischer et al., 1977; Palmer and Round, 1967).

All of the above suggest the possible existence of an endogenous oscillator responsible for self-sustained cycles of gene expression and circadian rhythmicity in these organisms. However, none of the
*Stramenopila* species sequenced (Armbrust, 2004; Bowler et al., 2008; Vieler et al., 2012) shows conservation of any known central component of characterized eukaryotic or prokaryotic clocks. *Stramenopila* possess SHQKYF- and MYB-related family transcription factors, a family to which belong some components of the plant clock (LHY and RVEs), as well as bHLH-PAS proteins with a structure partially similar to that of CLK, CYC, BMAL1 and NPAS2 of the animal central oscillator (Rayko et al., 2010; Thiriet-Rupert et al., 2016). Moreover, the divergence of the amino acid sequence is such that a conservation of function cannot be easily assumed. Moreover, the possibility that the TTFL of *Stramenopila* uses components that are completely different from those of *Archaeplastida* or animals cannot be excluded given the complex evolutionary history of these organisms and the high divergence of transcriptional oscillators in eukaryotes.

On the other hand, *Stramenopila* have conserved some proteins involved in post-translational regulation and input pathways of the timekeeping mechanism. For example, the inhibition of Casein Kinases alters the persistence of transcriptional rhythms in *Nannochloropsis spp.* (Poliner et al., 2019) in a way similar to what is seen in other phyla (Lin et al., 2002; Uehara et al., 2019). In addition, CPF1 cryptochromes in *P. tricornutum* and *Nannochloropsis gaditana*

show rhythmic expression under photoperiodic light/dark cycles (Coesel et al., 2009; Poliner et al., 2015; Smith et al., 2016). Moreover, as already mentioned in Paragraph I.3.c, the heterologous expression of *P. tricornutum* CPF1 in simian cells is able to complement the function of mutated endogenous CRYs in repressing CLOCK/BMAL1 activity, suggesting that this protein might perform a similar activity on diatom bHLH-PAS proteins. Still, this interaction has not been investigated yet. It is crucial to consider that cryptochrome exert a function in clock synchronization of evolutionarily distant organisms throughout the diversity of eTOL branches (Gould et al., 2013; Peschel and Helfrich-Förster, 2011; Somers et al., 1998; Stanewsky et al., 1998; Takahashi, 2017a). In addition, all the photoreceptors described in *Stramenopila* (Paragraph I.3.c) may play a role in input pathways but functional studies are missing on this front. Moreover, aureochromes do not simply act as photoreceptors but also as transcription factors (Kroth et al., 2017; Mann et al., 2020), implying a possible direct involvement in the TTFL as both light sensors and transcriptional regulators. None of this hypothesis has been tested yet. Globally, at the time of the beginning of my thesis the characterization of TTFL of diatoms and more generally of *Stramenopila* was still a largely unexplored field.
I.5 bHLH-PAS transcription factors
in diatoms

Overall, approximately 2% of the diatom proteome is predicted to encode for transcription factors, TFs (Rayko et al., 2010). This value is comparable to what can be observed in other evolutionarily distant unicellular algae (e.g. 2,1% in *C. reinhardtii* as for (Rayko et al., 2010). For comparison, in multicellular organisms where the level of complexity is increased due to cellular differentiation in different tissues, this value can be considerably higher (in *A. thaliana* 5,9% of the proteome is dedicated to TFs (Rayko et al., 2010; Riechmann et al., 2000)). In the case of *P. tricornutum*, the genes predicted to encode transcription factors total about 200, with the number varying slightly according to predictions (Rayko et al., 2010; Thiriet-Rupert et al., 2016). Among these, a relevant number is given by the Heat Shock Factors, the most numerous family followed by the Myb, bZIP and Zn finger factors that have undergone considerable expansion in diatoms (Rayko et al., 2010; Thiriet-Rupert et al., 2016).

Among the multitude of transcription factor families that can be found in diatoms, the basic-Helix-Loop-Helix (bHLH) family is of particular interest. The common element of these proteins, as their name indicates, is the presence of a highly conserved domain of 50-60 amino acids composed of a strand of 10-15 basic residues (basic region) followed by about forty amino acids that are structured in two amphipathic α-helices separated by an amorphous loop of variable length (the Helix-Loop-Helix region, (Jones, 2004; Murre, 2019; Pires and Dolan, 2010a). These proteins act as homodimers or heterodimers, with the two alpha helices representing the main dimerization interface. The DNA binding function, crucial for any transcription factor, is instead performed mainly by the

basic stretch located upstream of the HLH motif (Jones, 2004). The basic region is specifically tailored to interact with five- to seven-base DNA sequences. In general, the recognized sequences can be of the type CANN(N)TG (E boxes, where N corresponds to any nucleotide) (Carroll et al., 2018), CACNNG (N-boxes) (Paroush, 1994) or ACGTG/GCGTG (typically recognized by bHLH-PAS animal proteins which I will discuss below).

The relatively small bHLH domain is located in the amino terminal region of the protein and can associate with a variety of other downstream domains that determine the functionality of the protein and the specificity of dimerization (Ledent and Vervoort, 2001; Murre, 2019). These additional domains include leucine zipper motifs, Collier/Olf-1/EBF (COE) domains and PAS domains (Ledent and Vervoort, 2001). Although the α Helices of the bHLH domain represent the primary dimerisation interface at the level of the N terminus, the PAS folds act as additional surface of interaction and define the partner of choice (Kewley et al., 2004; Taylor and Zhulin, 1999). In bHLH-PAS proteins, the choice of the dimerization partner is in particular defined by the presence of the PAS domain, so that these proteins can homo/heterodimerize only with another member of the same family (Kewley et al., 2004; Michaud et al., 2000; Pongratz et al., 1998).

The first characterisation of these transcription factors dates back to 1989, when they were first identified in murine cells (Murre et al., 1989b, 1989a). Since, bHLH proteins were identified in the major eukaryotic phyla, including *Metazoa*, *Archaeplastida*, and fungi (Ledent and Vervoort, 2001; Murre, 2019; Pires and
Dolan, 2010b). Their functions are as diverse as the organisms where they can be found. Both bHLH TFs acting as transcriptional activators (e.g. Jang et al., 2017) and as transcriptional repressors (e.g. Kageyama et al., 2000) have been characterized. In animals they are involved in immune response, neurogenesis, myogenesis, and heart development (Jones, 2004; Massari and Murre, 2000; Murre, 2019, p. 201). In yeast they play a central role in glycolysis and phosphate uptake (Robinson, 2000). Finally, plant bHLH transcription factors have been shown to modulate secondary metabolism pathways, epidermal development, and responses to environmental light stimuli, such as in the case of processes like shade avoidance, photomorphogenesis, other processes regulated by photoreceptors and the regulation of the circadian clock (Castillon et al., 2007; Farré and Weise, 2012; Pires and Dolan, 2010a; Ramsay and Glover, 2005). In plants in particular, bHLH factors have undergone an important phenomenon of duplication and increase in gene copies, becoming the second most abundant and diverse families of transcription factors, while in unicellular *Archaeplastida* the number of copies of bHLH factors is far smaller (Pires and Dolan, 2010a, 2010b). Given the role played by plant bHLH factors in development and cell differentiation, it has been proposed that the amplification of this gene family occurred concomitantly with the transition from unicellularity to multicellularity, playing a crucial role in this evolutionary process (Pires and Dolan, 2010b).

*P. tricornutum*, in addition to TFs only containing the bHLH fold, also possess three factors associating PAS folds to the bHLH domain, proteins on which my thesis work focused. Before describing the bHLH-PAS proteins, it is essential to introduce the characteristics and functions of PAS domains. The PAS fold is an ubiquitously distributed functional domain that can be found in all eukaryotes as well as in archaea and

prokaryotes and takes its name from the first three proteins in which it was identified, the *Drosophila melanogaster Period* (PER) and *Single-minded* (SIM) and the mammalian *Arvl hydrocarbon Receptor Nuclear Transporter* (ARNT) (Henry and Crosson, 2011; Möglich et al., 2009; Taylor and Zhulin, 1999). It consists of 100-130 amino acids generating a five or six stranded β-sheet divided in two halves connected by an α-helix and flanked by a variable number of additional α-helices (Kewley et al., 2004; Möglich et al., 2009). PAS domains are not only found associated with bHLH DNA binding domains, but also with other functional domains such as histidine kinase, nucleotide cyclase, and response regulator domains (Henry and Crosson, 2011). All domains, therefore, are involved in signal transduction within the cell.

The PAS fold is a highly multifunctional structure, since it is in a plethora of molecular processes. The ability to interact with a chemically diverse range of metabolites is a hallmark of the PAS domain family. The chemicals bound by this domain are used either as a direct cue to initiate a cellular signaling response or represent functional cofactor moieties involved in the interaction with secondary physical or chemical signals such as gas molecules, redox potential, or photons (Henry and Crosson, 2011; Möglich et al., 2009). Among the ligands recognized by different PAS domains we count dioxin (as in ARNT proteins) (Procopio et al., 2002), metal ions (Ernst et al., 2001; Fass and Groisman, 2009), heme groups (Yoshioka et al., 2005) and various metabolites (such as malate, succinate or citrate (Henry and Crosson, 2011)).

It is important to note that PAS domains also play a central role in the activity of some photoreceptors. In fact, a specific subfamily of these protein folds (named Light-Oxygen-Voltage photoreceptor domains or LOV) is capable of binding a flavin
mononucleotide (FMN) cofactor that forms a cysteinyl-flavin C4(a) covalent adduct when activated by blue-light (Crosson et al., 2003; Purcell et al., 2007). The photoreceptive function of LOV domains was first identified in plant phototropins (Christie et al., 1999, 1998), but other LOV photoreceptors for blue light include the already mentioned LOV-HK proteins (found in prokaryotes as well as green algae (Djouani-Tahri et al., 2011)) and aureochromes, which as seen are present only in *Stramenopila* (Kroth et al., 2017).

bHLH-PAS proteins have been extensively studied in metazoans, where they are involved, for instance, in response to hypoxia and dioxin (like Hypoxia Inducible Factors and ARNTs (Jiang et al., 2001; Sonnenfeld et al., 1997; Takahata et al., 1998)), in the development of nervous system (like SIM) (Nambu et al., 1996) or in the control of circadian clock (as for CLOCK, CYCLE, NPAS2 and BMAL1, see Paragraph I.4.b for a detailed description, (Huang et al., 2012).

Even though bHLH-PAS TFs were originally thought to be uniquely present in metazoans (Kewley et al., 2004), these

proteins are not restricted to the animal kingdom, but have been recently found also in *Stramenopila* like diatoms (Rayko et al., 2010). Gene prediction analysis revealed the presence of three bHLH-PAS proteins in the model species *P. tricornutum* (bHLH1a, bHLH1b and bHLH2) out of a total of seven bHLH transcription factors (Rayko et al., 2010). A similar expansion of the bHLH family of proteins was also found in other diatoms such as *T. pseudonana* (5 paralogous bHLH genes of which 4 also contain a PAS domain, (Rayko et al., 2010)) or in *S. robusta* (13 bHLH genes of which 10 with a PAS domain (Bilcke et al., 2021a)). This is part of a more general expansion of transcription factor families observed in diatoms (Thiriet-Rupert et al., 2016). It is noteworthy that no bHLH-PAS protein has ever been described in any member of the *Viridiplantae* group. However, whereas the large majority of animal bHLH-PAS proteins contain two PAS folds, all known *Stramenopile* bHLH-PAS proteins contain only one PAS domain. Their patchy distribution among eukaryotic supergroups suggests that these domains might have fused more than once during evolution (Rayko et al., 2010).
Aims of the thesis

As for all autotrophic organisms, the life and ecological success of diatoms depend inextricably on their ability to respond to the light environment and its dynamic changes. Adaptation to such stimuli must necessarily pass through a fine-tuning of gene expression, in a regulatory mechanism in which transcription factors clearly play a key role. Despite the ecological relevance of diatoms and their importance in marine ecosystems, however, these regulatory mechanisms remain largely unexplored to date. For this reason, I have devoted my research work to integrate genetic resources with physiological and biophysical approaches to characterize the unknown regulators controlling two light-dependent processes of crucial importance to diatoms: rhythmic cellular processes and photosynthesis. Specifically, I based all my work on the marine diatom model species *Phaeodactylum tricornutum*.

On the one hand, I have characterized the ability of this organism to synchronize their physiology with the daily rhythmic cycles of light and dark. As described in Chapter I, paragraph I.4.d and in Chapter II.a, a large number of biological processes in these algae are modulated according to these cycles, such as cell division, pigment synthesis or photosynthetic metabolism, and more. However, at the time when I began my thesis the knowledge about diatom timekeeper was still in its infancy. The identification of regulators involved in this system of timekeeping were therefore essential to better understand the nature of these processes. As presented in Chapter II.b, early results from this line of work led to the identification of the bHLH-PAS protein bHLH1a (later renamed RITMO1) as one of the most rhythmic *P. tricornutum* transcription factors (TF). Alteration of its expression by overexpression leads to a perturbation of the timing and persistence of cellular fluorescence and gene expression. Based on these results, we proposed RITMO1 as the first regulator of diel biological rhythms identified in diatoms. However, the role of this protein in a putative circadian clock system remained to be defined. Using these initial findings as an entry point, I subsequently pursued a more in-depth dissection of the rhythmic processes of *P. tricornutum* and their control by the endogenous clock.

As presented in Chapter II.c, I decided to detail how this diatom adapts its rhythmicity to different photoperiods and light intensities to understand the flexibility to respond to a range of growth conditions reflecting the variety of environmental stimuli. I also generated lines in which the RITMO1 gene was obliterated by CRISPR/Cas9 targeted mutagenesis to determine how deletion of this protein disrupts cellular rhythms. Through these transgenic lines I was able to demonstrate that the circadian clock of *P. tricornutum* regulates, through RITMO1, a plethora of processes that include cell fluorescence and cell division, photosynthetic activity and, more generally, gene expression. These studies established RITMO1 as a component of the diatom clock.

The results obtained on RITMO1 are of cardinal importance as they demonstrate for the first time the existence of a molecular circadian clock in diatoms. They also insert the first element in the puzzle represented by the architecture of this endogenous oscillator. However, circadian timekeeping mechanisms cannot rely on a single component and, as described in the previous pages, require the participation of multiple regulators capable of generating self-sustained feedback loops. For this reason, part of my work has also focused on the search for possible partners of RITMO1. As reported in chapter II.d, I was hence able to demonstrate that a second bHLH-PAS transcription factor, bHLH1b, is able to physically interact with RITMO1 in yeast cells and, if obliterated by CRISPR/Cas9, leads to phenotypic alterations similar to those observed in RITMO1 mutants.
The second light-dependent process I have focused on is photosynthesis and the regulation of plastid activity. Considerable efforts have already been conducted by the scientific community to identify and characterize the proteins involved in chloroplast activity. As secondary endosymbionts, diatoms present several peculiarities in the photosynthetic apparatus and important knowledge has been accumulated on the proteins that are directly involved in its protection to excessive light stimuli. However, the regulatory mechanisms that modulate these processes and their response to the environment still remain largely unexplored. For this reason, during my Ph.D. I participated in the validation of a screening strategy aimed at identifying transcription factors involved in the regulation of photosynthesis in *P. tricornutum*, as reported in Chapter III. This led to the characterization of transgenic lines possessing an antisense fragment for the seven *P. tricornutum* TFs possessing a bHLH domain and to the identification of significant alterations in photosynthetic activity in *bHLH2* RNAi transgenic lines. Like RITMO1 and bHLH1b, bHLH2 is a transcription factor that possesses a PAS fold and a bHLH domain. The bulk of my work therefore focused on detailed dissection of the plastid physiology of *bHLH2*RNAi lines. Such work has allowed me to demonstrate that bHLH2 is a candidate regulator of plastid response to light stimuli with *bHLH2*RNAi lines exhibiting a defect in photosynthesis and growth that is found to be dependent on light intensity. In addition, I demonstrated how the expression of bHLH2, encoded in the nucleus, is regulated in turn by photosynthetic activity in a process of retrograde signaling. Notably, before being identified in *Stramenopila*, transcription factors containing both bHLH and PAS were thought to be a peculiarity of metazoans. The work presented in this thesis clearly reveals that bHLH-PAS proteins acquired functions critical to the life of phototrophic organisms during the evolution of secondary endosymbionts.
Bibliography

Ait-Mohamed, O., Novák Vanclová, A.M.G., Joli, N., Liang, Y., Zhao, X., Genovesio, A., Tirichine, L., Bowler, C., Dorrell, R.G., 2020. PhaeoNet: A Holistic RNAseq-Based Portrait of Transcriptional Coordination in the Model Diatom Phaeodactylum tricornutum. Front. Plant Sci. 11, 590949. https://doi.org/10.3389/fpls.2020.590949

Alabadí, D., Oyama, T., Yanovsky, M.J., Harmon, F.G., Más, P., Kay, S.A., 2001. Reciprocal Regulation Between *TOC1* and *LHY/CCA1* Within the *Arabidopsis* Circadian Clock. Science 293, 880-883. https://doi.org/10.1126/science.1061320

Allen, A.E., Dupont, C.L., Oborník, M., Horák, A., Nunes-Nesi, A., McCrow, J.P., Zheng, H., Johnson, D.A., Hu, H., Fernie, A.R., Bowler, C., 2011. Evolution and metabolic significance of the urea cycle in photosynthetic diatoms. Nature 473, 203-207. https://doi.org/10.1038/nature10074

Allen, J.F., de Paula, W.B.M., Puthiyaveetil, S., Nield, J., 2011. A structural phylogenetic map for chloroplast photosynthesis. Trends Plant Sci. 16, 645-655. https://doi.org/10.1016/j.tplants.2011.10.004

Anderson, C.R., Kudela, R.M., Benitez-Nelson, C., Sekula-Wood, E., Burrell, C.T., Chao, Y., Langlois, G., Goodman, J., Siegel, D.A., 2011. Detecting toxic diatom blooms from ocean color and a regional ocean model: REMOTE DETECTION OF TOXIC DIATOM BLOOMS. Geophys. Res. Lett. 38, n/a-n/a. https://doi.org/10.1029/2010GL045858

Andreatta, G., Tessmar-Raible, K., 2020. The Still Dark Side of the Moon: Molecular Mechanisms of Lunar-Controlled Rhythms and Clocks. J. Mol. Biol. 432, 3525-3546. https://doi.org/10.1016/j.jmb.2020.03.009

Andrés, F., Coupland, G., 2012. The genetic basis of flowering responses to seasonal cues. Nat. Rev. Genet. 13, 627-639. https://doi.org/10.1038/nrg3291

Anning, T., MacIntyre, H.L., Pratt, S.M., Sammes, P.J., Gibb, S., Geider, R.J., 2000. Photoacclimation in the marine diatom *Skeletonema costatum*. Limnol. Oceanogr. 45, 1807-1817. https://doi.org/10.4319/lo.2000.45.8.1807

Annunziata, R., Ritter, A., Fortunato, A.E., Manzotti, A., Cheminant-Navarro, S., Agier, N., Huysman, M.J.J., Winge, P., Bones, A.M., Bouget, F.-Y., Cosentino Lagomarsino, M., Bouly, J.-P., Falciatore, A., 2019. bHLH-PAS protein RITMO1 regulates diel biological rhythms in the marine diatom *Phaeodactylum tricornutum*. Proc. Natl. Acad. Sci. 116, 13137-13142. https://doi.org/10.1073/pnas.1819660116

Apt, K.E., Grossman, A.R., Kroth-Pancic, P.G., 1996. Stable nuclear transformation of the diatom *Phaeodactylum tricornutum*. Mol. Gen. Genet. MGG 252, 572-579. https://doi.org/10.1007/BF02172403

Armbrust, E.V., 2004. The Genome of the Diatom Thalassiosira Pseudonana: Ecology, Evolution, and Metabolism. Science 306, 79-86. https://doi.org/10.1126/science.1101156

Aschoff, J., 1999. Masking and parametric effects of high-frequency light-dark cycles. Jpn. J. Physiol. 49, 11-18. https://doi.org/10.2170/jjphysiol.49.11

Aschoff, J., 1954. Zeitgeber der tierischen Tagesperiodik. Naturwissenschaften 41, 49-56. https://doi.org/10.1007/BF00634164

Ashworth, J., Coesel, S., Lee, A., Armbrust, E.V., Orellana, M.V., Baliga, N.S., 2013. Genome-wide diel growth state transitions in the diatom Thalassiosira pseudonana. Proc. Natl. Acad. Sci. 110, 7518-7523. https://doi.org/10.1073/pnas.1300962110

Babenko, I., Friedrich, B.M., Kröger, N., 2022. Structure and Morphogenesis of the Frustule, in: Falciatore, A., Mock, T. (Eds.), The Molecular Life of Diatoms. Springer International Publishing, Cham, pp. 287-312. https://doi.org/10.1007/978-3-030-92499-7_11

Bailleul, B., Berne, N., Murik, O., Petroutsos, D., Prihoda, J., Tanaka, A., Villanova, V., Bligny, R., Flori, S., Falconet, D., Krieger-Liszkay, A., Santabarbara, S., Rappaport, F., Joliot, P., Tirichine, L., Falkowski, P.G., Cardol, P., Bowler, C., Finazzi, G., 2015. Energetic coupling between plastids and mitochondria drives CO2 assimilation in diatoms. Nature 524, 366-369. https://doi.org/10.1038/nature14599
Bailleul, B., Rogato, A., de Martino, A., Coesel, S., Cardol, P., Bowler, C., Falciatore, A., Finazzi, G., 2010. An atypical member of the light-harvesting complex stress-related protein family modulates diatom responses to light. Proc. Natl. Acad. Sci. 107, 18214-18219.
https://doi.org/10.1073/pnas.1007703107

Baker, C.L., Loros, J.J., Dunlap, J.C., 2012. The circadian clock of Neurospora crassa. Ederation Eur. Microbiol. Soc. Rev. 35, 95-110. https://doi.org/10.1111/j.1574-6976.2011.00288.x

Baldauf, S.L., 2008. An overview of the phylogeny and diversity of eukaryotes. J. Syst. Evol. 46, 11.

Ballottari, M., Truong, T.B., De Re, E., Erickson, E., Stella, G.R., Fleming, G.R., Bassi, R., Niyogi, K.K., 2016. Identification of pH-sensing Sites in the Light Harvesting Complex Stress-related 3 Protein Essential for Triggering Non-photochemical Quenching in Chlamydomonas reinhardtii. J. Biol. Chem. 291, 7334-7346. https://doi.org/10.1074/jbc.M115.704601

Bargiello, T.A., Jackson, F.R., Young, M.W., 1984. Restoration of circadian behavioural rhythms by gene transfer in Drosophila. Nature 312, 752-754. https://doi.org/10.1038/312752a0

Bar-On, Y.M., Phillips, R., Milo, R., 2018. The biomass distribution on Earth. Syst. Biol. 67, 1111-1129. https://doi.org/10.1073/pnas.1711842115

Basu, S., Patil, S., Mapleson, D., Russo, M.T., Vitale, L., Fevola, C., Maumus, F., Casotti, R., Mock, T., Caccamo, M., Montresor, M., Sanges, R., Ferrante, M.I., 2017. Finding a partner in the ocean: molecular and evolutionary bases of the response to sexual cues in a planktonic diatom. New Phytol. 215, 140-156. https://doi.org/10.1111/nph.14557

Beadle, G.W., Tatum, E.L., 1941. Genetic Control of Biochemical Reactions in Neurospora. Proc. Natl. Acad. Sci. 27, 499-506. https://doi.org/10.1073/pnas.27.11.499

Bertrand, M., 2010. Carotenoid biosynthesis in diatoms. Photosynth. Res. 106, 89-102.
https://doi.org/10.1007/s11120-010-9589-x

Biggins, J., Bruce, D., 1989. Regulation of excitation energy transfer in organisms containing phycobilins. Photosynth. Res. 20, 1-34. https://doi.org/10.1007/BF00028620

Bilcke, G., Osuna-Cruz, C.M., Silva, M.S., Poulsen, N., Bulankova, P., Vyverman, W., Veylder, L.D., Vandepoele, K., 2021a. Diurnal transcript profiling of the diatom Seminavis robusta reveals adaptations to a benthic lifestyle. Plant J. 107. https://doi.org/10.1111/tpj.15291

Bilcke, G., Van Craenenbroeck, L., Castagna, A., Osuna-Cruz, C.M., Vandepoele, K., Sabbe, K., De Veylder, L., Vyverman, W., 2021b. Light intensity and spectral composition drive reproductive success in the marine benthic diatom Seminavis robusta. Sci. Rep. 11, 17560. https://doi.org/10.1038/s41598-021-92838-0

Blaby-Haas, C.E., Merchant, S.S., 2019. Comparative and Functional Algal Genomics. Annu. Rev. Plant Biol. 70, 605-638. https://doi.org/10.1146/annurev-arplant-050718-095841

Blommaert, L., Chafai, L., Bailleul, B., 2021. The fine-tuning of NPQ in diatoms relies on the regulation of both xanthophyll cycle enzymes. Sci. Rep. 11, 12750. https://doi.org/10.1038/s41598-021-91483-x

Bolige, A., Hagiwara, S., Zhang, Y., Goto, K., 2005. Circadian G2 Arrest as Related to Circadian Gating of Cell Population Growth in Euglena. Plant Cell Physiol. 46, 931-936. https://doi.org/10.1093/pcp/pci100

Bowler, C., Allen, A.E., Badger, J.H., Grimwood, J., Jabbari, K., Kuo, A., Maheswari, U., Martens, C., Maumus, F., Otillar, R.P., Rayko, E., Salamov, A., Vandepoele, K., Beszteri, B., Gruber, A., Heijde, M., Katinka, M., Mock, T., Valentin, K., Verret, F., Berges, J.A., Brownlee, C., Cadoret, J.-P., Chiovitti, A., Choi, C.J., Coesel, S., De Martino, A., Detter, J.C., Durkin, C., Falciatore, A., Fournet, J., Haruta, M., Huysman, M.J.J., Jenkins, B.D., Jiroutova, K., Jorgensen, R.E., Joubert, Y., Kaplan, A., Kröger, N., Kroth, P.G., La Roche, J., Lindquist, E., Lommer, M., Martin-Jézéquel, V., Lopez, P.J., Lucas, S., Mangogna, M., McGinnis, K., Medlin, L.K., Montsant, A., Secq, M.-P.O., Napoli, C., Obornik, M., Parker, M.S., Petit, J.-L., Porcel, B.M., Poulsen, N., Robison, M., Rychlewski, L., Rynearson, T.A., Schmutz, J., Shapiro, H., Siaut, M., Stanley, M., Sussman, M.R., Taylor, A.R., Vardi, A., von Dassow, P.,
Vyverman, W., Willis, A., Wyrwicz, L.S., Rokhsar, D.S., Weissenbach, J., Armbrust, E.V., Green, B.R., Van de Peer, Y., Grigoriev, I.V., 2008. The Phaeodactylum genome reveals the evolutionary history of diatom genomes. Nature 456, 239-244.
https://doi.org/10.1038/nature07410

Bowler, C., Falciatore, A., 2019. Phaeodactylum tricornutum. Trends Genet. 35, 706-707.
https://doi.org/10.1016/j.tig.2019.05.007

Bradshaw, W.E., Holzapfel, C.M., 2007. Evolution of Animal Photoperiodism. Annu. Rev. Ecol. Evol. Syst. 38, 1-25. https://doi.org/10.1146/annurev.ecolsys.37.091305.110115

Brand, L.E., 1982. Persistent diel rhythms in the chlorophyll fluorescence of marine phytoplankton species. Mar. Biol. 69, 253-262. https://doi.org/10.1007/BF00397491

Brand, L.E., Guillard, R.R.L., 1981. The effects of continuous light and light intensity on the reproduction rates of twenty-two species of marine phytoplankton. J. Exp. Mar. Biol. Ecol. 50, 119-132. https://doi.org/10.1016/0022-0981(81)90045-9

Brown, F., Webb, M., 1948. Temperature Relations of an Endogenous Daily Rhythmicity in the Fiddler Crab, Uca. Physiol. Zool. 21. https://doi.org/10.1086/physzool.21.4.30152016

Bruce, V.G., 1974. RECOMBINANTS BETWEEN CLOCK MUTANTS OF *CHLAMYDOMONAS* REINHARDI. Genetics 77, 221-229.
https://doi.org/10.1093/genetics/77.2.221

Bruce, V.G., 1972. Mutants of the biological clock in Chlamydomonas reinhardtii. Genetics 70, 537-548. https://doi.org/10.1093/genetics/70.4.537

Bruce, V.G., 1970. The biological clock in Chlamydomonas reinhardtii. J. Protozool. 17, 328-334.
https://doi.org/10.1111/j.1550-7408.1970.tb02380.x

Bruce, V.G., Pittendrigh, C.S., 1956. TEMPERATURE INDEPENDENCE IN A UNICELLULAR "CLOCK." Proc. Natl. Acad. Sci. 42, 676-682.
https://doi.org/10.1073/pnas.42.9.676

Brunelle, S.A., Hazard Starr, A., Sotka, E.E., Van Dolah, F.M., 2007. Characterization Of A Dinoflagellate Cryptochrome Blue-Light Receptor With A Possible Role In Circadian Control Of The Cell Cycle. J. Phycol. 43, 509-518. https://doi.org/10.1111/j.1529-8817.2007.00339.x

Brzezinski, M., Olson, R.J., Chisholm, S.W., 1990. Silicon availability and cell-cycle progression in marine diatoms. Mar. Ecol. Prog. Ser. 67, 83-96.

Buck, J.M., Kroth, P.G., Lepetit, B., 2021. Identification of sequence motifs in Lhcx proteins that confer qE-based photoprotection in the diatom *Phaeodactylum tricornutum*. Plant J. 108, 1721-1734. https://doi.org/10.1111/tpj.15539

Buck, J.M., Sherman, J., Bártulos, C.R., Serif, M., Halder, M., Henkel, J., Falciatore, A., Lavaud, J., Gorbunov, M.Y., Kroth, P.G., Falkowski, P.G., Lepetit, B., 2019. Lhcx proteins provide photoprotection via thermal dissipation of absorbed light in the diatom Phaeodactylum tricornutum. Nat. Commun. 10, 4167. https://doi.org/10.1038/s41467-019-12043-6

Bünning, E., 1936. Die Endogene Tagesrhythmisches Grundlage Der Photoperiodischen Reaktion, Berichte der Deutschen Botanischen Gesellschaft 57, 590-607.

Bünning, E., 1935. Zur kenntnis der erblichen tagesperiodizität bei den primarblattern von Phaseolus multiflorus. Jahrb. Für Wiss. Bot. 81.

Burki, F., Roger, A.J., Brown, M.W., Simpson, A.G.B., 2020. The New Tree of Eukaryotes. Trends Ecol. Evol. 35, 43-55. https://doi.org/10.1016/j.tree.2019.08.008

Cahonn, L.B., 1999. The role of benthic microalgae in neritic ecosystems, in: The Role of Benthic Microalgae in Neritic Ecosystems. CRC Press, p. 40.

Carneiro, B.T.S., Araujo, J.F., 2012. Food entrainment: major and recent findings. Front. Behav. Neurosci. 6. https://doi.org/10.3389/fnbeh.2012.00083
Carré, I.A., Edmunds, L.N., 1993. Oscillator control of cell division in Euglena: cyclic AMP oscillations mediate the phasing of the cell division cycle by the circadian clock. J. Cell Sci. 104, 11. https://doi.org/10.1242/jcs.104.4.1163

Carroll, P.A., Freie, B.W., Mathsyaraja, H., Eisenman, R.N., 2018. The MYC transcription factor network: balancing metabolism, proliferation and oncogenesis. Front. Med. 12, 412-425. https://doi.org/10.1007/s11684-018-0650-z

Castillon, A., Shen, H., Huq, E., 2007. Phytochrome Interacting Factors: central players in phytochrome-mediated light signaling networks. Trends Plant Sci. 12, 514-521. https://doi.org/10.1016/j.tplants.2007.10.001

Chan, C.X., Bhattacharya, D., Reyes-Prieto, A., 2012. Endosymbiotic and horizontal gene transfer in microbial eukaryotes: Impacts on cell evolution and the tree of life. Mob. Genet. Elem. 2, 101-105. https://doi.org/10.4161/mge.20110

Chang, J., Carpenter, E., 1991. Species-specific phytoplankton growth rates via diel DNA synthesis cycles. V. Application to natural populations in Long Island Sound. Mar. Ecol. Prog. Ser. 78, 115-122. https://doi.org/10.3354/meps078115

Chauton, M.S., Winge, P., Brembu, T., Vadstein, O., Bones, A.M., 2013. Gene Regulation of Carbon Fixation, Storage, and Utilization in the Diatom *Phaeodactylum tricornutum* Acclimated to Light/Dark Cycles. Plant Physiol. 161, 1034-1048. https://doi.org/10.1104/pp.112.206177

Chen, C.-H., Dunlap, J.C., Loros, J.J., 2010. Neurospora illuminates fungal photoreception. Fungal Genet. Biol. 47, 922-929. https://doi.org/10.1016/j.fgb.2010.07.005

Cheng, P., 2001. Coiled-coil domain-mediated FRQ-FRQ interaction is essential for its circadian clock function in Neurospora. EMBO J. 20, 101-108. https://doi.org/10.1093/emboj/20.1.101

Cheng, P., He, Qun, He, Qiyang, Wang, L., Liu, Y., 2005. Regulation of the *Neurospora* circadian clock by an RNA helicase. Genes Dev. 19, 234-241. https://doi.org/10.1101/gad.1266805

Cheng, P., Yang, Y., Gardner, K.H., Liu, Y., 2002. PAS Domain-Mediated WC-1/WC-2 Interaction Is Essential for Maintaining the Steady-State Level of WC-1 and the Function of Both Proteins in Circadian Clock and Light Responses of *Neurospora*. Mol. Cell. Biol. 22, 517-524. https://doi.org/10.1128/MCB.22.2.517-524.2002

Chepurnov, V.A., Mann, D.G., Sabbe, K., Vyverman, W., 2004. Experimental Studies on Sexual Reproduction in Diatoms, in: International Review of Cytology. Elsevier, pp. 91-154. https://doi.org/10.1016/S0074-7696(04)37003-8

Chisholm, S.W., Brand, L.E., 1981. Persistence of cell division phasing in marine phytoplankton in continuous light after entrainment to light: Dark cycles. J. Exp. Mar. Biol. Ecol. 51, 107-118. https://doi.org/10.1016/0022-0981(81)90123-4

Christie, J.M., Reymond, P., Powell, G.K., Bernasconi, P., Raibekas, A.A., Liscum, E., Briggs, W.R., 1998. *Arabidopsis* NPH1: A Flavoprotein with the Properties of a Photoreceptor for Phototropism. Science 282, 1698-1701. https://doi.org/10.1126/science.282.5394.1698

Christie, J.M., Salomon, M., Nozue, K., Wada, M., Briggs, W.R., 1999. LOV (light, oxygen, or voltage) domains of the blue-light photoreceptor phototropin (nph1): Binding sites for the chromophore flavin mononucleotide. Proc. Natl. Acad. Sci. 96, 8779-8783. https://doi.org/10.1073/pnas.96.15.8779

Chukhutsina, V.U., Büchel, C., van Amerongen, H., 2014. Disentangling two non-photochemical quenching processes in Cyclotella meneghiniana by spectrally-resolved picosecond fluorescence at 77K. Biochim. Biophys. Acta BBA - Bioenerg. 1837, 899-907. https://doi.org/10.1016/j.bbabio.2014.02.021

Coesel, S., Mangogna, M., Ishikawa, T., Heijde, M., Rogato, A., Finazzi, G., Todo, T., Bowler, C., Falciatore, A., 2009. Diatom PtCPF1 is a new cryptochrome/photolyase family member with