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# Light sensing in the Ocean: studying diatom phytochrome photoreceptors

Carole Duchêne

► To cite this version:

Carole Duchêne. Light sensing in the Ocean: studying diatom phytochrome photoreceptors. Morphogenesis. Sorbonne Université, 2022. English. NNT: 2022SORUS164. tel-04145175

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https://theses.hal.science/tel-04145175

Submitted on 29 Jun 2023

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# Sorbonne Université

## École doctorale Complexité du Vivant

Laboratoire biologie du chloroplaste et perception de la lumière chez les micro-algues

## Light sensing in the Ocean :
studying diatom phytochrome photoreceptors

Par Carole Duchêne

Thèse de doctorat de Biologie

Dirigée par Dr. Angela Falciatore

Co-encadrante Dr. JAUBERT Marianne

Présentée et soutenue publiquement le 28 juin 2022

Devant un jury composé de :

<table><tr><td>GROSSMAN Arthur</td><td>Prof. Dr. Carnegie Institution for Science<br>Stanford University</td><td>Rapporteur</td></tr><tr><td>PETROUTSOS Dimitris</td><td>Chercheur CNRS</td><td>Rapporteur</td></tr><tr><td>BHAYA Devaki</td><td>Prof. Dr. Carnegie Institution for Science<br>Stanford University</td><td>Examinatrice</td></tr><tr><td>BABIN Marcel</td><td>Professeur d'Université, Université de Laval</td><td>Examinateur</td></tr><tr><td>CARBONE Alessandra</td><td>Professeur d'Université, Sorbonne Université</td><td>Examinatrice</td></tr><tr><td>FANKHAUSER Christian</td><td>Professeur d'Université, Université de Lausanne</td><td>Examinateur</td></tr><tr><td>FALCIATORE Angela</td><td>Directrice de recherche CNRS</td><td>Directrice de thèse</td></tr><tr><td>JAUBERT Marianne</td><td>Maître de conférences Sorbonne Université</td><td>Co-encadrante de thèse</td></tr></table>
*Choose a job you love, and you will never have to work a day in your life.*

Confucius
This thesis is obviously a team work, and I would like to thank all the people who
surrounded me, supervised and supported me during the whole duration of my thesis. It
has been a pleasure to develop this research project here.

I would like to start by thanking Angela, my thesis director, for giving me the opportunity to join this project (and many others!). Her enthusiasm for science and her passion for diatoms are a driving force for our research.

I thank Marianne for her patience, her rigor and her calm in all circumstances. Thank you for the freedom that was given to me in this thesis work. It was a pleasure to work with you.

I would like to underline the involvement of Jean-Pierre Bouly, who, even if he is not officially the supervisor of my thesis, has clearly been indispensable in the research that is presented here. Thank you also for the discussions and the hypotheses that always carry us further (and for the beers too).

I salute Alessandro, we started our theses (and masters!) together and we finish them two
months apart. Thank you for your presence, your style and your ideals.

I would like to thank my colleagues of the UMR7238, where I started my PhD experience and of the UMR7141: the basement community, including an office only of students; thanks to Sandrine and Soizic who make this lab run, and for your enthusiasm for microscopy. Thank you to all those with whom I ate OUTSIDE every lunchtime and in all weathers during these last two years. The pandemic has slowed our social interactions, but this daily break has remained convivial. Thank you for the lab meetings of several hours, because these discussions are extremely formative.

Thank you to the whole UMR7141 for this research atmosphere and this dynamic so
enriching. Thank you for the lab meetings of several hours, because these discussions are
extremely formative.

Thank you all for putting up with my whistling and singing.

I would like to warmly thank Dr. Chris Bowler, for your interest in this project, even in the
Arctic ice. I would like to thank you and Prof. Marcel Babin for the opportunity to live this
experience. I want also to thank Dr. Juan Pierella Karlusich for his advice and patience.

I want to thank Dr. Maurizio Ribera d'Alcalà for his enthusiasm about my results, and for
the discussions that followed.

Finally, I would like to thank all members of the jury for accepting to examine my work.
TABLE OF CONTENTS

<table><tr><td>Introduction</td><td>9</td></tr><tr><td>Diatom roles in the biogeochemical cycles of carbon and silica</td><td>10</td></tr><tr><td>Diatom origin and evolution</td><td>11</td></tr><tr><td>Life in the ocean: characteristics of the diatom environment</td><td>15</td></tr><tr><td>Cellular aspects of diatom life and their regulation by light</td><td>26</td></tr><tr><td>Light sensing</td><td>32</td></tr><tr><td>Model species and genetic tools</td><td>50</td></tr><tr><td>Objective of the Thesis</td><td>55</td></tr></table>

<table><tr><td>Chapter1: Photoreceptors repertoire and distribution in Ochrophyta</td><td>59</td></tr><tr><td>Abstract</td><td>61</td></tr><tr><td>Introduction</td><td>62</td></tr><tr><td>Results and Discussion</td><td>66</td></tr><tr><td>Conclusions</td><td>87</td></tr><tr><td>Material and methods</td><td>89</td></tr><tr><td>Bibliography</td><td>92</td></tr><tr><td>Supplementary Figures</td><td>104</td></tr><tr><td>Supplementary Data</td><td>116</td></tr></table>

<table><tr><td>Chapter 2: Marine diatom phytochromes are sensors of depth and phytoplankton concentration through sensing of underwater light variations</td><td>119</td></tr><tr><td>Abstract</td><td>121</td></tr><tr><td>Introduction</td><td>122</td></tr><tr><td>Results</td><td>125</td></tr><tr><td>Discussion</td><td>141</td></tr><tr><td>Material & Methods</td><td>146</td></tr><tr><td>References</td><td>152</td></tr><tr><td>Supplementary Figures</td><td>160</td></tr><tr><td>Supplementary Tables</td><td>172</td></tr><tr><td>Supplementary Data</td><td>173</td></tr><tr><td>Supplemental Method</td><td>174</td></tr></table>

<table><tr><td>Chapter 3: DPH in benthic diatom species</td><td>181</td></tr><tr><td>Results</td><td>183</td></tr><tr><td>Discussion</td><td>190</td></tr><tr><td>Material and methods</td><td>195</td></tr></table>
<table><tr><td>Supplementary Figure</td><td>197</td></tr></table>

<table><tr><td>Chapter 4: DPH sinalization</td><td>199</td></tr></table>

<table><tr><td>Results</td><td>200</td></tr></table>

<table><tr><td>Conclusion</td><td>215</td></tr></table>

<table><tr><td>Material and methods</td><td>215</td></tr></table>

<table><tr><td>Discussion</td><td>221</td></tr></table>

<table><tr><td>Bibliography</td><td>231</td></tr></table>

<table><tr><td>Annexes</td><td>251</td></tr></table>
LIST OF FIGURES AND TABLES

<table><tr><td>Introduction</td><td>9</td></tr><tr><td>Figure 1. Diatom evolution</td><td>12</td></tr><tr><td>Figure 2. Diversity of diatoms</td><td>14</td></tr><tr><td>Figure 3. Environmental context for phytoplankton life in the water column</td><td>17</td></tr><tr><td>Figure 4. Example of vertical profile of oxygen and light in two types of sediments</td><td>19</td></tr><tr><td>Figure 5. Ice associated algal communities and their dynamics</td><td>21</td></tr><tr><td>Figure 6. Examples of underwater light fields in different aquatic environments</td><td>23</td></tr><tr><td>Figure 7. Light field sediments and in ice</td><td>25</td></tr><tr><td>Figure 8. Diatom life cycle</td><td>27</td></tr><tr><td>Figure 9. The photosynthetic light reactions in diatoms</td><td>30</td></tr><tr><td>Figure 10. Sensing of light cues in diatoms</td><td>39</td></tr><tr><td>Figure 11. Phylogenetic analysis phytochrome photoreceptor</td><td>42</td></tr><tr><td>Figure 12. Signaling mechanisms in phytochromes</td><td>45</td></tr><tr><td>Figure 13. Algal phytochromes absorption spectra</td><td>49</td></tr><tr><td>Table 1. Sequenced genomes of centric diatoms</td><td>51</td></tr><tr><td>Table 2. Sequenced genomes of pennate diatoms</td><td>52</td></tr></table>

<table><tr><td>Chapter1: Photoreceptors repertoire and distribution in Ochrophyta</td><td>59</td></tr><tr><td>Figure 1. Photoreceptors repertoire in Stramenopiles</td><td>67</td></tr><tr><td>Figure 2. LOV-based photoreceptors in Ochrophyta</td><td>72</td></tr><tr><td>Figure 3. Ochrophyta CPF family protein tree</td><td>78</td></tr><tr><td>Figure 4. Phylogeny of the rhodopsins</td><td>83</td></tr><tr><td>Figure 5. Phylogeny of the phytochromes</td><td>85</td></tr><tr><td>Table 1. Details of the parameters and reference sequences used in our photoreceptor search</td><td>90</td></tr><tr><td>Figure S1. Stramenopile species tree</td><td>104</td></tr><tr><td>Figure S2. Bioinformatic analysis of the putative LOV-containing photoreceptors</td><td>105</td></tr><tr><td>Figure S3. LOV-based photoreceptors families in Ochrophyta</td><td>106</td></tr><tr><td>Figure S4. Analysis of the aureochrome and helmchromes</td><td>107</td></tr><tr><td>Figure S5. Analysis of the CPF family in Ochrophyta</td><td>108</td></tr><tr><td>Figure S6. Zoom on the CPD branch of the CPF tree</td><td>109</td></tr><tr><td>Figure S7. Zoom on the Cry-DASH branch of the CPF tree</td><td>110</td></tr><tr><td>Figure S8. The 6-4 photolyases</td><td>111</td></tr><tr><td>Figure S9. The plant-Cry like proteins</td><td>112</td></tr><tr><td>Figure S10. Sequence analysis of the CryP sub-families</td><td>113</td></tr><tr><td>Figure S11. Repartition of rhodopsins and phytochromes in Stramenopiles</td><td>114</td></tr><tr><td>Figure S12. Analysis of the rhodopsin photoreceptor types</td><td>115</td></tr><tr><td>Figure S13. Alignment of selected phytochrome sequences</td><td>116</td></tr></table>
<table>
  <tbody>
    <tr>
      <td><strong>Chapter 2: Marine diatom phytochromes are sensors of depth and phytoplankton concentration through sensing of underwater light variations</strong></td>
      <td>119</td>
    </tr>
    <tr>
      <td>Figure 1. PtDPH activity reporter system and experimental conditions</td>
      <td>126</td>
    </tr>
    <tr>
      <td>Figure 2. Action and inhibition spectra for PtDPH-mediated responses</td>
      <td>128</td>
    </tr>
    <tr>
      <td>Figure 3. Modelling of PtDPH activity in vivo</td>
      <td>130</td>
    </tr>
    <tr>
      <td>Figure 4. PtDPH activity in modeled environmental fields</td>
      <td>133</td>
    </tr>
    <tr>
      <td>Figure 5. PtDPH activity in situ measured field fields</td>
      <td>134</td>
    </tr>
    <tr>
      <td>Figure 6. DPH distribution is linked to latitude, temperature and optical parameters</td>
      <td>136</td>
    </tr>
    <tr>
      <td>Figure 7. Spectral properties of DPH from various diatoms</td>
      <td>139</td>
    </tr>
    <tr>
      <td>Table 1. Spectral and photochemical properties of different recombinant DPH</td>
      <td>140</td>
    </tr>
    <tr>
      <td>Figure S1. Experimental setup for the action and inhibition spectra</td>
      <td>160</td>
    </tr>
    <tr>
      <td>Figure S2. Control lines show no induction of YFP in the action spectra</td>
      <td>160</td>
    </tr>
    <tr>
      <td>Figure S3. Expression analysis of PtDPH-regulat and PtDPH-non regulated genes upon far-red and blue irradiation. P. tricornutum WT and DPH KO mutant</td>
      <td>161</td>
    </tr>
    <tr>
      <td>Figure S4. Light modeling examples</td>
      <td>162</td>
    </tr>
    <tr>
      <td>Figure S5. DPH activity in different modelled environmental condition</td>
      <td>163</td>
    </tr>
    <tr>
      <td>Figure S6. DPH activity in situ measured environmental light fields over a year time series</td>
      <td>164</td>
    </tr>
    <tr>
      <td>Figure S7. Biogeography of DPH and Aureochromes (Aureo) from centric diatoms</td>
      <td>164</td>
    </tr>
    <tr>
      <td>Figure S8. Metagenome Assembled Genomes (MAGs) geographical distribution of centric diatoms</td>
      <td>165</td>
    </tr>
    <tr>
      <td>Figure S9. DPH relative abundance compared to Aureochrome relative abundance</td>
      <td>166</td>
    </tr>
    <tr>
      <td>Figure S10. Correlation and individual generalized additive models analysis for DPH and Aureochromes relative abundance</td>
      <td>167</td>
    </tr>
    <tr>
      <td>Figure S11. Complex generalized additive models explaining DPH relative abundance with a combination of environmental parameters for each filter size</td>
      <td>168</td>
    </tr>
    <tr>
      <td>Figure S12. Alignment of a portion of the PAS and GAF domains of selected DPH and reference phytochrome sequences</td>
      <td>168</td>
    </tr>
    <tr>
      <td>Figure S13. Purified recombinant DPH</td>
      <td>169</td>
    </tr>
    <tr>
      <td>Figure S14. TpDPH restores blue and far-red-dependent YFP induction in PtDPH KO line</td>
      <td>169</td>
    </tr>
    <tr>
      <td>Figure S15. Projection of DPHs in modeled light fields</td>
      <td>170</td>
    </tr>
    <tr>
      <td>Figure S16. Method for DPH and diatom Aureochrome gene search</td>
      <td>171</td>
    </tr>
    <tr>
      <td>Table S1. Species from which DPH have been characterized in this study</td>
      <td>172</td>
    </tr>
    <tr>
      <td>Table S2. Sequences of primers used in this study</td>
      <td>172</td>
    </tr>
    <tr>
      <td>Table S3. SSN parameters for DPH and diatom Aureochrome search in Tara Oceans data</td>
      <td>173</td>
    </tr>
  </tbody>
</table>

<table>
  <tbody>
    <tr>
      <td><strong>Chapter 3: DPH in benthic diatom species</strong></td>
      <td>181</td>
    </tr>
    <tr>
      <td>Table 1. Pennate diatom species with phytochrome(s).</td>
      <td>183</td>
    </tr>
    <tr>
      <td>Figure 1. Absorption spectra and difference spectra between the Pr and Pfr forms for 4 A. coffeaeformis phytochromes.</td>
      <td>184</td>
    </tr>
    <tr>
      <td>Figure 2. Fluorescence emission spectra at 77K of P. tricornutum</td>
      <td>187</td>
    </tr>
    <tr>
      <td>Figure 3. DPH controls F710 amplitude in response to increased FR/R ratio</td>
      <td>188</td>
    </tr>
  </tbody>
</table>
<table><tr><td>Figure 4. Kinetics of induction of F710</td><td>188</td></tr><tr><td>Figure 5. 7K fluorescence spectra of centric diatom species</td><td>198</td></tr><tr><td>Figure 6. 7K fluorescence spectra of different pennate diatom species</td><td>190</td></tr><tr><td>Figure S1. DPH controls F710 amplitude in response to increased FR/R ratio</td><td>196</td></tr><tr><td>Table S1. Primers used to clone A. coffeaeformis DPHs</td><td>197</td></tr></table>

<table><tr><td>Chapter 4: DPH Signalization</td><td>199</td></tr><tr><td>Figure 1. Phytochrome chromophores biosynthesis pathways</td><td>201</td></tr><tr><td>Table 1. Characteristic of the candidate Heme oxygenase in Phaeodactylum tricornutum</td><td>202</td></tr><tr><td>Figure 2. Spectral properties of TweiDPH expressed SynHO1)</td><td>203</td></tr><tr><td>Figure 3. Scheme of the vector structure allowing to co-express T. weissflogii DPH photosensory module</td><td>204</td></tr><tr><td>Table 2. Characteristic of the candidate BVR in Phaeodactylum tricornutum</td><td>205</td></tr><tr><td>Figure 4. Spectral properties of TweiDPH expressed with SynHO1 and different PtBVR s</td><td>205</td></tr><tr><td>Figure 5. DPH localization. Venus-PtDPH-expressing P. tricornutum cells</td><td>206</td></tr><tr><td>Figure 6. DPH localization. PtDPH-NeonGreen-expressing P. tricornutum cells</td><td>207</td></tr><tr><td>Figure 7. Venus localization. Venus-expressing P. tricornutum cells</td><td>207</td></tr><tr><td>Figure 8. PtCPF1 localization. Venus-PtCPF1-expressing P. tricornutum cells</td><td>208</td></tr><tr><td>Figure 9. Elements from two-component systems</td><td>209</td></tr><tr><td>Figure 10. DPH-OPM interacts with PtHpt in Yeast Two_Hybrid</td><td>210</td></tr><tr><td>Table 3. Proteins with Response regulators domains in P.tricornutum genome</td><td>212</td></tr><tr><td>Figure 10. DPH-OPM interacts with PtHpt in Yeast Two_Hybrid</td><td>210</td></tr><tr><td>Figure 11. Regulation of ProHsf4.6a by light, cell concentration and agitation</td><td>214</td></tr><tr><td>Figure 12. Proposition of DPH signaling mechanisms in P. tricornutum</td><td>215</td></tr><tr><td>Table S1. Primers data used in this study</td><td>215</td></tr></table>
INTRODUCTION

Diatoms are unicellular photosynthetic eukaryotes that can be found in all wet environments, from freshwater to marine ecosystems, including brackish waters, and extended to sea ice and terrestrial soil(Armbrust, 2009). One of their characteristics is the presence of a silicified cell wall (the “frustule”) surrounding the cell, giving them a diversity of beautiful shapes (Babenko et al., 2022). Although microscopic, diatoms are major primary producers in the ocean and have tremendous roles in the cycle of carbon, silica and nutrients.

In the last 20 years, the availability of genomic information and genetic resources in model species such as *Thalassiosira pseudonana* and *Phaeodactylum tricornutum*, started to unravel the molecular secret of diatoms and unlock diatom potential for biotechnology application.

As photosynthetic organisms, light is for diatoms both a source of energy and of information about their surrounding environment. Light in the open ocean is blue-enriched due to the absorption of red and longer wavelengths by water; in more turbid waters (coastal regions for example), blue is absorbed by other components and the light field is enriched in green light. Light shapes different aspects of diatom life, such as photosynthetic acclimation, cell cycle control, sexual reproduction and movement. These traits are controlled by light-driven processes such as photosynthesis and photoperception, through specialized proteins that absorb light. Genomic studies revealed that diatoms possess different types of photoreceptors: blue light sensing Aureochromes and cryptochrome-like proteins, putative green-sensing rhodopsins, but also red/far-red sensing phytochromes, which is surprising given the light environment diatom live in.
Recent functional investigation showed that diatoms phytochromes (DPH) are bona fide red/far-red photoreceptors *in vitro*, and regulate gene expression in response to far-red light in the model diatom *Phaeodactylum tricornutum*. Despite the availability of DPH knockout in *P. tricornutum*, the physiological function of this photoreceptor is still unknown.

My PhD work was designed to get insights into the role and significance of DPH for diatom life in the marine environment. To address this question, different research approaches were developed. By combining bioinformatics, mathematical modeling of environmental light and DPH response to it, experimental approaches of genetics and biochemistry on molecular model diatoms, this work has provided a much complete view of DPH action in diatoms.

DIATOM ROLES IN THE BIOGEOCHEMICAL CYCLES OF CARBON AND SILICA

70% of our planet's surface is covered by ocean. Marine ecosystems contribute to about 50 Gt of net carbon fixation (GtC) per year, which is equivalent to the contribution of terrestrial ecosystems (Field et al., 1998). If the contribution of land and sea in the global primary production is roughly equal, primary produced biomass is drastically different (Bar-On et al., 2018). Indeed, primary producers represent 450 Gt C on land, sustaining 20GtC of consumers biomass, while in the marine environment there is 1 GtC of primary producers and 5 GtC of consumers (Bar-On et al., 2018). This reversed pyramid is explained by the high turnover rate of fixed carbon in the marine environment, and high efficiency in the transfer to higher trophic levels. Indeed, marine fixed carbon can fuel higher trophic levels with high efficiency (10% of the energy is transferred to higher trophic levels in the ocean, compared to 1% in terrestrial environments (Trebilco et al., 2013), or exported to the depth, where it can be re-mineralized as CO₂ or be buried in the sediments for geological times if reaches the bathypelagic layer.

Among phytoplankton (marine primary producers), diatoms play an important role. They are considered as responsible for 40% of marine primary production (Nelson et al., 1995)
and constitute about 40% of the particulate organic carbon that can be exported to the depth (Jin et al., 2006). This is due to their high sinking rate, which is linked to the diatom's large size and silicified cell wall, and to their tendency to from aggregates (Sarthou et al., 2005). However, large variations in cell size and silicification (Si/C ratio) exists both in different diatom species and in different environmental conditions, and all diatoms do not contribute to the carbon export in the same way (reviewed in Tréguer et al., 2017). Numerical models predicting carbon export in the ocean have to take into account this heterogeneity and the wide diatom diversity in the ocean (Malviya et al., 2016). These models are complex and difficult to accurately parameterize, and the exact contribution of diatoms to biological pump (export of carbon to the depth) remains to be quantified (Tréguer et al., 2017). The role of nanoplanktonic diatoms, which are difficult to observe due to their small size (<5µm), is gaining interest, and there is evidence that although small, these diatoms contribute to the export of carbon (Leblanc et al., 2018).

As the main silicified organism in the ocean, diatoms also play a tremendous role in the cycle of silica (Si), which is linked to the cycle of carbon and nitrogen in the ocean (Tréguer and De La Rocha, 2013). Diatom fix 240 Tmoles per year of dissolved Si in their frustule, of which 6.5 Tmoles are buried into the sediments (Struyf et al., 2009).

Diatoms as primary producers thus play a tremendous role in the marine trophic chain, and their silica cell wall gives them important roles in the ocean biogeochemical cycles.

DIATOM ORIGIN AND EVOLUTION

Diatoms belong to the Stramenopile, Alveaota, Rhizaria (SAR) eukaryotic lineage (Burki et al., 2020). The branching of SAR compared to other major eukaryotic lineages is still uncertain (Fig. 1). Inside the SAR supergroup, diatoms belong to the Stramenopile group, and more precisely to the Ochrophyta clade, which are photosynthetic Stramenopiles. In addition to diatoms, this clade includes a large diversity of ecologically important organisms, especially in marine environments: multicellular brown algae, which are the main primary producers in tidal and sub-tidal regions (next to the shores)(Bringloe et al., 2020), Chrysophytes, that show a variability of trophic modes and can be important
grazers or important primary producers depending on the species and the environment (Olefeld et al., 2018), or Pelagophytes, which can be important components of the picophytoplankton (size<2µm) in tropical regions (Guérin et al., 2021).

<figure><img src="image_2.png" /><figcaption>Figure 1. Diatom evolution. A, position of Stramenopiles in the Eukaryotic tree of life (from (Burki et al., 2020)). B, Close up on the Stramenopile lineage, and position of Ochrophyta (brown branches) and diatoms within Ochrophyta (Orange branches). The red dot indicates the acquisition of photosynthesis in the ancestor of Ochrophyta. Adapted from (Azuma et al., 2022; Dorrell et al., 2022). C, The complex history of endosymbiosis in diatom. a. Primary endosymbiosis and b. Higher order endosymbiosis, giving rise to a plast of red algal origin; Genes from green algae are found in the nuclear genome, suggesting previous association with a green alga. From (Armbrust, 2009)</figcaption></figure>

Primary endosymbiosis occurred about 900 million years ago with the association of a cyanobacteria and a eukaryotic host, giving rise to the Archaeplastidia (Shih and Matzke, 2013). Subsequently, secondary endosymbiosis event, i.e the engulfment and stable association of a photosynthetic eukaryote from the Archaeplastidia lineage by a (heterotroph) eukaryote, independently occurred multiple times during the evolution, with several examples of plastids of green algae origins (Jackson et al., 2018) and red algae
origin (Keeling, 2013). Higher order endosymbiosis events also exist especially in dinoflagellates (Alveolate). Different eukaryotic lineages carry a plastid of red algal origin, such as Cryptophytes, Haptophytes and Ochrophytes. Plastid phylogenies placed red-origin plastids of these algae as a single clade, which led to the idea that secondary endosymbiosis of a red algae chloroplast is a single event that led to the different eukaryotic lineages with red alga chloroplasts (Cryptophytes, Haptophytes and Ochrophytes; this is the chromalveolate hypothesis) (Keeling, 2013). However, large-scale hosts phylogenies place the different eukaryotic hosts on different eukaryotic branches, and the support for this hypothesis has diminished (Burki et al., 2016; Burki et al., 2020). One hypothesis could be that the red-alga derived plastid from different eukaryotic branches is the result of independent symbiosis of closely related red alga. Alternative hypotheses to conciliate plastid and host phylogenies proposed that a single secondary endosymbiotic event (a cryptophyte associating with a red alga) was followed by higher order endosymbiosis events (the ancestor of Ochrophyta adopting a Cryptophyte, followed by the ancestor of haptophyte adopting an early Ochrophyte, although the order is not clearly established): this is the “Cryptophyte first” hypothesis (Stiller et al., 2014).

In Ochrophyta, the early events leading to plastid acquisition are not clearly established, but there is a common agreement that these algae are the result from a single endosymbiosis event, giving rise to their plastid of ultimate red alga origin (red dot in Fig.1B) (Stiller et al., 2014; Burki et al., 2016; Dorrell et al., 2017; Strassert et al., 2021; Azuma et al., 2022; Dorrell et al., 2022).

Diatoms, and Ochrophyta in general, possess plastid-localized proteins, which are encoded by genes of red algal origin in the nucleus; these are supposed to result from endosymbiotic gene transfer from the red alga symbiont to the host. In addition, diatoms possess a number of genes (about 2% of *Phaeodactylum triconutum* genes) of green algal origin, addressed or not to the chloroplast. This could be the remains of an association with a green algae preceding the establishment of the endosymbiosis with the red algae, that gave rise to the actual chloroplast (Moustafa et al., 2009). Diatoms also have genes of bacterial origin (2.3% in *Phaeodactylum tricornutum*), supposed to be the result of horizontal gene transfer. Diatom genomes are thus a mosaic of genes from different
origins, which might explain their ecological success. For example: the LHCX proteins, involved in photoprotection (see also below, photosynthesis section), are supposed of green algal origin; ISIP2a, a membrane protein allowing iron uptake, is also related to green alga proteins; finally, diatom living in sea ice possess ice-binding proteins of bacterial origin (Dorrell et al., 2022).

<figure><img src="image_3.png" /><figcaption>Figure 2. Diversity of diatoms. Diatoms sampled during the Dark Edge cruise in the Arctic in October 2021 (coordinated by Prof. M. Babin). Upper left, *Fragilariopsis sp.* chains (pennate species), middle centric species, probably *Porosira* (upper 2) and *Thalassiosira* species. Right, pennate diatoms, probably *Navicula* (upper and right), the chain forming *Pseudo-Nitzschia* and the needle-like *Cynlindrotheca* (middle).</figcaption></figure>

Diatoms are historically separated into 2 groups based on their shape: the centric diatoms, which are cylindrical, and the more elongated pennate diatoms (Fig. 2). Centric species are further divided into radial and polar centric, and pennates into araphid and raphid based on the presence of the raphe, a structure along the frustule that enables gliding movements on a surface. However, this classification does not reflect diatom evolution (See Figure 1) (Medlin, 2016). Thanks to their silica frustule, diatom fossil records can be traced back, and with the combined use of molecular and fossil data, diatom important evolutionary transitions can be dated. Diatoms are clearly found in
fossil records from 180 million years ago (Ma) in the Jurassic period, but some report earlier rise of diatoms after the Permian-Triassic mass extinction crisis (250Ma) (Sims et al., 2006; Sorhannus, 2007). First diatoms were radial centric diatoms, and diversified during the Cretaceous. Multipolar centric, then araphid Pennate diatoms appear during the Cretaceous; the apparition of raphid penates is dated from the late Cretaceous. This group diversified rapidly and nowadays, raphid pennate species outnumber araphid pennate and centric species combined (Nakov et al., 2018).

Diatoms are today a major lineage of photosynthetic eukaryotes, and they are considered to be the most species-rich clade of algae (Kooistra et al., 2007; Mann and Vanormelingen, 2013; Nakov et al., 2018). They come in a diversity of size and beautiful shapes (Fig. 2) and occupy a wide range of ecological niches in the contemporary ocean (Malviya et al., 2016).

## LIFE IN THE OCEAN: CHARACTERISTICS OF THE DIATOM ENVIRONMENT

Phytoplankton growth is essentially controlled by light and nutrient availability (nitrogen, phosphorous, silica and iron, but also vitamin B12), plus eventually by CO₂ concentration and temperature. Diatoms have colonized very different environmental niches and can display different adaptive strategies.

## DIATOMS IN THE WATER COLUMN: PLANKTONIC LIFESTYLE

Diatoms can have a planktonic lifestyle, i.e. drifting in fresh or salted water. I will describe here the global framework to understand phytoplankton life in the oceans. In the water column, light and nutrients usually show opposite gradients with depth: light is high at the surface and decreases exponentially with depth; nutrients are low at the surface and high at depth (Fig. 3A). The “euphotic zone” defines the upper ocean layer that is illuminated (usually defined as the water layer where light intensity is higher than 1% of the surface light intensity) and where photosynthesis can take place.

In parallel, temperature is usually high at the surface and decreases at depth (Fig. 3). In a water column with no or little turbulence, the nutrients in the photic zone will be depleted
by phytoplankton growth and exported to the depth due to dead cells sinking. Phytoplankton growth is then nutrient-limited, and this can results in a "deep chlorophyll maximum" (DCM), with a higher phytoplankton concentration at a depth reflecting a trade off between light and nutrient requirements (Fig. 3A) (Mann and Lazier, 2006). This situation is common all year long in Tropical regions and in late summer in temperate waters. In temperate regions, seasonal variation will change the vertical structure of the water column. In winter, the upper oceanic layer is mixed to an important depth due to high turbulence (wind at the surface, storms) and nutrients from deeper waters are brought back to the surface (Fig. 3B). In this upper mixed layer, the physico-chemical parameters of the water are homogenous (temperature, density, nutrient concentration)(Mann and Lazier, 2006). However, light is in general lower in winter, for astronomic reasons (shorter photoperiod and lower sun angles, so a lower light intensity) and if the mixed layer is deeper than the photic zone, cells are periodically taken to the darker deep waters out of the photic zone. Phytoplankton growth is therefore considered to be light-limited. In spring, light limitation is alleviated either by the increase in day length and light intensity and/or by the mixed layer thinning that allows cells to stay longer in the photic zone. Nutrients are still high for the winter mixing events, and usually sustain spring blooms. Later in the season, increase in temperature will cause thermal stratification of the water column. In the photic zone, nutrients are quickly used for phytoplankton growth; some of it may be recycled with in photic zone though grazers ad cell lysis, but sinking of dead cells (eventually in aggregates and fecal pellets from grazers) will export nutrients to the deep and deplete the upper layer. Low mixing prevents the replenishment of nutrient stock at the surface and subsurface or deep chlorophyll maximum can appear in these conditions (Fig 3B) (Mann and Lazier, 2006). Finally, in autumn, the deepening of the mixer layer can sustain fall bloom in some regions.

Some marine environments show specificities that differentiate them from the global model presented above. In upwelling region, wind-induced currents cause nutrient-rich water from the deep to arise to the surface (Mann and Lazier, 2006). Coastal regions are
further influenced by tides and freshwater run-off from land, which also show seasonal
rhythms (equinox tides, river discharge increase in spring).

<figure><img src="image_4.png" /><figcaption>Figure 3. Environmental context for phytoplankton life in the water column. A. Biotic and abiotic parameters across the water column in stratified waters. Light is high at the surface, while nutrients are high at depth, leading to the formation of t maximum of chlorophyll and phytoplankton concentration at depth (DCM). Drawn by (Pierella Karlusich et al., 2020) with data from Bermuda Atlantic Time Series station. B. Seasonal variation of the chlorophyll profile with depth. In January, deep mixing brings enough nutrients to the photic zone to sustain phytoplankton growth. From winter to spring, increased stratification leads to the formation of a DCM, explained in A. Drawn from (Sauzède et al., 2015), data from the same station as A.</figcaption></figure>

Several studies showed that species succession during the season is remarkably conserved from year to year (Caracciolo et al., 2021; Longobardi et al., 2022). Both papers concluded that time of year per se, i.e. astronomical parameters such as photoperiod (Longobardi et al., 2022) and light intensity (Caracciolo et al., 2021), are primary determinants of phytoplankton community succession. Internal factors such as biotic interactions also control self-organization patterns and annual oscillations (Caracciolo et al., 2021).
Diatoms are usually considered to thrive in high nutrient, highly mixed environments (i.e. "spring-like" waters) (Margalef, 1978). Indeed, they often dominate the spring bloom and they are thought to be well adapted to these highly variable environments (especially to light variations with mixing (Lavaud et al., 2007; Lavaud and Lepetit, 2013)). However, diatoms are found in all marine regions, including tropical regions (Malviya et al., 2016). A recent review highlighted possible adaptation strategies to stratified water columns in diatoms (Kemp and Villareal, 2018). Indeed, diatom can be a major contributor of the DCM in stratified waters by adapting to low light levels ("shade flora" (Kemp et al., 2000; Goldman and McGillicuddy, 2003)), but also by regulating their buoyancy to perform vertical migration to acquire nutrient at depth and return to the near surface at a speed up to 7m/h (Moore and Villareal, 1996; Singler and Villareal, 2005). In addition, diatoms are able to store nutrients, including nitrate, in their vacuoles, allowing them to store nutrients during sporadic mixing events and to divide at higher rates than their competitors in nutrient scarce conditions (Dortch et al., 1984; Behrenfeld et al., 2021). Diatoms also possess a complete urea cycle, allowing them to rapidly recover from nutrient starvation (Allen et al., 2011). Finally, diatom-diazotroph association fuel diatom and phytoplankton community with newly fixed nitrogen, and alleviates nitrogen limitation (Singler and Villareal, 2005; Foster et al., 2011).

DIATOMS IN MARINE SEDIMENTS: BENTHIC ORGANISMS

The benthic zone is the bottom layer of a water body, including the sediment surface and its first centimeters. More generally, we call "benthic" any algae growing on a substrate (organic or not). This can include very different environments, as tidal mudflats, surf beaches, coral reefs, coastal sediments or organic surfaces such as sea turtle shells or macroalgae blades. Benthic diatoms play major roles in this ecosystem: they are primary producers at the base of the trophic chain, they mediate exchanges at the sediment-water interface, and they have major roles in stabilizing the sediments through the secretion of Extracellular Polymeric Substances (EPS) (Underwood and Kromkamp, 1999; Middelburg et al., 2000; Serôdio and Catarino, 2000; Underwood and Paterson, 2003).
Sediments are very peculiar environments compared to the water column. Light is attenuated by the sediments at the µm scale (compared to meters in the water column), and there are also steep gradients of temperature, nutrients, salinity, and if the microorganism population is dense, gradients of O₂, Dissolved Inorganic Carbon (DIC) and pH (Fig.4). In addition, temporal variation of these parameters with tides and day/night cycles can be extreme in the tidal zones (Cartaxana et al., 2016b; Cartaxana et al., 2016a; Marques da Silva et al., 2017).

Diatoms have colonized benthic environments and can dominate the microbenthos (i.e benthic microalgal community) in some regions, for example intertidal mudflats. Some species can shift between benthic and planktonic lifestyle and are called tychoplanktonic. Most of the diatom benthic species are pennate and are extremely diverse (An et al., 2020). Two types of benthic diatoms can be distinguished: epipsammic diatoms, that live attached to sediment grain and are not motile (or very little), as opposed to epipelic diatoms, free-living forms that can migrate daily over several millimeter of sediment. These are mainly raphid pennates that can glide on sediments thanks to their raphe.

<figure><img src="image_5.png" /><figcaption>Figure 4. Example of vertical profile of oxygen and light in two types of sediments (A. mud and B. coarse sand) containing diatoms (A. epipelic, B epipelic and epipsammic species). The photic zone in the muddy sediments is very thin, and was densely populated. In the sandy sediment, the photic zone is deeper, and we observe an increase in light intensity in the first 0.1mm at the near surface due to high scattering and low absorption From (Cartaxana et al., 2016b)</figcaption></figure>

Epipelic diatoms show impressive migratory behaviors to cope with these environmental constraints. They usually dominate in fine, soft sediments and silt habitat, and they are
able to move toward light, or away from it, to find optimal light conditions to perform photosynthesis (Fig. 4A). In dense photosynthetic communities, photosynthesis activity depletes inorganic carbon stocks and micromigration can allow alternate between high light/low DIC surface and low light/high DIC subsurface environment, with continuous replacements of the cells at the surface (Consalvey et al., 2004; Vieira et al., 2016; Marques da Silva et al., 2017). Deeper migratory patterns (at the mm scale) are also controlled by diurnal light rhythms (up during the light phase) and tidal rhythms (up during low tide, burial before tidal inundation). These rhythms can persist in continuous condition, highlighting an endogenous control of the timing of this behavior (Palmer and Round, 1967; Barnett et al., 2020). However, in the dark diatoms have to adapt their metabolism: they can grow heterotrophically (Lewin and Lewin, 1960) and in anoxic sediments, they can also perform nitrate respiration based on intracellular stored nitrates (Kamp et al., 2011; Merz et al., 2021).

In sediments with larger grain size, epipellic and epipsammic species can co-exist (Fig. 4B). Light penetrates deeper (see also next paragraph) and epipsammic diatoms (that do not migrate) adapt to high light through physiological photoprotective mechanisms, as opposed to “behavioral” mechanisms such as downward migration of epipellic diatoms (Cartaxana et al., 2011; Barnett et al., 2015). When photosynthetic organisms density is high, as occurs in biofilms, the community can self-organize in layers (niche differentiation). Benthic diatoms usually compose the uppermost layer, with green algae and cyanobacteria underneath. Anoxygenic phototrophs such as green and purple sulfur bacteria can be found below the cyanobacteria, where O₂ is depleted and H₂S concentrations are high (Stal et al., 1985; Stal et al., 2019).
<figure><img src="image_6.png" /><figcaption>Figure 5. A. Ice associated algal communities B. Time of phytoplankton blooms in the Arctic ecosystem at different latitudes, from (Daase et al., 2021)</figcaption></figure>

One other peculiar environments colonized by diatoms is sea ice (Fig5). Diatoms (especially pennate diatoms) dominate the sea ice community and can represent up to 90% of the algal cell abundance (Arrigo, 2016). Sea ice algae are usually found in the bottom 20cm of the ice sheet, where sufficient nutrients are available from the seawater below. Most common diatom genera in the bottom sea-ice community are *Nitzschia*, *Fragilariopsis* and *Navicula*. The sea ice interior is characterized by low nutrient and high salinity in brine pockets and channels, and thus less favorable to algal growth. Pennate diatom aggregates and chains of the centric diatom *Melosira* can also colonize the underside of the sea ice. The polar algae communities are subjected to drastic environmental changes along the seasons. In autumn, the ice starts to form when temperature fall below -1.76°C and traps protists into the sea ice (Różańska et al., 2008). Some algae can form cyst and resting stages/spores to survive the adverse conditions within sea ice. During winter, the polar night and the thickness of sea ice limit light availability for alga, and there is a selection mechanisms causing the overall diversity of the ice community to decrease. The return of the sun and lengthening of the days trigger the sea ice algal bloom, when low-light adapted algae within the sea-ice start to proliferate (Rozanska et al., 2009). At this time of the year, sea-ice bloom is often the sole source of carbon for heterotrophs. Further temperature increase causes the ice to melt, and algae are released to the water column (Leu et al., 2015). Different diatom types are thought to be adapted to the different stages of seasonal progression (Croteau et al., 2021). As in sediments, some pennate diatoms are motile and adjust their position in the sea ice to
optimize their growth (Aumack et al., 2014). Diatoms use EPS secretion and ice-binding
proteins to protect themselves from low temperature and adhere and migrate on ice.

THE PECULIARITY OF LIGHT IN AQUATIC ENVIRONEMENTS

The light field is defined both by light intensity and light spectral quality. In terrestrial environments, the light field depends on cloud cover, time of day, time of year, latitude and shade (notably by other photosynthetic organisms). In aquatic environments, an additional dimension has to be considered: depth.

LIGHT IN THE WATERCOLUMN

As already mentioned before, light intensity decreases with depth in the water column. The main physical processes determining the underwater light field are absorption and scattering, which will both reduce the light intensity. Major light-absorbing components in aquatic environments are water, Colored Dissolved Organic Matter (CDOM) and particulate matter as inanimate particles and phytoplankton (Kirk, 2011).

Water molecules themselves absorb light and can contribute to an important part of light absorption coefficient of natural waters (it can represent 68% of the absorption of Photosynthetically Active Radiations, PAR). The absorption spectra of pure water (Fig. 6) show high absorption in the long wavelength part of the light spectrum (above 500 nm). Salts present in seawater have negligible effect on light attenuation. CDOM, also called gilvin or yellow substance, derives from the decomposition of organic matter. In coastal regions, CDOM is a major component of light absorption coefficient, while it is less concentrated in clear oceanic waters. Source of CDOM is terrestrial organic matter (or resuspended organic matter from sediments). CDOM absorbs mainly in the blue, with an exponential decrease towards longer wavelengths (Fig. 6b and c).
<figure><img src="image_7.png" /><figcaption>Figure 6. Examples of underwater light fields in different aquatic environments and decomposition of their absorption spectra. A In clear oceanic waters (Pacific Ocean, station ALOHA), light penetrate deep (see spectra at 100m) and is blue enriched due to absorption by water. B, in coastal waters from the Baltic sea, absorption by CDOM and particles (gilvin+tripton) is higher, which leads low light penetration, but with higher penetration of green light. C. Finally, in the Groot Moost lake, a peat lake in the Netherlands, turbidity is so high that light is attenuated in the first meter, and the light filed is red-enriched. (Figure from Stomp et al., 2007)).</figcaption></figure>

Inanimate particulate matter is also called tripton or non-algal particles. It is composed of mineral particles and dead phytoplankton cells, and its absorption spectrum globally has the same shape as CDOM (Fig. 6). Photosynthetic pigments from living phytoplankton cells absorb light at specific wavelengths. Total phytoplankton absorption (both in shape and value) depends on the species composition of the phytoplankton community, their concentration, but also on their physiological state (cell size and intracellular pigment concentration). In general, phytoplankton absorption spectra show a strong absorption in blue and second peak in red corresponding to the chlorophylls (Kirk, 2011).

Scattering is the deviation of a photon from its original path following interaction with a particle or a molecule (Kirk, 2011). In aquatic environments, interaction with particles will increase the optical path for photons, and thus increase their probability of being absorbed; this will intensify the vertical attenuation of light. Scattering in natural waters
is roughly neutral towards wavelengths, with a slight attenuation towards long wavelengths.

Marine waters can be classified based on their optical properties (Morel et al., 2006). Case I waters are oceanic waters, where optical properties are directly linked to phytoplankton abundance. Coastal waters with limited terrigenous inputs can also be considered as case I waters. At low phytoplankton concentration, i.e. in oligotrophic waters, absorption of long wavelengths by water will result in blue-enriched light field at depth (Fig. 6a). Case II waters are waters for which resuspended sediments and dissolved organic matter (for example from river discharge) are important contributors to the optical properties. Absorption in the blue is very high in these waters, resulting in green-shifted light fields. (Fig. 6b). In some extremely turbid ecosystems, absorption of blue and red can lead to a red-enriched light field (Fig. 6c).

## LIGHT IN SEDIMENTS AND BIOFILMS

Less is known on light attenuation in marine sediments compared to light in the water column. The main protagonists involved in light attenuation are mineral particles (sand, silt, mud) and organic matter (living particulate and dissolved) (Cartaxana et al., 2016b). Here, water absorption is negligible compared to the other actors, but water is an important component as wet sediments scatter light less than dry sediments and light will penetrate deeper in wet sand (Kühl and Jorgensen, 1994). Both absorption and scattering are more intense than in the water column due to the high density of sediment particles and algae living in it, and the depth of the “photic zone” is limited to a few mm (Fig. 4).

The size of the mineral particles is crucial in sediment light attenuation (Kühl et al., 1994), through effect on scattering. Indeed, small particle size scatter light more compared to coarse grains. Light attenuation by sand is higher in the blue (maximum around 450nm), and decreases toward longer wavelengths. Presence of organic matter and microalgae will increase light attenuation in the blue range, with also a peak around 680nm due to chlorophyll absorption (Haardt H, Nielsen Gae, 1980; Kühl and Jorgensen, 1994). In biofilms, i.e. very dense communities of microorganisms, the layers of different organisms will also change the attenuation coefficient with depth due to their different pigment
composition (Fig. 7A) (Kühl and Jorgensen, 1992), for example chlorophylls and bacteriochlorophylls.

<figure><img src="image_8.png" /><figcaption>Figure 7. Light field sediments and in ice. A. Light under 1mm of coarse-grained costal sediment occupied by diatoms and cyanobacteria. Note that light is more absorbed at 680nm due the chlorophyll a. From (Lassen et al., 1992). B. Light under sea-ice. Maximum light penetration in the ice is at 460nm. Data from (Lund-Hansen et al., 2020).</figcaption></figure>

Intense scattering in sediments and in biofilms will also create a "subsurface irradiance maxima". Indeed, scalar irradiance, i.e. light intensity integrated from all directions, is higher than at surface in the first 0.5 millimeter below the sediments and can reach up to 280% of the surface (downwelling) irradiance in fine sand (Kühl et al., 1994). One other difference with the water column environment is the exposure to high UV light. If both the sea surface and the sediment surface are exposed to direct sunlight, the cells are diluted in the full volume of the mixer layer in the ocean, while they are concentrated to the surface and first mm in the sediment. Algae in sediment are thus exposed to much higher doses of UV light (Garcia-Pichel and Bebout, 1996).

As water, ice absorbs mostly in red part of the light spectra, resulting in blue-enriched light fields (Fig. 7B)(Ehn and Mundy, 2013). However, in the ice algae layer, the algae density is so high that absorption in the blue becomes important, and the light field can shift towards green. In addition, ice is a highly scattering media resulting in reduced light intensity.
LIFE CYCLE

All diatom species sequenced so far are thought to be diploid in their vegetative states and divide mitotically during the major part of their life cycle (Falciatore et al., 2020).

Diatoms have a “bloom and burst” life cycle, with periods of intense replication when the environmental conditions are favorable. The bloom's dynamics are not fully understood, and the bloom's demise can be due to different parameters (not mutually exclusive), for example nutrient depletion, grazing pressure, or pathogen attack (Behrenfeld et al., 2021).

Some diatoms can form resting stages to survive unfavorable environmental conditions and resume growth when better conditions are met. The cues that trigger resting stage formation are not clear for all species, but high cell density and nitrogen limitation are suspected (McQuoid and Hobson, 1996; Pelusi et al., 2020). Two types of resting stages can exist: resting spores, which are morphologically different from vegetative growing cells, and resting cells. Both have a reduced metabolism, and resting cells are able to successfully germinate after centuries (*Skeletonema marinoi* (Harnstrom et al., 2011)) and even millennia (*Chaetoceros muelleri* (Sanyal et al., 2021)) in sediments. Resting stages are important in diatom species succession, and resting stage stocks accumulated in sediments can form a reservoir of viable cells, seeding blooms in spring in coastal regions (Eilertsen et al., 1995)

Some aspects of the diatom life cycles are shaped by their silica cell wall, the frustule. The frustule is made of two pieces that fit together like a Petri dish and encapsulate the cell. The two pieces, also called theca, are slightly different in size: the larger one (like the Petri dish lid) is called the epitheca, and the smaller one is the hypotheca. Due to this rigid silica cell wall, daughter cells following mitotic division are different (see Fig. 8). One inherited the epitheca and will result in cell size comparable with the mother cell, while the cell that inherited the small hypotheca will form a new hypotheca inside the maternally inherited one (using it as a template), resulting in smaller cell size. This results in size reduction with mitotic division, which can end with cell death if the cell size becomes too small to
allow survival. In some diatoms, sexual reproduction is the only stage where size
restoration can occur (Bilcke et al., 2022) (Fig. 8).

<figure><img src="image_9.png" /><figcaption>Figure 8. Diatom life cycle. Clockwise from the red diatom: during mitotic division, each daughter cell inherits a theca and build a new hypotheca. This results in cell size reduction. When critical cell size is reached, or in response to other cues, sexual reproduction can occur, restoring cell size (yellow cells: example of the sexual reproduction in *Thalassiosira weissflogii*, redrawn from (Bilcke et al., 2022)).</figcaption></figure>

Sexual reproduction can be triggered when critical cell size is reached, but also in
response to environmental cues (nutrient, salinity, pheromones, light) (Amato, 2010;
Godhe et al., 2014; Moeys et al., 2016; Basu et al., 2017; Moore et al., 2017). Compared to
the diploid part of diatom life cycle, which can last years, sexual reproduction is short and
has rarely been observed in nature. Centric and pennate species show differences in their
sexual cycles (which are arguments to classify them too). In general, centric species are
homothallic, i.e. clonal cells can generate large oogonia or small, motile spermatocytes
(Fig. 8). Pennate are usually heterothallic, i.e. two mating type are required to perform
sexual reproduction (Bilcke et al., 2022). In raphid pennate, compatible mating type cells
must pair to form the gametangia that will undergo meiosis and gametogenesis. Here, the
two gametes mating types are indistinguishable (isogamy). In all diatom groups,
fertilization results in a large cell called auxospore that will resume mitotic division and
vegetative life phase. The model diatom *P. tricornutum* and *T. pseudonana* are presumed
to predominantly reproduce asexually in the lab and are rarely used to study sexual
reproduction (one exception is the study by Moore et al., 2017). Other species have arisen
as model species for sexual reproduction, mainly the pennate species *Seminavis robusta*
and *Pseudo-nitzschia multistriata*.

Numerous studies reported the effect of light on the diatom life cycle. Blue light is especially efficient in controlling mitotic cell cycle progression in *P. tricornutum* through blue light photoreceptor-mediated control of cyclin expression (Huysman et al., 2013). Resting spore germination is sensitive to photoperiod (Eilertsen et al., 1995) and light quality, especially to blue light (*Skeletonema costatum*, *Thalassiosira minima*, and *Chaetoceros sp*, (Shikata et al., 2009)) and blue and red light through photosynthesis activity in *Leptocylindrus danicus* (Shikata et al., 2011). Finally, sexual reproduction is controlled by red and far-red light in the diatom *Stephanopyxis palmeria* in a reversible manner (induction by red light, repression by far-red light applied after red light) (Ren et al., 2008). Red light also triggers sexual reproduction in the pennate diatom *Haslea ostreicola* (Mouget et al., 2009), but in *Seminavis robusta*, blue light is the more efficient waveband (Bilcke et al., 2021). Light intensity also matters: in *Seminavis robusta*, higher reproduction rate occurs at the lowest intensity tested (4µmol photons/m²/s, Bilcke et al., 2021) and in *Thalassiosira weissflogii* a period of darkness and/or low light are necessary to trigger sexual reproduction (Armbrust, 1990).

PHOTOSYNTHESIS

As mentioned earlier, there is a common agreement that diatom chloroplasts arose from
red algae. Most of the components of oxygenic photosynthesis are conserved in diatoms,
but they also possess unique features linked to their complex evolutionary history
(Büchel et al., 2022).

The chloroplasts possess a network of membranes containing the photosynthetic
apparatus: the thylakoids. Photosynthesis takes place in 2 phases: the “light reactions”,
where light energy is harvested and transformed into chemical energy, and the
“biochemical reactions”, when this chemical energy is used to fix carbon (CO2). The
photosynthetic chain in the thylakoids carries out the light reactions (Fig. 9). Protein complexes are embedded in the thylakoid membrane: photosystems (PS) I and II, cytochrome b6f complexes and ATP synthase. Photosystems are composed of core reaction centers containing a special pair of chlorophyll a and Light harvesting complexes (LHC) containing pigments. Light is absorbed by pigments in the LHC or directly by the 2 chlorophylls in the reaction centers and light energy transferred to the reaction centers where charge separation occurs. This fuels a flow of electrons in the photosynthetic chain from water ($H_2O$, with production of $O_2$) to NADP+, producing reducing power in the form of NADPH. The electron flow also allows the build-up a proton gradient across the thylakoid membrane. The return of $H+$ from the lumen to the stroma fuels the production of ATP by the ATP synthase.

Compared to the green algae and plants, diatom-specificity contributing to the light reactions are notably the spatial organization of protein complexes in the plastid and the protein and pigment content (Büchel et al., 2022). Diatom chloroplasts are embedded in 4 membranes, and their thylakoids are organized in stacks of 3 thylakoids (Fig. 9), and the different complexes are heterogeneously localized: PSI are rather in the stroma-facing membranes, while PSII are enriched in the core of the stacked membranes, facing other thylakoid membranes. ATP synthase is in the outer, stroma-facing membranes. This organization can change with light conditions (see also below).

The light harvesting complexes of diatoms differ from other eukaryotes (Büchel et al., 2022). LHC proteins can be grouped in 3 main families: LHCF, the major LHC group; LHCR, associated with PSI and related to red alga LCFs; and LHCX, that are of supposed green alga origin and are involved in photoprotection. In addition to chlorophyll a, LHCs bind chlorophyll c and fucoxanthins. This carotenoid pigment is specific for Ochrophyta and gives these algae their brownish color.
<figure><img src="image_10.png" /><figcaption>Figure 9. The photosynthetic light reactions in diatoms (the dotted line illustrates the linear flow of electron across the photosynthetic complexes).</figcaption></figure>

Excess light energy can damage photosystems, especially the PSII and therefore photosynthetic organism evolved sophisticated photoprotection mechanisms. In addition to efficient PSII repair systems, diatoms can also quickly adjust PSII activity and dissipate excess energy: this can be detected *in vivo* by a decrease in PSII fluorescence, and is called Non Photochemical Quenching (NPQ) (Lepetit et al., 2022). At least part of the NPQ capacity is related to a xanthophyll cycle involving the conversion of diadinoxanthin to diatoxanthin pigment under high light. This xanthophyll cycle is different from the one found in plants, both in terms of pigments (plants use violaxanthin, antheraxanthin and zeaxanthin pigments) and enzymatic reactions (two-step process in plants and only one in diatom). LHCX antenna are also involved in NPQ regulation, but the exact mechanisms is unknown (Bailleul et al., 2010).

Acclimation to both light intensity and light quality occurs in diatoms. High light acclimation is depicted by high maximal photosynthetic rate with high photoprotective mechanisms (high NPQ) (Schellenberger Costa et al., 2013). In *P. tricornutum*, blue wavelengths are sufficient to trigger this phenomenon, through aureochromes and cryptochromes photoreceptors (Juhas et al., 2014; Mann et al., 2017) which control expression of light-harvesting proteins. Few studies suggest that chromatic acclimation, i.e. acclimation to light quality, can also occurs and identified specific photosynthetic
hallmarks for cells grown in red light (Fujita and Ohki, 2004; Herbstová et al., 2015; Herbstová et al., 2017). In *P. tricornutum* and *Nitzschia closterium*, cells grown in red light showed a fluorescence spectra peak in FR light, associated with the presence of a specialized LHC in *P. tricornutum*, Lhcf15 (for more details, see chapter 3). A reorganization of the photosynthetic complexes in the thylakoid membrane was also described (Bína et al., 2016), with a stronger spatial heterogeneity. Schellenberg Costa and coauthors (Schellenberger Costa et al., 2013) compared chloroplast membrane proteins of *P. tricornutum* cells grown under blue or red light and identified proteins specific for the different light conditions (10 upregulated in blue light and linked to high light acclimation, 4 upregulated in red light including Lhcf15). Transcriptomic responses to changes in light quality, although at high intensity, go in the same direction, with an induction of many LHCF and LHCR genes in green and red, while blue light induced LHCX genes and genes involved PSII repair (Valle et al., 2014).

Fixation of CO₂ by Rubisco in the Calvin-Benson-Bassham cycle is the biochemical phase of photosynthesis. However, in aquatic environments, CO₂ concentrations are very low, as CO₂ dissolved in H₂CO₂, which dissociates in HCO₃⁻. The majority Dissolved Inorganic Carbon (DIC) in water is in the form of HCO₃⁻, which cannot cross membranes, while CO₂ can. Diatoms have developed efficient Carbon Concentration Mechanisms (CCM) to cope with the low CO₂ concentration in water (Kroth and Matsuda, 2022). These algae possess several HCO₃⁻-transporters, some of which can pump HCO₃⁻ inside the cell, and carbonic anhydrases converting HCO₃⁻ to CO₂ and vice-versa in the different compartments. Some of these enzymes are thought to be secreted and convert HCO₃⁻ to CO₂ at the cell surface. CO₂ passively crosses the plasma membrane and is converted to HCO₃⁻ in the cytosol, to which we should add HCO₃⁻ actively pumped at the plasma membrane. HCO₃⁻ is then transported inside the chloroplast by unidentified transporters, to the stroma where it is transformed back into CO₂ in the vicinity of RubisCO. This CCM is an important mechanism for diatoms in the wild, as mining surface seawater metagenomes and metatranscriptomes revealed an important number of carbonic anhydrase transcripts (Pierella Karlusich et al., 2021). Some studies showed that some diatoms possess genes homologs to genes involved in the C4 mechanisms from plant, but functional studies of these genes did not confirm the occurrence of this process (Haimovich-Dayan et al., 2013),
and mining environmental data showed that these genes rarely co-occur, limiting the
possible involvement of C4 in the environment (Pierella Karlusich et al., 2021).

DIATOM MOTILITY

Pennate raphid diatoms are able to glide on surface at high speed (up to 25µm/s). Not much is known about the mechanisms of diatom motility, but it involves secretion of mucilage through the raphe (Poulsen et al., 2022). Current models involve the secretion of mucilage via the Golgi and transport of vesicles to the plasma membrane. Extracellular mucilage is linked via transmembrane proteins to myosin motors, themselves connected to actin filaments. Myosin action builds a force along the actin filaments, which pushes the cell in the opposite direction.

Diatom movement is tightly controlled, especially by light. We already underlined diatom
movements in sediments. In natural communities, blue light is the most efficient
waveband to trigger positive phototaxy (Barnett et al., 2020). Numerous studies on
monocultures in laboratory conditions showed the effects of different wavebands: cells
are in general more attracted by blue light (*Craticula sp*, *Pinnularia sp*, *Nitzschia sp* (Cohn
and Weitzell, 1996); *Nitzschia communis* (Nultsch, 1971), *Nitzschia perminuta* (McLachlan
et al., 2009)) and repelled by red (Nultsch, 1971; Cohn and Weitzell, 1996), but this is
intensity- and strain-dependent. In addition, changes in illumination can change the
direction or the speed of the movement. Finally, the tip of the cell seems to be the site of
light perception (Cohn et al., 1999), and further signaling events involve calcium release
from intracellular socks (McLachlan et al., 2012).

LIGHT SENSING

Light is an essential source of information for organisms on Earth. All organisms
(photosynthetic or not) possess specialized proteins called photoreceptors that perceive
light and mediate the response to this environmental cue. These proteins usually bind a
pigment (also called chromophore) that enables them to absorb light at specific
wavelengths (Möglich et al., 2010). Upon light absorption, changes in the chromophore conformation impact protein conformation, changing its activity (protein-protein interaction, phosphorylation or other enzymatic activity for example). This will activate specific signaling cascade(s) leading to the biological responses.

A photoreceptor typically displays several protein domains or modules that mediate different parts of the photoreceptors function: light-sensing domains and signal transduction effector domains. Different classes of light-sensing domains are known, with different chromophores and light sensing properties. Three types of blue-light photoreceptors bind flavin chromophores: photoreceptors with a Light-Oxygen Voltage (LOV) domain, which bind FMN, sensors of Blue Light Using FAD (BLUF) and cryptochromes, which bind FAD. Other photoreceptors detect blue light via the 4-hydroxycinnamic chromophore and are called Photoactive Yellow Protein (PYP), while the UVB photoreceptor UVR8 perceives light through absorption by tryptophan residues. Rhodopsins perceive blue and green lights through a retinal chromophore, and finally phytochromes perceive red and far-red lights with a bilin chromophore. The combination of the same type of photosensing domain with different effector domains leads to protein with different functions. As example (see also below), the LOV-based photosensors are found in combination with a variety of effector domains across the three kingdoms of life (Glantz et al., 2016).

Land plants and green algae possess examples of 5 of the above mentioned photoreceptor types (no homologs of PYP or BLUF have been reported in Viridiplantae so far). Cryptochromes, which are photoreceptors known across all eukaryotes, are found in land plant and green algae. UVR8 photosensors are thought to be restricted to the green lineage. LOV domains exist in combination with a serine/threonine kinase domain (phototropins), F-box domain and Keltch repeat (Zeitlupe family), PAS domain (PAS/LOV in land plants), Histidine kinase domain (LOV-HK from Ostreoccocus and other mameliophyceae), but also in combination with other photosensory domain like phytochrome photosensory module (neochromes ferns and mosses). Different types of rhodopsins also exist in the green algae *Chlamydomonas reinhardtii*, while the phytochromes photoreceptors are conserved in land plants but not in all green algae.
Exploration of diatom genomes has led to the discovery of many putative photoreceptors in these organisms (Jaubert et al., 2017). Some are classical photoreceptors known from bacteria, land plant or animals, while others show unique domain combinations. Photoreceptor types in diatoms include cryptochromes, LOV-based aureochromes, rhodopsins and phytochromes. No homologs of the plant phototropins have been identified in diatoms. Although first identified in *P. tricornutum* (Bowler et al., 2008; Djouani-Tahri et al., 2011), close examination of the putative LOV-HK homologs in diatoms showed that they lack the critical amino acids for interaction with the chromophore. Diatoms have genes with similarity to UVR8, but these lack the critical tryptophans involved in light sensing and are thus not considered as UVB photoreceptors (Fernández et al., 2016).

In the following section, I will describe the current knowledge on the diatom photoreceptors, with a particular focus on phytochromes, which represent the main topic of my PhD. Some paragraphs (Cryptochromes, Aureochromes and Rhodopsins section, marked with *) have been extracted from a recently review published in the "Molecular Life of Diatom book", that I co-authored (Jaubert et al, 2022). The full chapter is available at the end of this manuscript, in the annex section. Additional information on new diatom photoreceptors and their distribution are described in the result session (Chapter 1).

# DIATOM PHOTORECEPTORS

## Aureochromes *

The aureochromes (AUREOs) are unique blue light photoreceptors that possess both an FMN-binding light-oxygen-and-voltage (LOV) domain (Crosson et al. 2003), and a bZIP domain typical for bZIP transcription factors (TFs) (Dröge-Laser et al. 2018) (Fig. 1). The blue light phototropin photoreceptors of green algae and plants also possess two LOV domains, but utilize a serine/threonine (Ser/Thr) kinase domain for signal transduction instead (Christie 2007). AUREOs were originally discovered in the xanthophyte alga *Vaucheria frigida* in 2007 (Takahashi et al. 2007). Their name refers to "aurum" (Latin for gold), because of the golden-brownish colour of most stramenopiles. Takahashi et al. (2007) identified two orthologs in *V. frigida*, VfAUREO1, and VfAUREO2. Using an RNA
interference approach to silence these two genes individually, they demonstrated that both AUREOs are involved in the regulation of photomorphic responses. Meanwhile, AUREOs have been identified in other stramenopiles (Ishikawa et al. 2009; Jungandreas et al. 2014) and in a raphidophyte (Ji et al. 2017), but not in non-photosynthetic oomycetes (Kroth et al. 2017). AUREOs are not present in red algae, which are considered to represent the endosymbiotic ancestors of stramenopile plastids (Archibald 2015). This indicates that the ancestral AUREO gene with its unique combination of LOV and bZIP domains may have been provided either by the putative host cell of the secondary endosymbiosis event, or that it evolved very early within the stramenopiles, possibly via domain shuffling (Di Roberto and Peisajovich 2014).

Stramenopile AUREOs differ in their structures. While AUREO1 proteins possess the typical LOV domain, AUREO2 proteins have a mutation within the LOV domain, which prevents non-covalent binding of flavin needed for light absorbance in the blue range. The reason is a steric hindrance from a methionine residue within the binding cavity (Banerjee et al. 2016a). AUREO2, therefore, is not a real photoreceptor, but could still be involved in light regulation, e.g. by forming a dimer with a light-sensing AUREO1 protein. In reciprocal experiments, genetic modification of AUREO2 from *P. tricornutum*-restored flavin binding (Serif 2017), while introducing a point mutation at the same site in PtAUREO1a, led to loss of flavin binding (Banerjee et al. 2016a). Based on this distinction and on phylogenetic analyses, aureochromes in diatoms and other organisms have been classified as either AUREO1-type (a/b/c etc.) or as AUREO2 (Schellenberger Costa et al. 2013b). The algae studied so far all possess one AUREO2 protein and one or more AUREO1 isoforms (Table 2).

Blue light absorption, both in aureochromes and phototropins, causes the formation of an adduct between the flavin and a nearby cysteine within a few microseconds, starting the signalling cascade (Toyooka et al. 2011; Kerruth et al. 2014). The domain topology of AUREOs is inverted as compared to most other characterized LOV proteins because the sensory domain of AUREOs is at the C-terminus of the receptor. The Jα helix of the AUREO-LOV domain allosterically regulates the fold of the N-terminally flanking A'α helix (Herman and Kottke 2015). Subsequent unfolding of the A'α helix exposes a high affinity dimerization site and enables the formation of the light state dimer of LOV (Herman et al.
2013; Herman and Kottke 2015; Heintz and Schlichting 2016). Indeed, in full-length AUREO, the Jα helix plays a crucial role in the formation of the light state of the receptor (Banerjee et al. 2016b). The activation of the LOV domain results in a loss of the helical secondary structure of the bZIP domain (Banerjee et al. 2016b), indicating that there is a direct communication between the two domains. If a DNA binding site is available, the helical fold of the bZIP domain is increased by light (Banerjee et al. 2016b), resulting in rigidification of the domain (Heintz and Schlichting 2016). In a complementary approach, Tian and colleagues showed the importance of a previously overlooked Cα helix in promoting the conformational protein changes (Tian et al. 2020). bZIP domains have a general tendency to dimerize, and are only capable of binding DNA as dimers (Tateyama et al. 2018). For some AUREOs, dimerization and DNA binding have been shown to be induced by blue light (Hisatomi and Furuya 2015; Banerjee et al. 2016b; Nakatani and Hisatomi 2018; Nakajima et al. 2021). PtAUREO1a occurs in the dark as a dimer/higher oligomer (Banerjee et al. 2016b) or as a monomer in equilibrium with a dimer (Heintz and Schlichting 2016). Light induces the dimerization of LOV domains and the association of the monomers (Kobayashi et al. 2020; Goett-Zink et al. 2020), which is the rate-limiting step in the process of DNA binding (Akiyama et al. 2016). Based on these properties, AUREOs could potentially be used as an optogenetic tool, for instance, to increase protein stability under blue light conditions (Hepp et al. 2020) or directly as a light-driven gene switch.

AUREO function has been extensively characterized in *P. tricornutum*, especially for PtAUREO1a that acts as a key regulator of the diatom cell cycle (Table 1; Chapter "Cellular Hallmarks and Regulation of the Diatom Cell Cycle"). Evidence from silenced and knockout lines of *P. tricornutum* further indicates that PtAUREO1a and PtAUREO1b might be involved in regulation of photoacclimation (Table 1; Schellenberger Costa et al. 2013b; Serif et al. 2017; Mann et al. 2017; Madhuri et al. 2019). PtAUREO1c might be a high light sensor in vivo because it recovers faster and is much less sensitive to light than PtAUREO1a (Bannister et al. 2019). Recent studies indicate that AUREOs may have a large impact on the cells. Changes in transcriptomes in response to a shift from red to blue light were analyzed in wild-type *P. tricornutum* cells, in PtAUREO1a knockout and in PtAUREO1a complemented lines (Mann et al. 2020). Wild-type cells react within minutes
by up- or down-regulating 75% of the genes, while this massive change in gene expression
is mostly inhibited in PtAUREO1a knockout strains (Mann et al. 2020). PtAUREO1a,
therefore, must have a specific function in cellular regulation that cannot be
complemented by other AUREOs. This raises the question of how a single photoreceptor
can affect such a large number of genes. Possibly, a cascade of TF transduces the initial
response of PtAUREO1a to blue light. This is supported by findings that the transcript
abundance of a large number of diatom TFs and photoreceptors (Rayko et al. 2010) is
strongly and rapidly affected by blue light in wild-type cells, but not in PtAUREO1a
knockout mutants (Mann et al. 2020). When common gene expression patterns are
analyzed, the aureochromes are placed in different clusters (Ait-Mohamed et al. 2020),
supporting the idea that AUREO1 isoforms may have different roles. There also is some
evidence that AUREOs might be involved in regulation of the diel cycle and the expression
of some AUREOs follows a different diurnal pattern (Banerjee et al. 2016b). The diurnal
rhythmic expression of PtAUREO1a and 1c can still be detected when the cells are kept in
the dark, while PtAUREO1b expression appears to be light activated, and PtAUREO2
oscillates only weakly throughout the day. The recently discovered diatom clock
component RITMO1/PtbHLH1a (Annunziata et al. 2019) is strongly induced by blue light
in wild-type cells, but not in the PtAUREO1a mutant, indicating that PtAUREO1a might be
involved in triggering the diatom clock in response to blue light.

Cryptochrome/Photolyase Family *

The cryptochrome/photolyase family proteins (CPFs) are widespread blue light-
absorbing flavoproteins with similar primary sequences, but very diverse functions. Most
CPFs non-covalently bind a flavin adenine dinucleotide (FAD) cofactor as a specific
prosthetic chromophore. Other chromophores such as 5,10-methenyltetrahydrofolic acid
(MTHF), 8-hydroxy-5-deazaflavin, or flavin-mononucleotide (FMN) may also be
associated with some CPFs as light antennae (Essen et al. 2017). CPF members have a
characteristic conserved photolyase-related (PHR) domain, but amino and carboxy
terminal extensions are highly variable in both length and primary sequence. Photolyases
(PL) are blue light-activated enzymes repairing UV-induced DNA lesions, such as
cyclobutane pyrimidine dimer (CPD PL) or (6-4) pyrimidine-pyrimidone photoproducts. The cryptochromes (CRY) of plants and some insects do not have any DNA repair activity, but are either blue light photoreceptors or light-independent components of the central circadian oscillator (Chaves et al. 2011). The activation of light-dependent CRY has been well studied. The photoperception process starts with a very fast photoinduced reduction of FAD. This redox- and light-dependent change at the core of the protein leads to conformational changes, allowing specific interactions with other protein partners. Globally, both light- dependent and -independent CRY, are involved in transcriptional regulation, respectively, by inhibiting transcriptional repressors or by inactivating transcription.

Phylogenetic relationships in the CPF family helped to identify five major super- classes (sc) which do not necessarily converge functionally. For instance, sc1 includes (6-4) PLs but also light-dependent animal CRYs and light-independent CRYs involved in the transcription/translation feedback loop of the circadian clock. Class I and III CPD photolyases group also together phylogenetically with proteins with different functions such as light dependent plant photoreceptor CRY and plant- like photoreceptor CRY in sc3, but are separate from class II CPD photolyases which are all found in sc4. The last two super-classes (sc2 and sc5) include all CRY-DASH (named after Drosophila, Arabidopsis, Synechocystis, Human) and the proteobacterial PL/CRYs (Fortunato et al. 2015; Ozturk 2017). A surprising result from genome sequence analyses was that diatoms do not possess canonical plant CRY photoreceptors, even though blue light is preponderant in the ocean and CRY regulate so many physiological processes in plants. However, further genomic and functional investigations in diatoms revealed novel CPF variants, including the animal-like CPF1 and plant-like CRYs (CryP) (see below) and several DASH CRYs (Table 2). The biological function of Cry DASH is not yet clearly defined in diatoms or indeed in other organisms, but several studies suggest that they might have a single-stranded DNA CPD PL activity, a signalling role, or be involved in the regulation of metabolism, consistent with their organellar localization (Kleine et al. 2003; Froehlich et al. 2010).

Only two diatom CPFs have been characterized in detail to date: CPF1 and CryP of P. tricornutum (Fig. 1b). Initially discovered in diatoms, these proteins are extensively
represented in metatranscriptomic data derived from marine environments (Coesel et al. 2021). CPF1 belongs to sc1, is localized in the nucleus, and has (6-4) PL DNA repair activity both in vitro (Coesel et al. 2009) and in vivo (De Riso et al. 2009). However, like plant CRY, CPF1 regulates the transcription of several genes acting in pathways modulating photoprotection, cell division, nutrient assimilation, etc., under blue light (Coesel et al. 2009), so is likely to be also a photoreceptor. CPF1 is also able to bind mammalian CLOCK protein in heterologous cells where it partially represses the CLOCK/BMAL1 heterodimer (Coesel et al. 2009) in the positive loop of the circadian clock (Kume et al. 1999).

<figure><img src="image_11.png" /></figure>

Figure 10 (Figure 1 from the book chapter). Sensing of light cues in diatoms. (a) Photoreceptor classes identified in diatom genomes. Cryptochrome-Photolyase, Aureochrome, Rhodopsin, and Phytochrome families are positioned schematically according to the range of wavelengths they usually absorb. The nature of the chromophores is indicated below the respective photoreceptor boxes, with question marks indicating putative chromophore. (b) Summary of characterized photoreceptors from *P. tricornutum* (Pt) and *T. pseudonana* (Tp). Domain architecture, chromophore, absorption spectrum, and regulated processes are indicated for: cryptochrome photolyase 1 (PtCPF1) (Coesel et al. 2009); plant-like Cry (PtCryP) (Juhas et al. 2014); aureochromes PtAUREO1a (Heintz and Schlichting 2016) and PtAUREO1c (Bannister et al. 2019) and diatom phytochromes (PtDPH and TpDPH) (Fortunato et al. 2016). Domains involved in the light sensing are indicated in blue or red

CryP belongs to the group of Plant-like photoreceptors CRY (sc3). Recombinant P. tricornutum CryP, produced in E. coli, binds FAD and MTHF chromophores (Juhas et al. 2014). Moreover, FAD photoreduction, a common mechanism in CPFs that bind FAD, is also active in vitro, suggesting that activation of CryP is light- dependent. Originally, CryP
was identified as a blue light regulator of light harvesting proteins directly involved in
photoprotection (LHCX) (Juhas et al. 2014). However, subsequent analyses of
transcriptional responses to illumination by blue light after prolonged dark, revealed that
gene expression was already deregulated in the dark in CryP knockdown lines compared
to wild type (König et al. 2017). Therefore, despite the presence of chromophores, CryP
may not be a major blue light photoreceptor but rather a component involved in the global
modulation of transcription, requiring other blue light photoreceptors to signal the light
responses.

Rhodopsins *

These light-sensing integral membrane proteins found in Archaea, bacteria, and
eukaryotes share a topology of seven transmembrane alpha helices within which a retinal
chromophore is covalently bound (Ernst et al. 2014). Rhodopsins exhibit a wide variety
of spectral tuning in the blue-green part of the spectrum depending on the nature of a few
influential amino acids interacting with the retinal (Man 2003; Ernst et al. 2014).
Rhodopsins function as light-driven ion pumps, ion channels, or light sensors (Grote et al.
2014). The discovery that H+-pump rhodopsin converts light into ATP has challenged the
assumed monopoly of photosynthesis as a phototrophy-enabling mechanism (Béjà et al.
2001; Finkel et al. 2013). Recently, a distinct group of microbial rhodopsins, the
heliorhodopsins, has been identified after analysing environmental genomic samples
(Pushkarev et al. 2018). Heliorhodopsins do not have the capacity for light-triggered ion
transport but they do have a long photocycle, suggesting that they could act as signalling
photoreceptors (Pushkarev et al. 2018). H+-pump rhodopsins and heliorhodopsins are
present in bacteria, Archaea, and algae and are highly represented in environmental
genomic data (Pushkarev et al. 2018; Coesel et al. 2021).

Diatom rhodopsin-like sequences falling into the H+-pump group, based on conservation
of key amino acids and phylogeny, were first identified in the transcriptome of Pseudo-
nitzschia granii (Marchetti et al. 2012), then in those of other species (Marchetti et al.
2015), and in the genome of F. cylindrus (Mock et al. 2017) (Table 2). Because P. granii
rhodopsin-like transcripts are highly abundant in low iron conditions, it has been
hypothesized that this proteorhodopsin-like protein could be involved in energy
production under conditions of iron deficiency that affect photosynthesis (Marchetti et al.
2012). It is noteworthy that a gene homologous to heliorhodopsin has been identified in
the genomes of *P. tricornutum* and other pennate diatoms, but not in centric ones
(Pushkarev et al. 2018) (Table 2). G-protein-coupled receptor rhodopsin-like genes,
homologs of receptors that transduce a wide range of stimuli including light, hormones,
volatile molecules, glycoproteins, nucleotides, and chemokines in eukaryotes (Costanzi et
al. 2009), have also been identified in various diatom genomes (Port et al. 2013), with
evidence of expansion of this gene family in some species (Osuna-Cruz et al. 2020).
However, information about the spectral and functional properties of these and other
diatom rhodopsins is still lacking, so their function as light sensors remains to be
established.

PHYTOCHROMES

Phytochromes are known for more than 60 years as master regulators of plant
photomorphogenesis, where they controls germination, de-etiolation, shade avoidance,
stomatal development, entrainment of the circadian clock, and flowering (non exhaustive
list!) in response to light and to the ratio of red to far-red wavelengths (Franklin and Quail,
2010). These photoreceptors were subsequently found in other organisms,
photosynthetic or not: prokaryotes such as cyanobacteria and anoxygenic photosynthetic
bacteria, but also non-photosynthetic bacteria; photosynthetic eukaryotes such as
glaucophyta and prasinophyte (primary endosymbiosis) or cryptophytes and
ochrophytes (secondary endosymbiosis), and heterotrophic eukaryotes such as Fungi
(Rockwell and Lagarias, 2020). Evolutionary, phytochromes are thought to originate from
bacteria (Rockwell and Lagarias, 2020), and transferred to eukaryotes at least twice
independently. Indeed, phytochrome phylogenetic studies showed that plant
phytochromes form a large and robust clade, sister to prasinophyte, glaucophyte and
cryptophyte phytochromes ("Eukaryotic phytochrome type 1" in Fig 11) while Fungi and
Stramenopile phytochromes form an independent clade ("Eukaryotic phytochrome type
2" in Fig 11). Phytochromes from the eukaryotic type 1 clade have been proposed to arise
from endosymbiotic gene transfer from cyanobacteria (Kooß and Lamparter, 2017), but
other studies resolve cyanobacterial phytochrome as an independent clade within
bacterial phytochromes (Duanmu et al., 2014; Li et al., 2015a). Within the eukaryotic type 1 clade, glaucophyte phytochromes are the first to diverge, followed by cryptophyte phytochromes; however, cryptophyte algae are not part of the archaeplastidia themselves (Burki et al., 2020). This incongruence between phytochrome tree and species tree has not been resolved. In the other eukaryotic branch, Fungi and Stramenopile phytochromes are sister clades, but the origin of these phytochrome has not been determined. Horizontal gene transfer, deletion but also duplication and diversification have shaped the evolutionary history of this photoreceptor (Rockwell and Lagarias, 2020).

<figure><img src="image_12.png" /><figcaption>Figure 11: Phylogenetic analysis phytochrome photoreceptor (based on the GAF and PHY proteins domains). Eukarotic Type 1 phytochrome contains phytochromes from Viridiplantae and Cryptophyte, while eukaryotic type 2 contains Fungi and Stramenopile phytochromes. These includes phytochromes from brown algae (Phaeophyceae), Diatom (Pennate and Centrales) and brown algal viruses. From (Fortunato et al., 2016).</figcaption></figure>

These photoreceptors share a common structure, with a conserved Photosensory Core Module (PCM) in the first part of the molecule, composed of PAS, GAF and PHY domains, and a more variable Output Module (OM)(Fig. 12). The chromophore is a linear tetrapyrrole derived from heme (bilin), which is covalently bound to a cysteine either in the PAS or in the GAF domain (Fig. 12). Different chromophore can be used: biliverdin (in
proteobacterial and fungal phytochromes for examples (Bhoo et al., 2001; Karniol et al., 2005; Blumenstein et al., 2005) or more reduced bilins, such as phytochromobilin (plants) or phycocyanobilin (streptophyte algae and cyanobacteria (Rockwell et al., 2014; Hughes et al., 1999). Heme oxygenase is the enzyme necessary to produce BV from heme, while other enzymes (Bilin Reductases) are required to produce the other chromophores. Light absorption triggers photoisomerization of the bilin around a double bound, from E to Z conformation or vice versa. Phytochromes exist in two states, a red absorbing form Pr with the chromophore in the Z conformation (in canonical plant phytochromes), and a far-red absorbing form Pfr, bound to chromophore in E conformation(Rüdiger and Thümmler, 1994). Absorption of red (R) light light by Pr will trigger the Pr→Pfr reaction, and absorption of far-red (FR) by Pfr will trigger to Pfr→Pr reaction (Fig. 12A). In the absence of light, some phytochromes will revert to its most stable form: this is called the dark or thermal reversion (Mancinelli, 1994). The dark state can be Pr (in canonical phytochromes) or Pfr (bathyphytochromes). The equilibrium between the two forms depends on the light environment, presence and binding of partners, temperature and dark reversion rate. In addition, phytochromes work as dimers *in vivo* (Klose et al., 2015; Brockmann et al., 1987)

## Plant phytochromes

Land plant possess variable copy number of phytochrome genes, which regulate different aspects of morphogenesis both in redundant and specific manners (Franklin and Quail, 2010; Legris et al., 2019). Seed plants possess 3 conserved types of phytochromes: PhyA, PhyB and PhyC. In the model plant *Arabidopsis thaliana*, subsequent duplication of PhyB gave rise to PhyE, found in all Angiosperms, and PhyD, which is specific for Brassicaceae (Mathews and McBreen, 2008). Phytochrome responses were first classified as Type I, represented by PhyA responses, and Type II. Type I responses are also called Very Low Fluence Responses, and function as a light- but not wavelength-sensitive response (Shinomura et al., 1996). Indeed, PhyA rapidly converts from Pr to Pfr upon light illumination of any wavelength, but is very unstable and degraded; this response is irreversible. PhyA also mediates High Irradiance Responses, which require long exposure to high light intensities. Type II responses, or Low Fluence Responses, are the classical phytochrome responses: they are induced by a pulse of red light, show R/FR reversibility,
sensitivity to the R/FR ratio and follow the reciprocity law, i.e the amplitude of response observed is a function of the total amount of photon received (Time of exposure*Light intensity) (Mancinelli, 1994). In addition, the rate of photoconversion and dark reversion is affected by ambient temperature, and phytochromes are involved in temperature sensing in plants (Jung et al., 2016; Legris et al., 2016). Structural and biochemical differences (thermal reversion) could explain the differences between phytochrome responses (Legris et al., 2019; Burgie et al., 2021).

Plant phytochromes bind phytochromoblin as chromophore with a cysteine in their GAF domain, and their output module is composed of 2 PAS domains and a Histidine-Kinase Related domain (HRKD) (Fig. 12B). Upon light exposure, phytochrome is shuttled to the nucleus and interacts with Phytochrome Interacting Factors (PIF), which are bHLH transcription factors. PIF will be phosphorylated, ubiquitinated and addressed to the proteasome for degradation (See Fig. 12 for more details on the phytochrome signaling cascade in plants). Whether all the phytochrome HRKD has a serine/threonine phosphorylation activity is debated (Boylan and Quail, 1996; Shin et al., 2016; Ni et al., 2017; Li et al., 2022).
<figure><img src="image_13.png" /><figcaption>Figure 12. Signaling mechanisms in phytochromes. A. Phytochromes can bind PCB, PφB or BV as chromophore. Illumintaiton triggers isomerisation around the C15-C16 double bound (see molecular structure on the right). Phytochromes exist in two conformation, Pr and Pfr. As they work as dimers, 3 different dimers type are found in equilibrum under illumination. Eventual thermal reversion might revert phytochrome to this most stable form. B. Different phytochrome structure found in Eukaryotic group 1 (first column), Eukaryotic group2 and the most common architechture for Bacteriophytochrome (Second column). Dotted contour indicates that this domain may not be present. C. Signalisation cascade. A. Simplified phytochrome signalisation cascade in plants. Upon illumination, phytochromes will be shuttled to the nucleus, where they interact with PIF transcription factors and inhibit them. PIFs will be phosphorylated and degraded or sequestered in the cytosol. In addition, phytochromes can interact with E3 ubiquitin ligase complexes such as the COP1/SPA1 complex. This will prevent the E3 ligase complex to address transcription factors to the proteasome for degradation. *Legend continues on next page*</figcaption></figure>
B. In bacteria, phytochromes such as RpbphP4 function as two-component systems, and will phosphorylate a response regulator domain (left). However, it has been shown recently that some phytochromes can work as phophatase (Deinococcus radiodurans DraBphP, middle) (Multamäki et al., 2021). Other bacteriophytochromes possess a REC domain fused at their C-terminal, suggesting that the signalisation cascade thus requires an additional phosphorelay (Hpt protein, right). C. In Fungi, the signaling cascade seem to begin as a two component system. In the dark YdpA, an Hpt protein, is phosphorylated and keeps Sska (REC) inactive. Upon illumination, YpdA dephosphorylates (YpdA interacts with FphA, but the light-dependency is unknown). SskA will interact with SskB, the first component of a MAPKinase cascade, which ends with the phosphorylation of SakA. SakA will be shuttled to the nucleus, where it will activates transcription factors (AtfA). FphA has also been shown to interact with LreB and VeA which are invlovled in blue-light signalling, and to have an impact on histone acetylation, potentially regulating gene expression.

## Bacterial phytochromes

Bacterial phytochromes show variations both in the PCM and in the OM (Auldridge and Forest, 2011) (Fig. 11). Most bacterial phytochromes have the PAS-GAF-PHY architecture, but some lack the PAS, like Cph2 from the cyanobacteria Synechocystis, and others only have the GAF domain (Cyanobacteriochromes, CBCR) (Fig 12). Bacteria phytochromes use biliverdin as a chromophore, bound at the N-terminal extremity (Bhoo et al., 2001), while cyanobacteria phytochromes and CBCR use phycocyanobilin bound to the GAF domain (Yeh et al., 1997; Hughes et al., 1997). Additional variation of the PCM includes the presence of a PYP (Photoactive Yellow Protein) upstream of the PCM; this additional domain binds another chromophore (p-hydrocinnamic acid) and absorbs blue light, conferring new light-sensing properties to these chimeric photoreceptors. The Output module is a typical two-component system domain, with histidine kinase eventually followed by a response regulator domain. This system is a phosphorelay-signaling cascade, with ATP-dependent phosphorylation of a Histidine residue in the HK domain; the phosphate group is then transferred to an aspartate in a response regulator (RR) protein (Yeh et al., 1997; Giraud et al., 2005) (Fig. 12). Phosphorylation of the RR affects its activity (usually, DNA binding and transcription regulation). Bacteriophytochrome genes are often part of operon containing also heme oxygenase for BV production and response regulator genes, and sometimes also the phytochromes-regulated (Bhoo et al., 2001; Giraud et al., 2002; Lamparter et al., 2002). Other output architecture exists, but their signaling mechanisms have not been elucidated.
Different aspects of bacterial physiology can be regulated by phytochromes. In cyanobacteria, one major response is complementary chromatic adaptation (CCA) (reviewed in Wiltbank and Kehoe, 2019). CCA is the process by which cyanobacteria remodel their light harvesting machinery (phycobilisomes) to optimize their light harvesting capacity according to the incoming light quality (Grossman, 2003). Different types of CCA exist, regulated by CBCR (Kehoe and Grossman, 1996). CBCR can have very different absorption spectra, enabling the detection of green, teal, yellow and red light for example (Wiltbank and Kehoe, 2019). A special case of CCA, far-red light photoacclimation, is regulated by a phytochrome with red/far-red absorption spectra (Gan et al., 2014; Gan and Bryant, 2015).

Anoxygenic photosynthetic bacteria also use phytochrome to regulate their photosynthetic apparatus (Giraud et al., 2002; Giraud et al., 2005; Jaubert et al., 2007). Phytochromes control both photosystem and light harvesting complexes synthesis, in response to far-red light in *Rhodopseudomonas palustris* and *Bradyrhizobium sp*. In some *Rps. palustris* strains, some phytochromes have lost their light-sensing abilities, but mediate the response to redox conditions (Vuillet et al., 2007).

In non-photosynthetic bacteria, the physiological role of phytochromes is less known. In *Agrobacterium fabrum*, recent studies showed that 2 phytochromes (Agp1 and Agp2) are involved in the control of conjugation and plant infection (Bai et al., 2016; Xue et al., 2021). In the non-photosynthetic bacteria *Deinococcus radiodurans*, phytochrome regulates the synthesis of carotenoid pigment that protects the bacteria during growth under high light (Davis et al., 1999). Recently, bacteriophytochrome from a deep-sea bacterium *Croceicoccus marinus* was proposed to promote growth in infra-red (940nm) by modulating the cell's metabolism (Liu et al., 2021).

## Fungal phytochromes

Phytochromes seem to be widespread in Fungi, at least in Ascomycota (Schumacher, 2017). Their structure is close to Bacteriophytochromes (Fig. 12B), but with a long N-terminus extension, and they bind biliverdin as chromophore. In the model *Aspergillus nidulans*, the phytochrome FphA has a red/far-red absorption spectra and represses
sexual reproduction in red light (Blumenstein et al., 2005). It is localized in the cytoplasm
and signals through interaction with the phosphotransfer protein YdpA, as a two-
component phosphorylation relay, followed by the HOG (high osmolarity glycerol) MAP
kinase cascade and activation of transcription factors (Yu et al., 2016) (Fig. 12C). FphA
also interacts with blue-light photoreceptors and with chromatin modification complexes
(Purschwitz et al., 2009; Hedtke et al., 2015). Recombinant FphA shows no dark reversion
*in vitro* (Consiglieri et al., 2019), but it was suggested that FphA is also a temperature
sensor in Fungi (Yu et al., 2019).

Algal phytochromes

Phytochromes have been found in many marine algae, and their characterization is gaining interest in recent years (Duanmu et al., 2014; Rockwell et al., 2014). Duanmu et al. (2014) showed that phytochrome from the green algae *Micromonas pusilla* (marine prasinophyte) is shuttled to the nucleus during the light period, suggesting conserved mechanism with land plant phytochromes. However, this phytochrome exhibits light-induced auto-phosphorylation activity (probably histidine kinase). This phytochrome and other prasinophyte phytochromes (*Dolichomastix tenuilepis*, *Tetraselmis astigmatica*, *N pyriformis*, *Prasinoderma coloniale*) bind phycocyanobilin and show variation of their absorption spectra, from red/far-red (648/734nm, *T.astigmatica*) to absorption spectra shifted towards shorter wavelength, i.e., orange/far-red (586-614/690-718 nm) (Fig. 13) (Rockwell et al., 2014). This shift towards shorter wavelength is supposed to be an adaptation of phytochrome absorption spectra towards light abundant in the aquatic environment, i.e. spectral tuning (fig 13). Other algal phytochromes have been spectrally characterized: 2 phytochromes from glaucophytes, showing peculiar absorption properties and photocycle with blue/red absorption shifts; *Ectocarpus siliculosus* phytochrome 1 (brown algae), which binds phytochromobilin and has a red/green photocycle. These variations in absorption spectra are once again discussed in the scope of spectral tuning, considering that phytochromes in aquatic environments are tuned towards blue and green wavelengths (Fig. 13) (Rockwell et al., 2014))
<figure><img src="image_14.png" /><figcaption>Figure 13. Algal phytochromes absorption spectra. PtDPH absorption spectra in compared to different algal phytochromes from (Rockwell et al., 2014). PtDPH Pfr is clearly the most FR-shifted phytochrome. Cpar and Gwit, *Cyanophora paradoxa* and *Guillardia theta* (Glaucophytes); Dten and Pcol, *Dolichomastix tenuilepis* and *Prasinoderma coloniale* (Prasinophytes); Esil, *Ectocarpus siliculosus* (Phaophyceae) B. Hypotheses on DPH activation in the environment</figcaption></figure>

Contrary to algal phytochromes presented above, phytochromes from the 2 marine diatom models *Thalassiosira pseudonana* (Tp) and *Phaeodactylum tricornutum* (Pt) bind biliverdin and have a red/far-red absorption spectra (686/764 nm for TpDPH, 700/750nm for PtDPH, Fig. 13), shifted towards longer wavelength compared to plant phytochromes (Fortunato et al., 2016). Other DPHs found from an initial exploration of available genome information are structurally similar to TpDPH and PtDPH and phylogenetically close to bacterial and fungal phytochromes. Surprisingly, pennate and centric phytochromes do not belong to the same clade, but pennate phytochromes are sister to phaeophyceae, while centric phytochromes are sister to phaeophyceae+pennate phytochromes. Interestingly, giant brown algal viruses carry phytochrome genes that branch close to these groups, suggesting that horizontal gene transfer could have contributed to the evolution of diatom phytochromes. In addition, we can note that not all diatoms possess phytochromes, but that centric diatoms do have it in one copy while pennate phytochromes can be present in several copies.

Before I started by PhD, functional studies in *P. tricornutum* using knockout (KO) lines have shown that diatom phytochrome (DPH) regulates gene expression in response to far-red light (Fortunato et al., 2016). The far-red light induced genes (80 genes) are of unknown function (55%) or involved in transcription and signaling (25%). DPH does not seem to be involved in red light gene regulation in *P. tricornutum*. DPH KOs do not show alteration of physiology so far, so the consequences of DPH activation by far-red light are
unknown. *In vitro* both Pt and Tp DPH show no dark reversion, but PtDPH has been shown to auto-phosphorylate in response to FR light, suggesting a phosphorylation cascade as in two-component systems. No homologs of PIFs have been found in diatom genomes, suggesting different phytochrome signalization pathway.

Hypotheses have been made concerning the light that could be perceived by DPH in the marine environment (Fortunato et al., 2016). DPH could sense direct solar R and FR light, but this would be limited to the first meters of the water column, which leads to the idea that DPH could be a surface sensor. Alternatively, DPH could sense R and FR photons from fluorescence of photosynthetic organisms; in that case, DPH could perceive changes in its own chloroplast photosynthetic activity, or light from surrounding photosynthetic organisms. This could bring information on cell density in an algal bloom for example. However, it was underlined that below the first meters the R/FR light ratio is rather constant in the marine environment, and thus DPH activity is probably not based on sensing this waveband ratio.

## MODEL SPECIES AND GENETIC TOOLS

### THE GROWING NUMBER OF DIATOM GENOMES, TRANSCRIPTOMES AND META-DATA

The first diatom genome to be sequenced was the one from the cosmopolitan centric diatom *T. pseudonana* (Armbrust et al., 2004). Whole-genome sequence of a second model diatom, *P. tricornutum* soon followed (Bowler et al., 2008). These species were also chosen because they were amenable to genetic transformation and show fast growth under laboratory conditions. Both their genomes are relatively small (32.1 and 27.4 Mb for *T. pseudonana* and *P. tricornutum*, respectively) with 11776 and 12177 coding genes, respectively. Some peculiarities of diatoms already arose from the analysis of their genomes: the presence of red and green algal genes, an important number of bacterial genes from horizontal gene transfer, in addition to species-specific genes. Even today,
with way more genomes available, the fraction of genes not shared between centric and pennate species is very high (40%), highlighting the diversity and rapid evolution of diatoms (Tirichine et al., 2017; Mock et al., 2022).

<table><caption>Table 1. Sequenced genomes of centric diatoms</caption><thead><tr><td>Centric species</td><td>Reference and interest</td></tr></thead><tbody><tr><td>Thalassiosira pseudonana</td><td>(Armbrust et al., 2004), first diatom sequenced</td></tr><tr><td>Thalassiosira oceanica</td><td>(Lommer et al., 2012), adaptation to iron-poor waters</td></tr><tr><td>Cyclotella cryptica</td><td>(Traller et al., 2016; Roberts et al., 2020), biofuel production</td></tr><tr><td>Skeletonema costatum</td><td>(Ogura et al., 2018; Sorokina et al., 2022), harmful algal blooms and abundant coastal diatom</td></tr><tr><td>Skeletonema marinoi</td><td>(Johansson et al., 2019), chain forming and abundant coastal diatom</td></tr><tr><td>Chaetoceros tenuissimus</td><td>(Hongo et al., 2021), widely distributed blooming diatom, very small</td></tr><tr><td>Attheya, Chaetoceros calcitrans, C. decipiens, C. muelleri, C. neogracile, Skeletonema dorhrnii, S. marinoi, S. menzelii, Thalassiosira guillardii, T. nordenskioeldii, T. weissflogii</td><td>(Nelson et al., 2021), large sequencing project to get insights from diverse microalgae, with a focus on the role of viruses in algal evolution</td></tr><tr><td>Chaetoceros gracilis</td><td>(Kumazawa et al., 2022)</td></tr></tbody></table>

A lot of other diatom species were sequenced to study specific aspects of diatom life and explore diatom diversity, with different interests (evolutionary, ecological, biotechnologies), summarized in Table 1 and 2 below.

We can also mention ongoing projects such as the Chaetoceros sequencing project "Most Abundant Diatom Genus in the World's Ocean" and the 100 Diatom genomes project with very different diatom species.

Databases and tools have started to emerge for the molecular study of diatoms. One of the first databases was the Diatom Portal, gathering gene expression microarray data from *T. pseudonana* and *P. tricornutum* (Ashworth et al., 2016). A new platform gathering expression data from RNASeq experiments is being developed (Diatomicbase, Villar et al,
unpublished but available at the end of this manuscript). In another project, PhaeoNet integrated Pt RNASeq data to provide co-expression analyses (Ait-Mohamed et al., 2020). Comparative genomics platform PLAZA Diatom is available to compare gene families across diatom genomes (Osuna-Cruz et al., 2020).

<table><caption>Table 2 Sequenced genomes of pennate diatoms</caption><thead><tr><td>Pennate species</td><td>Reference and interest</td></tr></thead><tbody><tr><td><i>Phaeodactylum tricorntutum</i></td><td>(Bowler et al., 2008), first pennate genome; model species</td></tr><tr><td><i>Pseudo-nitzschia multiseries</i></td><td>harmful algal blooms</td></tr><tr><td><i>Pseudo-nitzschia multistriata</i></td><td>(Basu et al., 2017), sexual reproduction</td></tr><tr><td><i>Fragilariopsis cylindrus</i></td><td>(Mock et al., 2017), adaptation of polar regions</td></tr><tr><td><i>Fistulifera solaris</i></td><td>(Tanaka et al., 2015), biofuel production</td></tr><tr><td><i>Seminavis robusta</i></td><td>(Osuna-Cruz et al., 2020), first benthic diatom</td></tr><tr><td><i>Synedra acus</i> (<i>Fragilaria radians</i>)</td><td>(Galachyants et al., 2015), araphid pennate, freshwater</td></tr><tr><td><i>Nitzschia inconspicua</i></td><td>(Oliver et al., 2021), biofuels and bioproducts; suitable for large scale aquaculture</td></tr><tr><td><i>Amphora coffeaeformis</i>, <i>Cylindrotheca fusiformis</i>, <i>Halamphora</i>, sp (2 strains), <i>Navicula incerta</i>, <i>Navicula pelliculosa</i></td><td>(Nelson et al., 2021), large sequencing project to get insights from diverse microalgae, with a focus on the role of viruses in algal evolution</td></tr><tr><td><i>Haslea ostrearia</i></td><td>(Gabed et al., 2022) Unique marenine pigment production, epiphytic diatom</td></tr></tbody></table>

Genomic data are growing, but transcriptome resources are still the reference for many diatom species. The MMETSP (Marine Microbial Eukaryote Transcriptome Sequencing Project) gathered data from 411 microalgal strains including 92 diatoms (Keeling et al., 2014). This valuable resource was re-assembled three times (Johnson et al., 2019; Guita Niang et al., 2020; Van Vlierberghe et al., 2021) to produce robust and clean (from bacteria contamination notably) references with the latest bioinformatics tools.
Lately, a new type of genetic data arose: environmental genomics. The massive sequencing of complete communities allowed the assembly of expressed transcripts forming atlas of expressed genes, and metagenomes-assembled genomes and sequencing of single cells allowed the assembly of genomes of uncultured organisms. However, the annotation and use of these data depends on our understanding of lab strains (ie genomes and MMETSP transcriptomes). The main contributions to these types of data (for eukaryotes) are the Tara expeditions: *Tara* Oceans and *Tara* Oceans Arctic circle, for which data are already available (Carradec et al., 2018; Seeleuthner et al., 2018; Delmont et al., 2020); data from other expeditions (Pacific Ocean, Mediterranean Sea) are on their way. Sampling was performed with a repetitive protocol that allows sample comparison, and a lot of other data are available (HPLC, physico-chemical parameters of the water column (optics, nutrient, carbon chemistry) to link genetics data to environmental conditions. Another project recently made public is the Sea of Change project (Martin et al., 2021).

## MOLECULAR MODEL SPECIES

As the first diatoms whose genome was sequenced, *T. pseudonana* and *P. tricornutum* are the most studied diatoms in molecular studies. Both species are small (10µm) marine diatoms, isolated from coastal waters, close to estuaries. Some studies suggest that *T. pseudonana* is in fact a freshwater species that re-colonized marine environments (Alverson et al., 2011). In the ocean, the *Thalassiosira* genera is one of the most abundant diatom genera (Malviya et al., 2016). *P. tricornutum* is not an abundant species in the open ocean, but several strains were isolated from coasts around the world (De Martino et al., 2007), suggesting a widespread distribution. This species has the unique feature of existing 3 distinct morphotypes, oval, elongated and triradiate, and is able to shift from one morphotype to the other depending on the strain and the environmental conditions. *P. tricornutum* is though be a benthic diatom in its oval form, and planktonic in its triradiate and elongated forms. Contrary to other diatoms, *P. tricornutum* does not have a strict requirement for silica and its cell wall is only poorly silicified. Sequencing of ecotypes in both *T. pseudonana* and *P. tricornutum* (Koester et al., 2007; Rastogi et al., 2019) unveiled their dominant reproduction mechanisms (sexual for *T. pseudonana*,
asexual for *P. tricornutum*) through population genetics, and allow the study of gene polymorphisms, eventually linked to adaptation to specific environmental conditions.

GENETIC ENGINEERING

Genetic transformation has been set up for the model species *T. pseudonana* and *P. tricornutum* (Apt et al., 1996; Falciatore et al., 1999; Poulsen et al., 2006), and also for other sequenced diatom species (Moosburner et al., 2022). The first successful method to transform diatom was by biolistic transformation, with the delivery of a DNA vector on gold or tungsten beads. The insertion of the vector into diatom genomes is stable but random, and often in multiple copies, and this can lead to the random disruption of nuclear genes and variable levels of expression of the transgene (George et al., 2020). DNA can also be introduced by electroporation (Falciatore et al., 2020).

Lately, the use of a yeast derived sequence allowed the design of an episome for the delivery of genetic material by bacterial conjugation (Karas et al., 2015). In most of the cases, the plasmid is not integrated into the genome but maintained as an episome. Some variations of expression within a clone are observed, but expression is comparable between clones (George et al., 2020). Antibiotic selection has to be maintained to keep the episome, and without selection, the episome is loss. As consequence, this method can be used for transient expression of deleterious transgenes and a more homogenous expression between independent transgenic lines.

First modulations of gene in diatoms by reverse genetic approaches were done in diatoms by gene overexpression. Several plasmids were designed with different promoters to allow high expression of the transgene in the cell (reviewed in (Falciatore et al., 2020). Down regulation of a gene was also achieved by gene silencing (De Riso et al., 2009; Shrestha and Hildebrand, 2015). However, gene silencing is not fully controlled in diatoms and does not always determine clear reduction of expression of the targeted genes (De Riso et al., 2009; Shrestha and Hildebrand, 2015). Genome editing techniques were recently developed, first with TALEN (Transcription activation Like Effector Nuclease)(Daboussi et al., 2014), and more recently with the CRISPR-Cas9 method
(Hopes et al., 2016; Nymark et al., 2016). Genome editing is achieved by introducing in diatom a vector containing the Cas9-coding gene and guide RNA genes, that have to be expressed in the cell, either by biolistic transformation and vector integration or as episome (Hopes et al., 2016; Nymark et al., 2016; Sharma et al., 2018). The Cas9 can also be directly delivered as ribonucleoproteins and introduced into the cell by biolistics, resulting in DNA-free edition of the genome (Serif et al., 2018).

Finally, plastid transformation has been reported in *P. tricornutum* (Materna et al., 2009; Xie et al., 2014), which should enable the study of chloroplast-encoded genes.

OBJECTIVE OF THE THESIS

Light regulates different aspects of diatom life. Accordingly, diatoms possess different
photoreceptors that are involved in light regulation, including a plethora of blue light
photoreceptors, but also red/far red light photoreceptors, the diatom phytochromes
(DPH).

At the time I joined this project, new information on some photoreceptors identified in the genomes of some diatom species began to emerge (Jaubert et al., 2017). Particularly, the discovery of a red and far-red light photoreceptor in diatom model species, using far-red light as an active signal, was particularly surprising, as aquatic environments are less transparent than the atmosphere to these radiations (Fortunato et al., 2016). However, a complete information on the full possible photoreceptors repertoire in diatoms was still lacking, which seemed particularly relevant given the enormous diversity of diatoms and their complex evolution. Therefore during my PhD training, I addressed several questions, and their results are described in different chapters of this thesis:

*What is the repertoire of diatom (and more largely Ochrophyta) photoreceptors? Can we link this repertoire to the algae environment? What is the evolutionary origin of these photoreceptors, considering the complex history of Ochrophyta? And more specifically, what is the origin of diatom phytochromes?*
In chapter 1, to draft an answer to these questions, I gathered data from Ochrophyta genomes and transcriptomes and started looking for known photoreceptor domains. The resulting data showed that some photoreceptor types are conserved in Ochrophyta (aureochromes, a group of stramenopile-specific LOV-based sensors, and cryptochromes), while others are restricted to some algal groups only. Diatoms in particular show a duplication of several cryptochrome-like proteins, some of which might be new photosensors. Phytochromes and rhodopsins probably evolved from horizontal gene transfer, possibly through viral infection. Although the evolutionary origin of diatom photoreceptors has not been resolved yet, this study has provided comprehensive information on the photoreceptor repertoire in Ochrophyta, which is the subject of a manuscript, in preparation.

Still very little is known about phytochrome action in diatoms. To gain insights into the physiological function of DPH and its relevance for diatom life in the oceans, I addressed in the second chapter the following question:

*What is the light phytochrome responds to in the marine environment known to lack red and far-red lights? Do different phytochromes of different diatom species have the same spectral properties? Where the diatom containing phytochrome lives? What is the contribution of phytochrome to the diatom life?*

In Chapter 2, I combined studies in the lab to establish the *in vivo* action spectra of DPH in *Phaeodactylum tricornutum* to projection of DPH activity in modeled and measured marine light fields. This showed that blue light is the prominent waveband favoring DPH activation in the marine environment, and red, as well as green lights, act mostly in reverting blue light activation. I confirmed the presence of DPH in the open ocean by taking advantage of the *Tara* oceans data (Carradec et al., 2018), and by characterizing novel DPH from different species and environmental sequences, I showed these DPH are mostly from centric diatoms and are shared very conserved light-sensing properties. Overall, these results provide completely novel insights both for DPH functioning and its putative role in marine diatoms. We hope to publish these results very soon after the defense.
As this study showed the quasi-absence of pennate diatom phytochromes in the open ocean, we considered the possibility that DPH presence in pennate diatom is associated to the distribution of some of these species in the benthic environment.

*Is there a link between DPH and the light field in benthic environments?*

As noticed in chapter 1, many pennate diatoms have several copies of DPH genes and are mostly benthic. I therefore also explored the photosensing properties of the DPH homologs in the benthic diatom *Amphora coffeaeformis*. I successfully cloned 3 of these DPHs and showed that they all possess a red-far red light spectra but with variations of the absorption maxima and of the photocycle. To get insights into the function of these DPH, we explored the response of *P.tricornutum* and other pennate diatoms to red-enriched light field like the ones occurring in sediments, and showed the involvement of DPH in acclimation to red light.

Finally, as underlined in this introduction, knowledge on the DPH transduction cascade is missing, and the integration of this bacteriophytochrome in a eukaryote is a puzzle. In the last chapter, I addressed the following question:

*Who are the actors of the different DPH signaling steps?*

I started the *in vitro* characterization of *P.tricornutum* chromophore-producing enzymes, i.e heme oxygenase and putative biliverdin reductases. I also addressed the question of DPH localization *in vivo*, and identify the first component of its signaling cascade by Yeast Two Hybrid. We noticed that other factors, such as cell density in the culture and agitation, also regulate DPH-regulated genes, suggesting that these genes are the targets of signaling networks integrating several environmental cues.

I will discuss the results obtained in these different chapters as a whole in a final conclusion.
CHAPTER 1: PHOTORECEPTORS
REPERTOIRE AND DISTRIBUTION IN
OCHROPHYTA

Carole Duchêne, Jean-Pierre Bouly, Angela Falciatore, Marianne Jaubert

CNRS, Sorbonne Université, Institut de Biologie Physico-Chimique, Laboratoire de Biologie du chloroplaste et perception de la lumière chez les microalgues, UMR7141, F-75005 Paris, France

With the rapidly increasing genomic resources for diatoms (genomes and transcriptomes), it was timely to systematically mine for their photoreceptor content using the different characteristic domains of these proteins.

Some very basic questions can be asked, such as inventorying the different known
photoreceptors types that are found in this eukaryotic branch. As this has only little been
addressed, we expect to find new photoreceptors. Expending more widely this analysis to
the Ochrophytes allowed to determine what part of this repertoire is specific to diatoms,
or specific to some other ochrophyte lineages, and what is shared among Ochrophyta. As
light fields can be highly different in aquatic environments, it would be interesting, when
possible, to link the photoreceptor repertoire of an algal group to its ecological niche
(oceanic, freshwater, brackish), as well as their lifestyle (planktonic, benthic) or trophic
characteristics (obligate phototroph, mixotroph, heterotroph).
My contribution to this study consisted in gathering the available genomes and
transcritpomes sequences, performing HMM search to look for photoreceptors light-
sensing domains, and Sequence Similarity Networks. I performed simple phylogenies, to
approach the question of the evolutionary origins of the different photoreceptors. When
needed, I ran additional analyses, except the homology modelling.

Manuscript in preparation
Carole Duchêne, Jean-Pierre Bouly, Angela Falciatore, Marianne Jaubert

CNRS, Sorbonne Université, Institut de Biologie Physico-Chimique, Laboratoire de Biologie du chloroplaste et perception de la lumière chez les microalgues, UMR7141, F-75005 Paris, Franc

ABSTRACT

Ochrophyta are a diverse class of photosynthetic eukaryotic algae resulting from
secondary endosymbiosis and including ecologically important groups, such as the
unicellular diatoms and the multicellular brown algae. Light regulates many aspects of the
life of these organisms, likely through photoreceptor-mediated signalling mechanisms.
Photoreceptor proteins have been identified in the genomes of model microalgal species,
but the amount of genomic and transcriptomic data from marine phytoplankton
organisms is rapidly increasing, allowing a more complete exploration of their occurrence
and possible diversification. Using a large genome and transcriptome dataset, we provide
here a detailed repertoire of the different photoreceptors classes of Ochrophyta: the light-
oxygen-voltage (LOV)-based sensors, cryptochromes, rhodopsins and phytochromes. We
show different distribution patterns, with some photoreceptors conserved across
Ochrophyta (aureochromes, a new group of LOV photosensing protein or some
cryptochrome/photolyase family members), but also families specific to some algal
groups (some LOV-based photoreceptors, diatom-specific duplication of
cryptochrome/photolyase family members), while other photoreceptors are sparsely
distributed in Ochrophyta (phytochrome and rhodopsin), likely the result of various
horizontal gene transfer events. Hence, it appears that Ochrophyta have evolved different
sets of photoreceptors, most likely contributing to their adaptation to specific ecological
niches.
INTRODUCTION

Light is essential for life on Earth, both as a primary source of energy for phototrophs, but also as a source of information for most organisms, shaping different aspects of their physiology and life cycle. Accordingly, organisms have evolved biological mechanisms to perceive and respond to light (in colors and intensity) that suit different ecological niches (Björn, 2015). Light is absorbed by specific proteins called photoreceptors, which usually contain a chromophore molecule that gives photoreceptors their light-sensing properties (Möglich et al., 2010). Following light absorption, changes in the chromophore conformation are reflected in the structural changes of the protein, triggering signaling cascade(s) leading to cellular response.

Known types of photoreceptors can be classified based on their light-sensing abilities and the nature of the chromophore they bind (Möglich et al., 2010). Most UV-A-Blue light photoreceptors use flavin-based chromophores to perceive light. Flavin-based photoreceptors can be subdivided in three categories. Cryptochrome photoreceptors are known in all kingdoms of life (Bacteria, Archaea and Eukaryotes). They are part of a larger protein family called Cryptochrome/Photolyase Family (CPF) also including DNA photolyases (blue-light activated enzymes that repair UV-B DNA damage such as cyclobutane pyrimidine (CPD) dimer or 6-4 pyrimidine-pyrimidone photoproducts), Cry-DASH for which single and double strand DNA repair, and photoreceptors activities have been associated in a taxa-dependent manner (Kiontke et al., 2020), and light-insensitive transcription repressors (animal type II cryptochrome) (For review see (Chaves et al., 2011)). These proteins usually contain a non-covalently bound Flavin Adenine Dinucleotide (FAD) as cofactor, and eventually additional antenna chromophores (Essen et al., 2017). Other important flavin-blue light photoreceptors include LOV-based photoreceptors. LOV (Light-Oxygen-Voltage) domain, a subclass of Per ARNT Sim (PAS) super family domain, binds flavin mononucleotide (FMN), and can be found in
combination with different effector domains in all kingdoms of Life (Krauss et al., 2009; Losi and Gärtner, 2012; Glantz et al., 2016). The diversity of effector domains associated with a similar sensing domain allows modularity and customized biological response. To illustrate this aspect, one can consider the typical LOV-containing photoreceptors such as phototropins (green lineage specific photoreceptors), with 2 consecutive LOV domains associated with serine/threonine kinase (Christie, 2007), helmchrome proteins coupling two LOV domain and a Regulator of G protein Signaling (RGS) in tandem (Fu et al., 2016) or aureochromes (stramenopiles specific) that combine a LOV-domain with a DNA-binding bZIP domain, thus being a blue-light-driven transcription factor (Takahashi et al., 2007; Kroth et al., 2017). Last flavin-based group of blue-light photoreceptors are BLUF sensors (Blue Light Using Flavin), which are essentially known in Bacteria but also found in several Euglenozoa (Gomelsky and Klug, 2002; Iseki et al., 2002; Masuda and Bauer, 2002).

PYP (Photoactive Yellow Protein) also absorbs blue light but binds 4-hydroxycinnamic acid as chromophore. However, it has only been reported in bacteria so far (Meyer et al., 2012).

Plant UV-B light photoreceptors, UVR-8, do not bind an additional chromophore as their photosensing properties are based on light absorption by the amino acid tryptophan in the protein itself. Study of the phylogeny of these photoreceptors suggests that these are present in Viridiplantae only (Fernández et al., 2016).

Phytochromes are typically red/far-red light sensors that contain covalently attached bilin derivatives as chromophore to perceive light (Rockwell et al., 2006). They are known in bacteria and eukaryotes, and are typically composed of a photosensory module (PSM) containing the PAS, GAF and PHY domains, followed by highly variable output modules (for review see Rockwell et al, 2006; (Rockwell and Lagarias, 2020). Eukaryotic phytochromes form two separate branches on a phylogenetic tree, one containing Archaeplastida and Cryptophyta phytochromes while the other contains fungal and Stramenopile phytochromes (Duanmu et al., 2014; Fortunato et al., 2016). Bacterial phytochromes branch in between these two groups, and the origin of eukaryote phytochrome is a debated issue (Duanmu et al., 2014; Li et al., 2015; Kooß and Lamparter,
2017). Cyanobacteria possess an additional group of specific phytochrome-related photoreceptors (Cyanobacteriochromes, CBCR), which only require a bilin-binding GAF domain, and show a wide variation of absorption spectra reflecting potential light environments adaptation(Ikeuchi and Ishizuka, 2008).

All photoreceptors characterized and mentioned above are intracellular or plasma membrane-associated. Rhodopsins on the other hand are multi-pass membrane proteins. They contain 7 trans-membrane helices and bind a retinal chromophore giving them the ability to sense green and blue light. Rhodopsins are classified in 2 groups, microbial rhodopsins (type I) and animal rhodopsins (type II), that do not share strong sequence homology (Ernst et al., 2014). Animal rhodopsins (opsins) are part of a wider group of membrane proteins also containing chemical receptors (G protein coupled receptors). Microbial rhodopsins can mainly have two biological roles, a bioenergetic one with the use of light as source of energy (through the generation of an electrochemical gradient) or a photosensory function that uses light as source of signal. Several chemical activities can be associated with the light perception properties such as ion transport (H+, Na+, Cl-) associated with the two biological functions (bioenergetics roles or photoperception), light-induced enzymes (photoperception). The recently discovered heliorhodopsins have no ion transport activity and no effector domain, and their function remains unknown (Pushkarev et al., 2018) (For review see (Rozenberg et al., 2021)).

Finally, if different photosensory protein domains can be combined with diverse effector domains, combinations of photosensing domains have also been uncovered in recent years, giving rise to proteins with new light-sensing abilities. Examples of chimeric photoreceptors are present in bacteria, with Ppr, a chimeric photoreceptor from *R. centenum* in which a Photoactive Yellow Protein is fused at amino-terminal of a phytochrome (PYP+PHY) (Jiang et al., 1999), in ferns with neochrome, an association of LOV (amino-terminal) and phytochrome domains (LOV + PHY) (Kawai et al., 2003), or in green algae with the dualchrome a recently characterized photoreceptor composed of a cryptochrome and a phytochrome (CRY+PHY) (Makita et al., 2021).

In aquatic environments, photoperception raises different challenges. Underwater light field varies with depth, due to the wavelength-dependent absorption properties of
water, and the content in suspended particulate material (such phytoplankton and
mineral) and coloured dissolved organic matter (Kirk, 2011)(Kirk, 1994; Mobley, 1994).
This provides a full variety of photic niches from solar spectra at the surface, to
blue/green light in clear oceanic waters or red-enriched light field in turbid waters.
Photoreceptor-like sequences from marine microalgae have been recently identified by
omics approaches but their physiological properties have been characterized on a very
few of them (Jaubert et al., 2017; Petroutsos, 2017). Initial studies have revealed that
some algal photoreceptors are similar to those known in plants but new variants with
different spectral tuning and algal-specific light sensors have also been found, changing
current views and perspectives on how photoreceptor structure and function have
diversified in phototrophs experiencing different environmental conditions (for review
(Jaubert et al., 2017). Stramenopiles are a class of eukaryotes containing a large number
of photosynthetic algae (Ochrophyta) that are of ecological importance, especially in the
marine environment (de Vargas et al., 2015). Ochrophyta are a monophyletic clade and
include a diversity of organisms, from the unicellular diatoms which are important
primary producers in the open ocean, to the brown algae kelp forest in the coastal regions
(Dorrell et al., 2022). Moreover, beyond different potential adaptation to specific light
environments, Ochrophyta can have different trophic modes: photoautotrophy,
mixotrophy and heterotrophy (some genus have to potential for both auto- and hetero-
trophy, while a few of them have completely loss their photosynthetic abilities)(Beisser
et al., 2017; Onyshchenko et al., 2019; Kamikawa et al., 2021; Onyshchenko et al., 2021).
These algae result from secondary endosymbiosis of a red algae, and their genome (at
least in diatoms) contains red algal genes transferred from the rhodophyte symbiont to
its host, but also genes of green algal origins suggesting previous association with a green
algae(Dorrell et al., 2022). Horizontal gene transfer from bacteria also occurred at a high
rate in Ochrophyta, and their genome is often considered as an evolutionary mosaic
(Dorrell et al., 2021). There are initial evidences that this branch of Eukaryotes harbors
specific types of photoreceptors, such as aureochromes (bZIP-LOV photoreceptor) (Kroth
et al., 2017), but there is currently no global view of the light sensing capabilities of these
organisms. With the increasing number of omics data (genomics, transcriptomics) from
marine phytoplankton, Ochrophyta thus appear especially interesting from an ecological
and evolutionary point of view to deeply explore marine photoreceptors and get novel insights on the relevance of light sensing for life in aquatic environments.

To this aim, in this work we gathered genetic data (transcriptome and genomes) from different Ochrophyta to explore diversity and distribution of photoreceptors in this branch of the eukaryotic Tree of Life. We focused on LOV-containing proteins, cryptochromes, phytochromes and microbial rhodopsins as no UVR8-, BLUF- or PYP-like photoreceptors have been identified. Even if the physiological roles of photosensing have not been assessed, we uncover new LOV-containing proteins with original domain association as well as a potential new cryptochrome sub-family. Moreover, while some photoreceptors are ubiquitous into Ochrophyta, some appear phyla-specific. Hypotheses on the acquisition of some photoreceptors in Ochrophyta are also discussed.

# RESULTS AND DISCUSSION

## DATASET AND CONSERVATION OF PHOTORECEPTORS

We gathered nearly 9 million genes from 69 Stramenopiles genomes and 260 transcriptomes available on a variety of platforms (NCBI, JGI, MetDB, private repository, see Supp.Data 1 for full description of the dataset). Among this dataset, we used 276 Ochrophyta datasets, to which we added 33 genomes from Oomycota (sister group of Ochrophyta) and 20 genomes and transcriptomes Bigyra (sister group of Gyrista, (i.e. Ochrophyta + Oomycota) to try to infer the origins of the different photoreceptors in this group of eukaryotes. As diatoms (Bacillaryophyta) are one of the most studied groups, they represent the majority of the organisms in our dataset (44% (147/329) of the transcriptomes and genomes). Most of the Ochrophyta studied here are phototrophs, with the notable exception of some Chrysophyceae species (12 species, but for some the exact trophic mode is not known) and one diatom strain (*Nitzschia sp*) which are heterotrophs (photosynthesis loss). Moreover, one diatom strain (the diatom *Fragilaria accus*) and some Chrysophyta are known to live in freshwater, the others being marine or brackish.
<figure><img src="image_15.png" /><figcaption>Figure 1. Photoreceptors repertoire in Stramenopiles. Presence of different photoreceptor types visualized on a multi-gene phylogenetic tree of Stramenopile species, colored according to the main algal groups. Outer circles indicate the presence of different photoreceptor types (squares: LOV-based, round: CPF, stars rhodopsins and triangles phytochrome). (White background in the Stramenopiles tree refers to two Eustigmatophyceae and one Xanthophyceae, See FigS1 for the tree with the species names)</figcaption></figure>

We looked for known photoreceptor's light sensing domains in the dataset by using Pfam HMM models or custom HMM models (See Material and Methods and Supp.Data 2). A Sequence Similarity Network (SSN) (Atkinson et al., 2009) was computed for each photoreceptor type (including also reference sequences from other organisms) to classify proteins into sub-groups. Additional stringent filtering criteria specific for different photoreceptor types were further applied for the presence of conserved amino acids interacting with the chromophore, overall structure of the protein, and the presence of transmembrane helices. The protein domain architecture was also examined and eventually followed with phylogenetic tree reconstruction.
A very contrasted picture emerges regarding the photoreceptor domain content among the different Ochrophyta groups (Fig.1). Indeed, some photoreceptors appear ubiquitous in Ochrophyta such as aureochromes and Cry-DASH cryptochromes, while others (helmchromes, rhodopsins, phytochromes) are sparsely distributed on the Ochrophyta species tree. Interestingly, genomes of Oomycetes, sister group of Ochrophyta, exhibit no CPF of the 6-4 photolyase class and no aureochrome and other LOV-based sensors, while Bigyra also have some of the Cry-DASH class, and some LOV-based photoreceptors of the aureochrome family, suggesting a particularly rich content in photoreceptors amongst the Ochrophyta. It is to note that regarding the transcriptomes data, some photoreceptor's domain types may not have been detected in this dataset because they were not expressed. On the other hand, some species appear with high copy numbers of some genes, possibly due to assembly problems. For example, manual examination of phytochrome sequences showed that some truncated sequences, when put together, allow the reconstruction of a single photoreceptor. This manual examination was not done for all photoreceptor types, so the copy numbers are to consider with care. Details of each type of photoreceptors are presented in the following sections.

## LOV-BASED PHOTOSENSING: UBIQUITOUS AND SPECIFIC FAMILIES

LOV-domain model was designed with reference sequences from Glantz et al., 2016. Search for this domain in the stramenopile database resulted in 5800 sequences with LOV-domain hits.

Conserved amino acids defined to be necessary for the photochemical activity have been used as functional criteria (Glantz et al., 2016). More precisely the conservation of highly conserved FMN binding motif (GRNCRFLQ), especially the Cys involved in the adduct formation with FMN was verified. As shown by a representative alignment of the different sub-groups of LOV-domain (FigS2A), a strong conservation of the amino acids involved in FMN binding is observed, suggesting these proteins might be functional even if specific substitution affecting photochemicals properties cannot be excluded. Moreover, the presence of other associated protein domains was examined. To classify these genes, we first conducted a SSN using the full length sequences, including reference sequences from
plants, fungi, bacteria and other algae (sequences from Glantz, 2016), with an alignment
score of 40 allowing separation of proteins with less than 45 % identity. This resulted in
a separation of proteins, with 3 sub-groups containing no stramenopile sequences but
only reference sequences from plants with the Adagio/Zeitlupe sub-group, and two sub-
groups of fungal proteins including White Collar and Vivid proteins. These groups were
removed for further analysis, allowing the identification of 6 sub-groups (alpha, gamma,
zeta, theta, Stramenopile LOV sub-group and one larger group, FigS2B) containing
Stramenopile sequences and carrying the LOV-FMN binding motif (FigS2A).

3 small sub-groups (of 10 to 40 sequences) contained only sequences of one alga type (alpha: Phaeophyceae, gamma: Pelagophytes, theta: pennate diatoms) (FigS2). Zeta sub-group contained a mix of different algae sequences, including reference sequences from Glaucophytes.

One sub-group contained 130 sequences from stramenopiles only and will be called
Stramenopile-specific LOV sub-group (Fig1 and S2).

Last, one large sub-group contained more than 4000 proteins including reference
sequences from Viridiplantae, aureochromes and helmchromes from Ochrophyta. This
group was further subjected to a second, more stringent SSN step with an alignment score
of 60 allowing separation of Viridiplantae and Ochrophyta photoreceptors (FigS2C;
named SSN60_group1 to 5 in FigS3).

Except for aureochromes and Stramenopile-specific LOV proteins, each of the resulting
groups seem to be restricted to one branch on the Ochrophyta species tree (examples of
the small sub-groups mentioned above, see also below and FigS3). The SSN approach used
here does not enable us to decipher the relations between groups, and we would need to
perform cautious phylogeny on the LOV-domain(s) to decipher the origin and
evolutionary path of LOV-based photoreceptors. This analysis with 2 sequential SSN
resulted in 13 LOV subgroups that will be discussed below.
Aureochromes: ubiquitous blue-light photoreceptors in Ochrophyta

Proteins possessing bZIP domains and grouping with characterized aureochromes in SSN were annotated as aureochromes. These photoreceptors were first identified in the multicellular xanthophyte *Vaucheria frigida* as photomorphogenesis regulators, and were recently shown to be master regulators of blue-light responses in the model diatom *Phaeodactylum tricornutum* (Mann et al., 2020). These proteins have been identified in all Ochrophytes, some Bygira, but not in Oomycetes as previously described (Kroth et al., 2017).

Previous work distinguished different groups of aureochromes, named from *V. frigida* auerochromes: “group 1” and “group 2” (Takahashi et al., 2007; Schellenberger Costa et al., 2013). Based on functional characterization, both group 2 aureochromes from *V. frigida* and *P. tricornutum*, are not able to bind the flavin chromophore and thus not considered as bona fide blue light photoreceptors (Takahashi et al., 2007; Banerjee et al., 2016). To explore whether these two proteins might reflect all type 2 aureochrome, we looked at the conservation of a methionine in position M301, replacing a Valine in PtAureo1a (V253) and shown to impair FMN binding in PtAureo2 (Banerjee et al., 2016). In our alignment of type 2 sequences, this position was occupied by a cysteine in VfAureo2, but other proteins had a Valine or an Isoleucine, suggesting that some of the group 2 aureochromes from our dataset could be functional photoreceptors in several organisms (FigS2A). It has also to be noted that differences in the bZIP DNA binding site were also observed between group 1 and group 2 aureochromes suggesting different DNA binding abilities (FigS4B).

We made phylogenetic tree of the different aureochromes found in our data (Fig.2 A, see FigS4 for the unrooted tree). The sequences branched per “Aureotype” with three main branches: type 2, type 1a, and type 1b and c. Type 1b and c are close to each other, forming a clade named 1b/c. On each branch, sequences form clades by algal group, suggesting ancient multiplication and further evolution within the different species. Interestingly, in our data, all groups of algae have a copy of “type 2” aureochrome, one or more copies of “type 1a” and one or more copies of “type b/c” (Fig.2 A). As example, in the species *V. littorea*, in addition to type 2 and type 1a, we report a third aureochrome of type 1 b/c (see
red arrows in Fig2A) while in *V. frigida*, the species in which aureochromes were first discovered, only one of each type 1 and type 2 aureochrome has been characterized (Takahashi et al., 2007; Ishikawa et al., 2009). Interestingly, we also found aureochrome in heterotrophic species such as Chrysophyceae, and the diatom that has lost photosynthetic capacity (*Nitzschia sp* in our study) as already reported for *Nitzschia putrida*, another heterotroph diatom genome (Kamikawa et al., 2021).

Compared to diatoms, which all seem to have one type 2 and three type 1 (1a, 1b, 1c), Pelagophyte, Dictyophytes and Chrysophyte have only two copies of type 1 (1a-like and 1b/c like). Phaeophyceae show expansion of this family with 5 or 6 aureochromes, with two potential type 2, one being a small group containing phaeophyceae sequences close to *Ectocarpus siliculosus* (Esil)Aureo4 and branching at the base of the type 2 aureochrome; two type 1a (see sister Phaeophyceae branches in the 1a branch, also named EsilAureo1 and EsilAureo3 in *Ectocarpus siliculosus* annotation), one sister to diatom type 1b (also named EsilAureo 5) and one close to diatom type 1c (not found in *Ectocarpus siliculosus*, but found in the other *Ectocarpus* strain in our dataset). Whether all Phaeophyceae possess all 6 types of aureochromes has not been investigated in detail, but could be linked to the evolution of multicellularity.

Here, we show that these photoreceptors are ubiquitous within Ochrophyta, but not found in Oomycetes. Type 2, 1a and 1b/c thus seem common aureochrome types in Ochrophyta, suggesting that this multiplication was already present in their ancestor, while multiplication further occurred in brown algae and diatoms. Moreover, the presence of aureochromes in 4 species of Cafeteria/Halocafeteria genus, belonging to Bigyra clade, suggests that this photoreceptor might have originated in the common ancestor to Bigyra and Ochrophyta, with a loss in Oomycetes. Alternatively, this photoreceptor could have been transferred to Cafeteria after their divergence from Oomycetes and Ochrophyta. The exact evolutionary relations between these groups need to be further examined. Moreover, as the tree presented here is rooted with Cafeteria's Aureochrome, this might not be the appropriate outgroup to infer evolutionary history (See FigS4 for the unrooted tree, which shows the same topology).
<figure><img src="image_16.png" /><figcaption>Figure 2. LOV-based photoreceptors in Ochrophyta. A, Aureochrome phylogenetic tree calculated from the alignment of bZIP and LOV domains. The tree was rooted with Cafeteria aureochrome sequences; see Fig S4 for unrooted tree. Branch colors correspond to aureochrome groups according to annotation in *P*.</figcaption></figure>
*tricornutum*, Names are annotated according the *P. tricornutum* annotation (DiatomAureo 1a, b, c and 2) or *Ectocarpus* species (Phaeophyceae « Phaeo »Aureo1 to 6). Red arrows indicate *Vaucheria littorea* aureochromes. B. Unrooted phylogenetic tree of the new group of LOV-based photoreceptors that seems to be widespread in Ochrophyta (full length protein was used). In both trees, the colors of the taxa correspond to the algal groups to which the genes belong (*Vaucheria* and Eustigmatophytes on black), and purple circles on the branches indicate local-bootstraps support >0.7

## A new group of LOV-based photoreceptors widespread among Ochrophyta

A group of LOV domain-containing proteins was identified in Ochrophyta from SSN. These short proteins (on average 190 amino acids long) are all made of a single LOV domain (Fig S3B) with a conserved chromophore binding site (FigS2A). The LOV domain is present after 64 amino acids (on average), which might recall the structure of the Vivid proteins in Fungi (Yu and Fischer, 2018).

These proteins are present in a large number of species (3rd circle in Fig1, see also FigS3A): diatoms, brown algae and Raphidophyceae, but absent in Pelagophytes and Dictyophytes branches. This might suggest a common origin of this putative photoreceptor in the ancestor of diatoms, brown algae and Raphidophyceae, with a loss in the Pelagophyte/Dictyophyte branch. Phylogenetically (Fig2B), these proteins form separate branches according to species, with 2 groups of diatom genes. Some diatoms such as *Thalassiosira oceanica* possess genes in both branches, but one branch contains mostly genes from *Thalassiosirale* species.

## Other LOV-based: Branch-specific specialization?

As indicated above, SSN analysis allowed us to identify several interesting groups of LOV domain-containing proteins, found restricted to some Ochrophyta groups (Fig. S2 and FigS3).

The “Alpha” sub-group, identified in the first SSN (alignment score 40) contains sequences from Phaeophyceae and from Vaucheria (closely related to Phaeophyceae). Most of these sequences possess a PAS_9 (PF13426) domain, followed by the LOV domain with FMN
binding site, an architecture common in all three domains of life (Glantz et al., 2016), and
suggested to have photoperception roles in *Arabidopsis thaliana* (Ogura et al., 2008).

The "Gamma" sub-group, restricted to Pelagophytes, contains proteins with a long N-
terminal region followed by the LOV domain. No known conserved domains could be
identified in this N-terminal region.

The Zeta sub-group encompasses a mix of proteins from Labyrinthulomycetes (Bigyra), from one diatom species (*Odontella aurita*) and reference sequences from Glaucophytes and Chlorophytes (Glantz et al., 2016). From the Interpro database, this architecture is also known in two proteins from Rhizaria. These proteins possessed GTP cyclohydrolase II domain, which is involved in riboflavin biosynthesis (Ren et al., 2005).

An important group of LOV-based proteins for which experimental studies exist are the
helmchromes (Fu et al., 2016), which contain PAS_9 and RGS domains (Regulator of G
protein Signaling). These proteins are thought to be involved in phototaxis of flagellated
cells, and are detected by immunofluorescence in the flagella of sperm cells in some
Phaeophyceae and in the flagella of some Chrysophyceae. Here, these photoreceptors
were not found ubiquitously in Ochrophyta: they are present in some branches of
Chrysophyceae, Phaeophyceae, Pelagophytes and Dictyophytes, but they are absent from
diatoms and from Raphidophytes. Our distribution of helmchrome in Ochrophyta agrees
with experimental data detecting the presence of this protein in Phaeophyceae and
Chrysophyceae (Fu et al., 2016), and also with the presence of helmchromes in
Pelagophytes and Dictyophytes reported recently (Coesel et al., 2021). The domain
architecture of these helmchromes is RGS-LOV-PAS9-RGS-LOV-PAS9, although
sometimes sequences are too short to span all the domains. Fungi also possess RGS-LOV
domain proteins of unknown function, which form a separate group in a phylogenetic tree
with helmchromes (FigS4), suggesting independent evolution. Interestingly, these fungal
proteins are recruited to the plasma membrane upon blue-light illumination (Glantz et al.,
2016; Glantz et al., 2018), suggesting that helmchromes could perform their photosensing
function through the same type of mechanisms.

Although concerning only a few diatom strains, some proteins showed new domain
association such as LOV domains fused to CAP-GLY (Cytoskeleton Associated Proteins
with glycine-rich domain), which might bind the microtubules (Weisbrich et al., 2007) (FigS3). In the Interpro database, this structure was not reported for species other than diatoms, suggesting that this new LOV protein class is specific for diatoms. This is an interesting domain combination, as such proteins could regulate light-induced intracellular movements of organelles in the absence of phototropins, which are known to regulate plastid movement in Viridiplantae (Christie, 2007).

Finally, by looking for LOV domain-based photoreceptors, we identified a wide variety of proteins, from the ubiquitous photoreceptor aureochromes to novel putative photoreceptor families restricted to some species only. Eukaryotic LOV-containing proteins are thought to have evolved from bacterial gene transfer, either through endosymbiotic gene transfer from the mitochondria (plant phototropins and fungal LOV-photoreceptors) or from the chloroplast (plant Zeitlupe) (Krauss et al., 2009). This hypothesis has been challenged for the origin of Zetilupe, but plant phototropins remain a sister group to alpha-proteobacteria LOV proteins in more recent analyses (Losi et al., 2015). Aureochromes group close to the plant phototropins (Ishikawa et al., 2009; Djouani-Tahri et al., 2011), suggesting common origin. However, independent horizontal gene transfer (HGT) is possible for the other LOV photoreceptors identified here. For example, the LOV-HK photoreceptors from the green algae *Ostreococcus* and *Micromonas* are closer to bacterial genes than to phototropins, suggesting more recent horizontal gene transfer from bacteria (Djouani-Tahri et al., 2011). The origin of the LOV domains in Ochrophyta (common origin with plant phototropins vs independent HGT), and the possible domain shuffling events (Di Roberto and Peisajovich, 2014), will need to be further analyzed with phylogenetic studies of the LOV domain only. In addition, we found here new eukaryotic representatives of LOV-based photoreceptors, particularly represented in Dictyophyceae and Pelagophyceae (FigS3), that might be included in broader studies on the origin of LOV domain in Eukaryotes.
CPFs constitute a large group of flavoproteins with an impressive diversification (Chaves et al., 2011; Fortunato et al., 2015). CPF proteins share a common domain called Photolyase Homology Region (PHR) but are associated with a multitude of functions. CPF can be phylogenetically separated into 5 different superclasses regrouping proteins with different functions : (i) the bacterial Fes-BCP or Cry-Pro class, (ii) the 6-4 photolyase class including animal light-dependent and light-independent cryptochromes, (iii) the class I/III CPD photolyases including the plants cryptochromes (photoreceptor) and plant-like cryptochromes, (iv) the class II CPD photolyases, and (v) Cry-DASH (Drosophila Arabidopsis Synechococcus Homo) class. Due to the large diversity among the different members of this family, CPF domain architectures alone did not lead to sufficient information for a functional prediction (i.e., photolyase vs blue light photoreceptors and light-independent transcriptional regulation). Therefore, several criteria such as SSN, phylogenetic tree, or biochemical information need to be used and combined to give insights into the possible function of these proteins. A new functional classification of this protein family based on the use of computational multiple profile models has been recently proposed (Vicedomini et al., 2022). The model diatom *Phaeodactylum tricornutum* possesses 7 CPF members: PtCPF1, a 6-4 photolyase with gene regulation activity (Coesel et al., 2009), PtCryP, a plant-like cryptochrome having a role in gene expression regulation (Juhas et al., 2014; König et al., 2017), PtCPF2 and 4 belonging to the Cry-DASH sub-family, two class II CPD, PtCPD3 and PtCPD4, and PtCPD1, a NCRY (New CRY) protein which might represent a new class of light photosensing protein (Emmerich et al., 2020; Vicedomini et al., 2022).

We tested several HMM domain models (FAD binding domain 7-PF03441, DNA photolyase-PF00875, and custom model using COG0415 sequences from the conserved protein domain family PHRB). Only the one using COG0415 sequences could recognize all 7 members of the CPF family in the model diatom *Phaeodactylum* and was further used for the search in our database. The resulting SSN of the dataset (1830 sequences including references) separated proteins into 13 groups (FigS5) when applying an alignment score of 90.
Interestingly, sub-classes containing only diatoms and Bolidophyceae sequences (group 1 and 2) and a mix of stramenopile ones (group 3 to 5) did not group with other known sequences, even with lower alignment scores (FigS5). Reconstruction of a phylogenetic tree agreeing with the SSN (Fig3), placed group 1 and 2 close to Plant-like Cryptochrome including PtCryP (Juhas et al., 2014), and these were further named "CryP2a" and "CryP2b".

Group 3 to 5 form an independent clade. All sequences in this clade had DNA photolyase domain (PF00875) followed by alpha beta hydrolase 6 domain (PF12697), and most of them are possibly addressed to the chloroplast based on HECTAR prediction (Gschloessl et al., 2008) (FigS5A). Mining other proteins showing this structure on Interpro retrieved unreviewed proteins from land plants and green algae (Viridiplantae). This observation was already done in a work on photolyases by Emmerich et al, 2020. As these proteins did not possess FAD binding domain, they are unlikely to function as photoreceptors.

CPF from the 6-4 photolyases, Cry-DASH class and class II CPD photolyase sub-families are present in all Ochrophyta and Bigyra (see Fig1 and S5). However, Oomycetes lack Cry-DASH as already observed in some fungi such as *Aspergillus nidulans* (Schumacher, 2017; Corrochano, 2019). Plant-like cryptochromes group is more sparsely distributed but appears conserved in diatoms. Interestingly duplication of Plant-like cryptochromes, CRY-DASH and class II CPD photolyase seems to be uniquely found in diatoms and in their sister group, the bolidophyceae, suggesting duplication occurred in their common ancestor. Last, NCRY proteins are only found in diatoms, Bolidophyeae and Raphidophyceae.
<figure><img src="image_17.png" /><figcaption>Figure 3. Ochrophyta CPF family protein tree. Tree was rooted at the base of the CryPro class. Branch colors correspond to major CPF class, see also FigS5 to 8 for zoom on different branches. Taxa colors correspond to the algal groups to which the genes belong, and purple circles on the branches indicate local bootstraps support >0.7</figcaption></figure>
Looking more in detail the Class II-like photolyase CPD, duplication seems to have occurred in Chlorophyta and Diatoms, forming a sister clade to the main CPD photolyase class II branch (Fig3, FigS6). Due to a strong divergence of the primary sequence, multi sequence alignment and homology modeling of the structure did not allow to identify active sites of photolyase activity. However, while 25% of the diatom class II CPD photolyases are predicted to be addressed to organelles (22/87 diatom proteins with mito/chloro/signalP prediction), the duplicated Class II CPD photolyase in diatoms do not seem to be addressed to the organelles (0 mito/chloro/signalP prediction with HECTAR) (FigS5). These results, in agreement with functional characterization of PHR1 and PHR2 in Chlamydomonas and PiPhr1 and PiPhr2, from the antarctic diatom *Phaeodactylum tricornutum* ICE-Hand suggest that only diatoms and Chlorophytes may have specific nuclear and chloroplast-targeted double strand CPD DNA photolyase activity (Small and Greimann, 1977; Petersen et al., 1999; An et al., 2021).

Similar results have been observed with CRY-DASH proteins whose function is still unclear (ssDNA photolyase or light sensors). We clearly distinguished 2 clades of Cry-DASH in Ochrophyta, one CRY-DASH-like clade containing only diatoms and another one containing sequences from all Ochrophyta (Fig3, FigS7). This includes a diatom clade, a brown algae clade, then a group with mixed Pelagophytes, Dictyophytes and Raphidophytes. Cry-DASH proteins have been found to be localized in chloroplasts or mitochondria in many organisms. Here, 44% of the diatom CRY-DASH proteins were predicted to be addressed to organelles (PtCPF2 group), while only 16% of the diatom CRY-DASH-like ones were (PtCPF4 group, FigS5). This might also suggest cellular compartmentalization and possible functional specialization. Interestingly, like for the class II CPD photolyases, duplication also occurred in Chlorophyta, and the duplicated Chlorophyta genes are on the same branch as CRY-DASH-like diatom genes.

Only one gene copy of the 6-4 photolyase class has been identified in Ochrophyta, a potential common feature to photosynthetic organisms, but this was also the case in Bigyra. The Ochrophyta 6-4 photolyases form a clade with a Chrysophyceae, a Phaeophyceae and 2 diatom clades (one sister to Phaeophyceae, the other basal to these), while Oomycete proteins group close to Viridiplantae (Fig3, FigS8). In *Phytophtora* species in particular, we could identify up to 3 copies (FigS8). As in insects, this could
mean sub-functionalization of the different copies with either 6-4 photolyase activity or
cryptochrome functions. To support this idea, we calculated the isoelectric point (pI) of
the 6-4 photolyase family members from oomycetes, considering that proteins with
different pI might have different functions (Kiraga et al., 2007; Mohanta et al., 2019). Most
photoreceptors of the CPF family (animal or plant) are characterized by an acidic pI. We
found that the 3 groups of oomycota sequences have different pI: mean±sd of 5.37±0.44
for group CPF1aa, 7.3±0.99 (CPF1ab) and 6.05±0.8 (CPF1b, FigS8B). Moreover, CPF with
photoreceptor functions potentially exhibit a C-terminal extension, a feature found in at
least two copies of the proteins present in Oomycetes. Indeed, the DNA photolyase
domain starts at position 105 (±11) for CPF1aa, position 3 (except one sequence) for
CPF1ab and position 96(±23) for CPF1b. This suggests at least two different functions are
carried by these proteins, and that CPF1aa probably are photoreceptors. Although
speculative, it would also seem surprising that multiple proteins with a DNA repair of
pyrimidine-pyrimidone (6-4) photoproducts which represent only ~20% of the DNA UV
damage, would have been kept in Oomycetes while no CPD photolyase has been detected,
suggesting that at least one of these oomycetes 6-4 photolyase family members might be
a photoreceptor.

Plant-like Cryptochromes are the CPF that branch closest to the plant blue light
photoreceptor cryptochromes (Fig. 3, S9 and (Juhas et al., 2014)). In the diatom *P.
tricornutum*, both *CryP* knock-down and knock-out lines showed that this protein has a
role in gene expression regulation (Juhas et al., 2014; König et al., 2017). Two groups of
stramenopile sequences can be distinguished from the SSN and on the phylogenetic tree,
forming 2 sister clades (named CryP1 and CryP2) themselves sisters to plant
cryptochromes. CryP1 group contains a few sequences from Raphidophyta, Pelagophytes
and Dictyophytes but mostly diatom ones, with the representative PtCryP. CryP2 group
contains only diatom and Bolidophyte sequences, including the "CryP-like" sequence from
the model diatom *Thalassiosira pseudonana*. Most diatoms have a gene from each group,
such as *Thalassiosira oceanica* or *Navicula sp* (FigS9). Moreover, the CryP2 itself is
separated into 2 groups (in both SSN and phylogenetictree), called CryP2 a and b. By
looking for different biochemical properties and different FAD fixation sites, CryP1 has a
much higher average pI (8.2±1.13 for CryP1 compared to 6.09±0.7 for CryP2, FigS9)
suggesting that CryP1 and CryP2 would bear different functions. Sequences of the FAD
binding sites show also some differences between CryP1, CryP2a and b (FigS10A) and
structure reconstruction on Swiss-model for one of each CryP sub-families members gave
slightly different results suggesting also different photochemical properties (FigS10B).
Overall, in Ochrophyta, we have seen duplication in diatoms into two sub-family (CryP1
and CryP2) probably associated with neofunctionalization of putative new light
photosensors.

Finally, the last class is the NCRY, with PtCPD1 as diatom representative. Its position in
the phylogenetic tree is not clear, either linked with the CRY-DASH or the CPD photolyase
class I family in different studies (Lucas-Lledo and Lynch, 2009; Oliveri et al., 2014;
Fortunato et al., 2016; Vicedomini et al., 2022). This gene was found mostly in diatoms,
with a few sequences from Phaeophyceae and Raphidophyceae. This group was identified
by SSN by Emmerich 2020, but only one functional study exists. Experimental approaches
on NCRY from *Vibrio cholera* have shown that this protein was able to bind FAD as
chromophore but was lacking photolyase activity. In addition, an *in silico* model of the
NCRY sequences showed that it presents a unique FAD active site, the absence of few
amino acids involved in CPD interactions, and the presence of a C-terminal extension
(Vicedomini et al., 2022). All these characters support the existence of a novel functional
class within the CPF, with new photochemical properties in diatoms.

Here we showed that Cry-DASH and (6-4) photolyases are well conserved amongst
ochrophyta, while there is an expansion of the different CPF families in diatoms (NCRY,
CRY-DASH-like, CPD Photolyase-like class II, and CRYP2) (FigS5). Given the complexity of
the CPF, we cannot decipher whether these are photoreceptors but members of CryP2
group might be good candidates as new photosensible proteins.

RHODOPSINS: DIVERSITY AND SPARSE DISTRIBUTION

We found 3 types of rhodopsins in Ochrophytes: proteorhodospins, histidine kinase
rhodopsins (HK rhodopsins) and heliorhodopsins (Fig. S11, S12 and 4). All proteins
reported here (86 proteins) have at least 5 trans-membrane helices as predicted by TMHMM, and were separated by SSN.

As already reported before, some diatoms possess Proteorhodopsins that may function as proton pumps using light to produce ATP. It has been proposed that these proteins could therefore have a bioenergetic role, particularly useful in iron-limited oceanic regions (Marchetti et al., 2012; Marchetti et al., 2015). We found in our dataset a similar distribution of Proteorhodopsins as reported in Marchetti et al, 2015, with the presence of this family in diatoms (with only a few additional new diatom sequences from species not included in previous analyses), in Cryptophytes and in Haptophytes (Fig S12). Within this family, some Proteorhodopsins are known to show spectral tuning, at least in bacteria: their absorption spectra is shifted towards blue if a glutamine is at position 106 (of the gamma-proteobacteria rhodopsin isolated at the Hawaii Ocean Time series station, Uniprot reference Q9AFF7) (Man, 2003). Here all diatom sequences had a leucine or a methionine at this position, suggesting green-absorbing spectra.

A group of Ochrophyta rhodopsins possessed additional signaling domains, with HATPase and Response Regulator domains (HK rhodopsins). HK rhodopsins are reported here in Raphidophytes and Pelagophyte, and have been previously described in green algae as *Chlamydomonas reinhardtii*, where it seems to work as a UV/BL photoreceptor (Luck et al., 2012). However, its biological function in green algae remains unknown. Phylogenetically, Stramenopile enzyme rhodopsins form a clade grouped with enzyme rhodopsins from other marine phytoplankton groups (*Emiliana huxleii*, Coccolithophyceae, Cryptophytes and Mameiophyceae) (Fig4A). The sequences from green algae *Chlamydomonales*, including *C. reinhardtii* histidine kinase rhodopsin 1, are sister to this group.
<figure><img src="image_18.png" /><figcaption>Figure 4 Phylogeny of the rhodopsins. A. HK rhodopsins and B heliorhodopsins, Taxa colors correspond to the algal groups the genes belong to, with a red background for viral sequences, and purple circles on the branches indicate plocal bootstraps support >0.7. Both trees were rooted with other type 1 rhodopsin from (Pushkarev et al., 2018), collapsed clades.</figcaption></figure>
We also studied the presence of heliorhodopsins, which has been identified in a functional screen of environmental sequences and has already been found in *P. tricornutum* (Pushkarev et al, 2018). Heliorhodopsins have a slow photocycle, suggesting that it would function as photoreceptor (Pushkarev et al., 2018). The majority of stramenopile heliorhodopsins identified here groups with other eukaryotes rhodopsins (Fig4B), including Chlorophyta, Glaucophyta and Euglenozoa sequences. Within this branch, at least 3 groups of diatom heliorhodopsins are found. A few stramenopile heliorhodopsins group with Fungi or Haptophyte. We underline that at the base of the clade containing haptophyte sequences, 2 sequences come from Haptophyta viruses, suggesting horizontal transfer of heliorhodopsin gene between viruses and their host.

## PHYTOCHROMES: CONSERVED BACTERIAL PHYTOCHROME (BPHP)-LIKE STRUCTURE BUT SPARSE CONSERVATION

The photosensing module of phytochrome is composed of three domains including the specific PHY domain which connects the chromophore binding domains PAS-GAF with the output module. Search for Pfam PHY domain (PF00360) resulted in proteins all showing the same domain architecture (except for one brown alga *Cladosiphon* sequence, and some sequences that are truncated): (PAS)-GAF-PHY-HisKA-HATPase-(REC), which represents the structure of bacteriophytochromes. Full length sequences were aligned and used to design stramenopiles- and phytochrome-specific domain models for each domain except for the PAS domain, which has been found not well-conserved in some diatoms (Fortunato et al., 2016). These Stramenopile- and phytochrome-specific models were used in combination with SSN to identify truncated sequences. The presence of the conserved cysteine residue involved in the linkage of the bilin chromophore was verified. Most of the proteins had the Cys residue at the N-terminal extremity of the protein, conserved with the chromophore-binding cysteine in bacteriophytochromes, but brown algae and some phytochromes from the diatom *Amphora coffeaeformis* also possessed an additional cysteine in the GAF domain, involved in chromophore binding in plant and cyanobacteria phytochromes (Rockwell et al., 2014; Fortunato et al., 2016). In addition, we noticed that Chrysophyte sequences (Ochromonas species) only possessed the
cysteine in the GAF domain, suggesting that these phytochromes bind a different chromophore and have different light-sensing abilities (FigS12).

<figure><img src="image_19.png" /><figcaption>Figure 5 Phylogeny of the Stramenopile phytochromes. Full-length sequences were aligned and the tree was rooted with fungal and bacterial phytochromes (collapsed clade). Taxa colors correspond to the algal groups the genes belongs to, with a red background for viral sequences, and purple circles on the branches indicate local bootstraps support >0.7</figcaption></figure>

Phylogenetic tree of phytochrome full-length proteins reveals a complex story (Fig 5). As reported before (Duanmu et al., 2014; Fortunato et al., 2015), all stramenopile phytochromes seem to share a common origin, branching as sister clade to fungal phytochromes (Fig5). However, the relative position of phytochrome branches does not reflect the species phylogenetic relationships. The most striking example is the relation between diatoms, brown algae and chrysophyceae. In the phylogenetic species tree, diatoms are separated into pennates species forming a monophyletic clade, and centric
species that are paraphyletic, branching as sister groups basal to pennate. Bolidophyceae form the sister group to all diatoms (Kooistra and Medlin, 1996; Kessenich et al., 2014), but in our search of phytochrome, Bolidophyceae do not possess any phytochromes. Brown algae diverged much earlier in the history of Ochrophyta. Pennate phytochrome sequences form a monophyletic clade, sister to chrysophyceae and multicellular brown algae phytochromes; the resulting group (pennate-brown algae-chrysophyceae) being sister to centric phytochromes. Interestingly, some sequences from brown algae viruses are branching just outside the Pennate+brown algae+chrysophyceae group (see red background in Fig5). This latter point could suggest that virus-mediated gene transfer may have also contributed to phytochrome evolution amongst ochrophytes. The ancestor of pennate and centric diatom could have acquired phytochrome, which then diverged into centric and pennate branches. Brown algae and chrysophyceae would have acquired phytochromes from a pennate species, possibly through viral horizontal gene transfer.

Another interesting point is the phytochrome copy number (FigS11). From first studies (Fortunato et al., 2016) and further investigation of this omic dataset, it is clear that not all Ochrophyta possess a phytochrome gene. Even in diatoms, within both the pennate and centric diatoms, there are species which do not possess this putative photoreceptor. At present, we have not been able to find a clear explanation linking the presence/absence of the phytochrome and the species that possess it. Multiplication seems also to have occurred several times in different algal lineages. In the centric diatoms, there seem to be only one phytochrome gene copy, while pennate species have up to 4 different phytochromes suggesting that gene duplication occurred after separation between centric and pennate diatoms. Moreover, brown algae and Raphidophyceae also have several phytochrome genes (Fig5, FigS13). As suggested already by previous analyses (Fortunato et al., 2016; Osuna-Cruz et al., 2020), it seems that gene duplication occurred in benthic species preferentially (Seminavis and Amphora especially). Because these environments are heterogeneous, characterized by sharp and sometimes dynamic gradients of light, oxygen, nutrient availability, and redox state (Stockdale et al., 2009; Cartaxana et al., 2016; Marques da Silva et al., 2017), it is possible to speculate that phytochrome gene duplication represented an adaptation to thrive in this complex environments. Considering the huge biodiversity of diatoms and the important amount of
novel information deriving from environmental genomics, it will be interesting to further
investigate phytochrome distribution in different diatom species, and try to associate it
to their environmental distribution and diversity of life style and strategies.

The origin of stramenopile phytochromes remains puzzling. When compared to other
phytochromes, there is clearly a stramenopile specific phytochrome branch, sister to
fungal phytochromes, but separated from other eukaryotic algal phytochromes
(Archaeplastida and Cryptophyceae) by bacterial phytochromes. This would suggest a
common origin for stramenopile phytochrome. However, within the stramenopile
phytochromes, the phytochrome phylogenetic tree does not follow the phylogenetic
species tree. One possibility could be acquisition of phytochrome in one ochrophyta
species, then within-ochrophyta horizontal gene transfer through viral infection.

CONCLUSIONS

In this study, we have addressed the presence of different photoreceptors types in
Ochrophyta. We have shown that the LOV-based photosensing seems to be very
important in this eukaryote branch, with Aureochrome photoreceptors present in all
Ochrophyta. Moreover, we also discover a Stramenopile-specific group containing a single
LOV domain as potential ubiquitous photoreceptors in diatoms, while phyla-specific
proteins appeared in some algal groups. However, whether all these photoreceptors
evolved from a unique ancestral protein has not been assessed here, but could be done in
an other study focusing on the phylogeny of the LOV domain. We also uncovered new
domain combinations, for example with cytoskeleton-interacting domains, with possibly
new functions for LOV-based photoreceptors. Members of the cryptochrome-photolyase
family are also present in all species studied here, and among Ochrophyta, diatoms show
gene multiplication in several CPF classes (CryP, Cry-DASH and class II CPD photolyase).
In particular, we have been able to show a multiplication of Plant-like cryptochrome in
diatoms with the potential presence of new photoreceptors (CRYP2) and duplication
associated with potential specific subcellular localization (Cry-DASH and class II CPD
photolyase).
The conservation of phytochromes and rhodopsins is much more sparse on the Ochrophyta species tree. Interestingly, each of these proteins can be found in giant virus genomes (phycodnaviridae). There are several arguments in favor of a viral transfer for both rhodopsins and phytochromes. Rhodopsins are found in algal viruses, including Chlorophyta and coccolithophore viruses, and transfer of rhodopsin genes between virus and host have been demonstrated in other algae (Needham et al., 2019; Rozenberg et al., 2020). For phytochromes, the presence of a copy of a giant phycodnavirus genome carrying a phytochrome in the *Ectocarpus* genome is a good argument for virus-mediated horizontal gene transfer (Cock et al., 2010). Giant phycodnavirus are known to infect brown algae (McKeown et al., 2017), raphidophytes (Ogura et al., 2016) and pelagophytes (Moniruzzaman et al., 2014), and have been suggested to infect chrysophytes too (Endo, 2020), but not diatoms. These can be infected by small ssDNA and ssRNA viruses, but the giant phycodnaviridae are thought to be too big to get through the pores of the frustule (Tomaru and Nagasaki, 2011). Interestingly, if we focus on pennate diatom phytochromes, we can see that these genes are sparsely distributed on the diatom phylogenetic tree, but the genes form a clade on a phytochrome phylogenetic tree. This could mean that the ancestor of pennate diatoms had a phytochrome, but that it was repeatedly lost in the current lineages. However, we also noticed that some pennate diatoms that carry phytochrome are benthic species, living in dense sea floor communities, where viral pressure is high (Stal et al., 2019; McMinn et al., 2020). Viral-mediated gene transfer could therefore be very successful in this environment, and bring phytochrome and heliorhodopsin genes to these species. On this basis, one can infer that pennate diatom phytochromes form a clade not because they come from the same diatom ancestor, but because these diatoms share the same environment and are infected by the same viruses. Mining the available giant DNA virus databases (Moniruzzaman et al., 2020; Schulz et al., 2020) for rhodopsins and phytochromes genes and comparing them to Ochrophyta genes will be the next step in this analysis.

Our database is composed of a wide range of Ochrophyta species. Because our search is based on known characterized photoreceptors domains, it is nearly certain that other photoreceptors types exist and could be identified by enriching this search with additional genomic data now available from algae (Nelson et al., 2021), or environmental
sequences derived from meta-omic data and single cell genomes (Carradec et al., 2018; Delmont et al., 2020). It would be also interesting to add to this search dinotoms (dinoflagellates that possess a chloroplast of diatom origin) to identify conserved photoreceptors and eventual transfers from diatoms to the dinoflagellate host genome (Hehenberger et al., 2016). Finally, it would be informative to look more into species characteristics of the different ochrophyta groups possessing different photoreceptor classes, such as trophic mode and habitat (freshwater or marine, planktonic or benthic), and identify putative blind spots in our analysis.

Finally, representing the first comprehensive study of the photoreceptor repertoire in an important group of eukaryotes that have so far been largely ignored in photobiology research, this study provides important support for the discovery of new photosensitive proteins and for the study of their evolution and functional diversification in aquatic environments.

## MATERIAL AND METHODS:

### Database:

All transcriptome and genome resources used in this analysis are listed in Supplementary Data 1. We used only genomes with protein predictions and assembled transcriptomes. Protein prediction was performed with Transdecoder (version 5.5.0, default settings) on assembled transcriptomes if needed.

### Photoreceptor search (summarized in Table1):

We used the hmmsearch algorithm (HMMER 3.2.1) to retrieve candidate sequences. HMM model details are as follow:

**LOV:** the LOV domain is recognized by the PAS_9 model in Pfam (PF13426), however this model also recognizes other domains (lacking the flavin binding site notably). To define a specific LOV domain signature, we aligned all LOV-containing sequences from Glantz et
al, 2016 to the PAS_9 model with hmmalign (--trim option) and constructed LOV hmm profile with hmmbuild.

CPF: We tested Pfam models PF and PF models, but a custom model recognized more sequences. This custom model was built by aligning reference sequences from COG0415 (ncbi conserved domain database) with mafft (v7.247) and built with hmmbuild.

Rhodopsins: Pfam models PF01036 (bac-rhodopsin) and PF18761 (heliorhodopsins) have been used.

Phytochromes: Pfam model PF00360 (PHY) has been used to retrieve full length sequences, as defined by sequences with GAF and REC domains, aligned with mafft). Alignment of the full-length sequence has been used to build stramenopile specific models for each domain (GAF, PHY, HisKA, HATPase, REC).

<table><caption>Table 1. Details of the parameters and reference sequences used in our photoreceptor search</caption><thead><tr><td>Photoreceptor type</td><td>HMM domain</td><td>SSN alignment score</td><td>Reference sequences</td></tr></thead><tbody><tr><td>LOV</td><td>custom (PF13426 and reference sequences from Glantz 2016)</td><td>40 and 60</td><td>(Glantz et al., 2016) and haptophyte RGS-LOV sequences reported by (Coesel et al., 2021)</td></tr><tr><td>CPF</td><td>custom (alignment of the COG0415 sequences from ncbi conserved domain database</td><td>90</td><td>(Vicedomini et al., 2022)</td></tr><tr><td>Rhodopsins</td><td>PF01036 (bac-rhodopsin) and PF18761 (heliorhodopsins)</td><td>20</td><td>(Govorunova et al., 2017; Pushkarev et al., 2018) enriched in eukaryotic sequences from Interpro for the phylogenetic trees</td></tr><tr><td>Phytochromes</td><td>PF00360 and custom from Stramenopiles sequences</td><td>80</td><td>(Fortunato et al., 2016)</td></tr></tbody></table>

Candidate sequences were clustered at 90% identity with Cdhit (http://weizhong-lab.ucsd.edu/cdhit-web-server/cgi-bin/index.cgi) and submitted to Sequence similarity network (SSN) at EFI/EST (https://efi.igb.illinois.edu/efi-est/, fasta option) (Zallot et al., 2019). SSN alignment scores were chosen to separate proteins at about 40% identity and
the score has been increased if necessary. We proceeded with trial and error to find
optimal alignment scores.

Reference sequences were added to the SSN (see Table1) to help to visualize known
photoreceptor types.

Other domain examination and sequence visualization was done in CLCworkbench 8.0.1
(Pfam database version 33, built-in hmmer algorithm version 3.1b1).

Phylogenetic reconstruction:

Alignment of selected proteins was done with muscle (v3.8.31), except for CPF, where
sequences were aligned to a reference alignment of the DNA photolyase and FAD-binding
domains with hmmalign with the --trim option. Alignements were trimmed with trimAL
(v1.2) (80% conservation), and tree was reconstructed with FastTree (Price et al., 2010)
(version 2.1.11), default mode (JTT+CAT; local support values: Shimodaira-Hasegawa test
with 1000 resampling).

For species phylogeny, we used the mutligene method from Dorell et al, 2021: 63 genes
from *Phaeodactulym tricornutum* were used as blastp queries to identify homologs in each
strain of our database (best blast hit for each strain was used). Genes were aligned
individually with mafft, trimmed with trimAL (-gt 0.2) and alignments concatenated. Tree
was reconstructed with FastTree.

Trees were visualized with iTOL v6 (https://itol.embl.de, (Letunic and Bork, 2021).

Complementary analyses:

Sequences logos (aureochromes binding site and CryP family) were done at
https://weblogo.berkeley.edu/logo.cgi. Protein targeting predictions were done with
HECTAR (Gschloessl et al., 2008), diatom sequences only, at http://www.sb-
roscoff.fr/hectar/. Isoelectric points were calculated with the online tool IPC2
((Kozlowski, 2021), http://ipc2.mimuw.edu.pl). Transmembrane prediction for the
rhodopsins were done with TMHMM 2.0 at
https://services.healthtech.dtu.dk/service.php?TMHMM-2.0.

Structure homology modelisation was performed with SwissProt ((Waterhouse et al., 2018), https://swissmodel.expasy.org).

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