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SUPPLEMENTARY FIGURES

<figure><img src="image_20.png" /><figcaption>Figure S 1. Stramenopile species tree, as in Fig1, FigS3, S5 and S13 with the details of species names. Bold labels indicate genomic data, circles indicate local bootstraps support >0.7.</figcaption></figure>
<figure><img src="image_21.png" /><figcaption>**Figure S 2.** Bioinformatic analysis of the putative LOV-containing photoreceptors. A Alignment of selected sequences from LOV-containing Ochrophyta proteins, with references LOV containing proteins (Arabidopsis phototropins and known aureochromes from *Ectocarpus siliculosus* and *Phaeodactylum tricornutum*). The cysteine required for chromophore interaction is marked with a red spot; the position suggested to prevent photoreceptor activities in Aureo2 proteins is marked with a green spot. Shades of blue indicate sequence conservation. B and C Sequence similarity network of the LOV candidate proteins. Colors indicate the network connectivity, i.e to how many sequences each sequences is linked. This helps to detect the different groups B, alignment score 40; C alignment score 60 performed on the sequences in the large group in B.</figcaption></figure>
<figure><img src="image_22.png" /><figcaption>Figure S 3. LOV-based photoreceptors families in Ochrophyta. A. Repartition of the LOV families on the Stramenopile tree of species. Outer circles represent the presence of the different photoreceptor types and the bar height is proportional to the number of copies. Except for aureochromes and the Stramenopile LOV group, all other LOV families appear specific to one branch of algae. B, domain architecture of the LOV-containing proteins.</figcaption></figure>
<figure><img src="image_23.png" /><figcaption>Figure S 4. Analysis of the aureochrome and helmchromes. A Unrooted tree of the aureochromes photoreceptors. B, DNA-binding motif of the bZIP domains of Aureochrome 1a, 1b/c and 2 clades. C. Tree of the helmchrome sequences (RGS-LOV alignment). In A and C, label colors correspond to the algal groups the genes belongs to, and purple circles on the branches indicate local bootstraps support >0.7</figcaption></figure>
<figure><img src="image_24.png" /><figcaption>Figure S 5 Analysis of the CPF family in Ocrhophyta. A Sequence similarity network of the CPF (alignment score 90). B Targeting prediction by HECTAR on diatom CPF proteins. Proportion of sequences addressed to the organelles are prediction to the chloroplast, mitochondria or presence of a signal peptide. C. CPF repertoire visualized on the ohcrophyta tree of species. Outer circle represent the presence of the different photoreceptor types and the bar height is proportional to the number of copies.</figcaption></figure>
<figure><img src="image_25.png" /><figcaption>Figure S 6. Zoom on the CPD branch of the CPF tree presented in Fig3</figcaption></figure>
<figure><img src="image_26.png" /><figcaption>Figure S 7. Zoom on the Cry-DASH branch of the CPF tree presented in Fig 3</figcaption></figure>
<figure><img src="image_27.png" /><figcaption>Figure S 8. The 6-4 photolyases : A. Zoom on the 6-4 photolyase branch of the CPF tree presented in Fig3. B Isolecetric poitn calculated with IPC2 for oomycetes CPF1, separated per genus. Note that *Alphanomycete* CPF1 do not branch with other Oomycetes CPF.</figcaption></figure>
<figure><img src="image_28.png" /><figcaption>Figure S 9 The plant-Cry like proteins : A. Zoom on the plant Cry branch of the CPF tree presented in Fig3.
B. Isoelectric point calculated for the different type of Diatom CryP</figcaption></figure>
<figure><img src="image_29.png" /><figcaption>Figure S 10 Sequence analysis of the CryP sub-families. A Sequences logo of the FAD binding site of the different sub-families. PtCryP has been shown to bind FAD as chromophore and to be capable of a photoreduction of the FAD upon illumination with blue light (Juhas et al., 2014). Comparison of the representative motifs of CryP1, CryP2a and CryP2b shows that all three subclasses share amino acids involved in FAD binding such as R38, D66, D68 or N72, the latter one interacting with the N5 of the isoalloxazine ring of the FAD (numbers from the logo) (Zhong, 2015). Moreover, while CryP1 has a methionine at position 39 that has been shown to be essential for CPD binding, CryP2a/b have a leucine. The absence of this M39 with the absence of N71 and W78 suggest that CryP2a/b proteins cannot interact with CPDs (Tan et al., 2015).</figcaption></figure>

B. Spatial arrangement of tryptophan and tyrosine which may be involved in an electron transfer chain. CryP1 and CryP2b homology models based on the crystal structure of a bacterial class III photolyase from *Agrobacterium tumefaciens* (Scheerer et al., 2015) PDB 4u63.1. CryP2a homology models based on the crystal structure of *Arabidopsis thaliana* BIC2-CRY2 complexa (Ma et al., 2020) PDB 6k8k.1. In the CPF protein, FAD is embedded inside the protein, and transfer of electrons from the solution (photoreduction) occurs through an intramolecular electron transfer chain involving mostly tryptophan and tyrosine and is
an indispensable process for the CPF enzymatic or photoperception activity (Chaves et al., 2011). Notable
differences are observed in the amino acids that may be involved in the electron transfer chain. While W10,
W53, and W76 seem to be conserved between all CryP, alternative electron transfer pathways are observed
in CryP1 and CryP2a with conserved W or Y such as W87, W59 allowing to reconstitute in silico a complete
electron transfer chain.

<figure><img src="image_30.png" /><figcaption>Figure S 11. Repartition of rhodopsins and phytochromes visualized on the Stramenopile species tree.
Outer circles represent the presence of the different rhodopsins and phytochromes and the bar height is
proportional to their number of copies.</figcaption></figure>
<figure><img src="image_31.png" /><figcaption>Figure S 12. Analysis of the rhodopsin photoreceptor types. A. Sequence similarity network of the rhodopsin family. B alignment of the portorhodopsin sequences. The glutamate residue responsible for the spectral tuning is marked with a blue star. C. Phylogenetic tree of the proteorhodopsins identified in this study. Label colors correspond to the algal groups the genes belongs to, as in Fig4, and purple circles on the branches indicate pseudo-bootstraps support >0.7.</figcaption></figure>
<figure><img src="image_32.png" /></figure>

Figure S13 Alignment of selected phytochrome sequences from Stramenopile and reference phytochromes (Bacteriophytochrome from Deinococcus radiodurans, plant phytochrome from Arabidopsis thaliana PHYA, cyancobacterial phytochrome Synechocystis Cph1 and fungal phytochrome Neurospora crassa phytochrome 2). First part of the alignment is the potential chromophore binding site in the N-terminal part of the protein, while the second part is in the GAF. Conserved cysteine binding chromophore are highlighted in red.

SUPPLEMENTARY DATA

Supplementary Data 1: List of the strains used here and source for downloading the genome/transcriptome

Supplementary Data 2: List of the photoreceptor genes identified here. Each sheet corresponds to a photoreceptor type: "LOV", "CRY" (CPF family), "Rho" rhodopsins, "PHY" phytochrome. Protein name is the concatenation of the strain (in the form genus-species-strain) and the original protein/transcript name in the original dataset. Available on MyCore at

https://mycore.core-cloud.net/index.php/s/hD5zBsV1yhGkvfH
CHAPTER 2 : MARINE DIATOM
PHYTOCHROMES ARE SENSORS OF
DEPTH AND PHYTOPLANKTON
CONCENTRATION THROUGH SENSING
OF UNDERWATER LIGHT FIELD
VARIATIONS

Carole Duchêne¹, Jean-Pierre Bouly¹, Juan Pierella Karlusich², Julien Sellès¹, Chris Bowler²,
Maurizio Ribera d'Alcalà³, Angela Falciatore¹, Marianne Jaubert¹

1. 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

2. Ecole Normale Supérieure, PSL Research University, Institut de Biologie de l'Ecole Normale Supérieure, CNRS UMR 8197, INSERM U1024, F-75005 Paris, France

3. Stazione Zoologica Anton Dohrn, 80121 Naples, Italy

Phytochrome photoreceptors from *T.pseudonana* and *P.tricornutum* display a red/far-red
absorption spectra, which led to complex hypotheses concerning the light source that
could modulate DPH activity in the ocean, given the poor abundance of these wavelengths
in this environment. To get insights into the function DPH could play, we wanted to
determine the conditions in which DPH can be active in the environment, we aborded the
questions by different approaches. On one side, we determined the action spectra of the
DPH from the model diatom *Phaeodactylum tricornutum* thanks to the use of a reporter
system in which the fluorescent reporter eYFP is under the control of a DPH-regulated
promoter in *P.tricornutum*, allowing to build a mathematical model describing DPH
activity in a given light field. On the other side, we investigated in which environment are
found DPH-containing diatoms, which could explain the adaptive value that could confer
DPH activity. For this, we collaborated with Chris Bowler and Juan Pierella Karlusich to
dig into the Tara Oceans resources. Both approaches brought complementary and
convergent information toward the environmental impact on DPH activity.

In this study, I realized the wide majority of the work, from the setting of the experimental
conditions to the realization of fluence response curves of the action spectra, expression
analysis of different DPH-regulateed genes, and the mathematical modeling of DPH
activity based on already established equations, using different light field I modeled or
environmental data provided by Maurizio Ribera d'Alcala. I analyzed the sequences
retrieved from the Marine Atlas for Tara Oceans Unigenes (MATOU) and quantified their
respective abundance, and characterization of novel DPH recombinant proteins.

Manuscript in preparation
MARINE DIATOM PHYTOCHROMES ARE SENSORS OF
DEPTH AND PHYTOPLANKTON CONCENTRATION
THROUGH SENSING OF UNDERWATER LIGHT FIELD
VARIATIONS

Carole Duchêne¹, Jean-Pierre Bouly¹, Juan Pierella Karlusich², Julien Sellès¹, Chris Bowler²,
Maurizio Ribera d'Alcalà³, Angela Falciatore¹, Marianne Jaubert¹

1. 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

2. Ecole Normale Supérieure, PSL Research University, Institut de Biologie de l'Ecole Normale Supérieure, CNRS UMR 8197, INSERM U1024, F-75005 Paris, France

3. Stazione Zoologica Anton Dohrn, 80121 Naples, Italy

ABSTRACT

Aquatic environments are more penetrative to blue and green wavelengths while red and far-red ones are rapidly attenuated. So marine organisms are expected to have adapted their photosensing capabilities to the most abundant wavebands. However, marine diatoms possess photoreceptors of the phytochrome family (DPH) that respond to red/far-red light. To shed light on this puzzling evidence, we set up a reporter system in the model diatom *Phaeodactylum tricornutum* to determine the action spectra of PtDPH activity. We assessed that this photoreceptor responds, effectively, not only to red and far-red bands, but also to blue and green lights. Projecting PtDPH capability, quantified by the photochemical parameters determined *in vivo*, in different ocean light fields, we determined that, counterintuitively, PtDPH activity increases with depth, while being sensitive to the concentration of optically active components, as phytoplankton concentration. We then investigated the occurrence of DPH *in situ*, using the ocean wide data set from the *Tara* Oceans expedition, and found that DPH containing, and expressing, diatoms were almost exclusively present in temperate and polar regions, which experience high annual variability in phytoplankton abundance and a variable
photoperiod. We expanded the number of characterized DPH family members with
sequences found to be abundant in the open ocean. We found that their photochemical
properties are strongly conserved and similar to those of PtDPH, and that they show a
similar activation pattern in the environment. Overall, these results set a milestone in
marine photoreception, showing light sensitivity of DPH along the whole photic zone, and
open new perspectives both for DPH functioning and putative role in marine diatoms.

Keywords: Phytochrome, diatoms, light sensing, oceans

INTRODUCTION

For photosynthetic organisms, the perception of their light environment is crucial for
multiple aspects of their life, from the regulation of photosynthesis to cellular and
developmental processes such as acclimation, adjustment or synchronization of
physiology and metabolism (Briggs and Spudich, 2005). Photosensing relies on
photoreceptor proteins that perceive particular bands of the light spectrum (Möglich et
al, 2010). Among them, the phytochromes are red/far-red light sensing proteins, present
in terrestrial plants but also in photosynthetic and non-photosynthetic bacteria, fungi and
diverse algae such as streptophytes, prasinophytes, glaucophytes, cryptophytes and
heterokonts (Butler et al, 1959; Hughes et al, 1997; Yeh et al., 1997; Bhoo et al, 2001;
Giraud et al, 2002; Blumenstein et al, 2005; Froehlich et al, 2005; Falciatore and Bowler,
2005; Rockwell et al, 2014). Phytochromes are characterized by their ability to
photoreversibly convert between an active and an inactive state: upon the perception of
red light, the red-absorbing form Pr switches to a far-red absorbing form, Pfr, which can
subsequently revert back into the Pr form upon far-red light (Butler et al, 1959). Thus, the
activity level of phytochrome is related to the photoequilibrium formed upon a given light
exposure (Morgan and Smith, 1976). A well-known example of phytochrome
photoequilibrium dependent response in plants is the perception of the low R/FR ratio
under a canopy, allowing the detection of neighbors (i.e., competing phototrophs) and
mediating shade avoidance responses (Casal, 2012). The photoequilibrium state as well
as the phototransformation rates between the two forms can be predicted mathematically
at any wavelength, based on the phytochrome photoconversion cross sections ($\sigma_{\lambda}$) (that
depend on the quantum yield of photoconversion ($\phi$) and the extinction coefficient of each
form ($\epsilon_{\lambda}$)), and the spectral fluence rate of a light source (Butler, 1972). These
photochemical properties can be estimated from measurements made *in vitro* on
recombinant phytochromes (Giraud et al, 2010) but also *in vivo* (Schmidt et al., 1973).
However, the determination of action spectra for specific biological responses that reflect
phototransformation reactions (Hartmann 1966, Beggs, 1981, Shinomura et al, 1996)
have shown some differences between *in vitro* and *in vivo* approaches due to distortions
of the light transmitted within cells (Kazarinova-Fukshansky, 1985; Kusuma, 2021)
or/and biological events acting downstream of phytochrome photochemical reactions
(Klose, 2015; Olson, 2017).

Phytochromes are dimeric chromoproteins which sense light via heme-derived linear
tetrapyrroles as chromophore (Quail, 1991; Rockwell et al, 2006). These chromophores
covalently bind to a conserved cysteine residue in the N-terminal photosensory module
(PSM), composed of PAS (Per-Arnt-Sim), GAF (cGMP phosphodiesterase/ adenylyl
cyclase/FhlA) and PHY (phytochrome-specific) domains (Montgomery and Lagarias,
2002; Karniol et al, 2005; Lamparter et al, 2002, Blumenstein et al, 2005). Upon excitation
by light, changes in the chromophore generate protein conformational changes which
activate a more diversified C-terminal output module initiating a signaling cascade
(Rockwell and Lagarias, 2006).

The presence of phytochromes in marine phytoplankton was quite unexpected
because water differentially absorbs light bands in a manner inconsistent with
phytochrome activation. For example, light intensity decreases exponentially with depth,
and short (UV) and long (red and far-red) wavelengths of the spectra are attenuated more
strongly than green and blue wavelengths (Mobley, 1994). Additionally, the presence of
other components in the seawater, such as phytoplankton, and their pigments, inorganic
particles and colored dissolved organic matter (CDOM) also alter the light field because
they absorb and/or scatter specific wavelengths depending on their biooptical properties
(Kirk, 2011). These properties give rise to a strong vertical structuring of light in the ocean
with a continuum of spectra, going from a high intensity solar spectrum near the surface to a dim blue-green light field at depth, therefore progressively deprived of the red-far red band. Absorption spectra variations have been shown for some algal phytochromes from prasinophytes, glaucophytes and brown algae, that are tuned to the shorter wavelengths that are more abundant in the water column, with orange-yellow/far-red, red/blue or far-red/green photocycles, respectively, which has been interpreted as an adaptive spectral tuning of the photoreceptors (Rockwell et al, 2014).

Among marine phytoplankton, diatoms are one of the most abundant and diverse groups, and are considered responsible for about 20% of the primary production on Earth (Field et al, 1998; Malviya et al, 2016; Pierella Karlusich et al, 2020). These secondary endosymbiosis-derived microalgae have been able to colonize very different aquatic niches. They particularly thrive in turbulent environments (Margalef, 1978; Esposito et al, 2009), that can imply drastic variations in light conditions, both in term of intensity and quality. Diatoms exhibit a palette of photoreceptors including blue light photoreceptors of the Cryptochrome/photolyase family (CPF), the stramenopile-specific Aureochrome family, and putative green light photoreceptors of the Heliorhodopsin family (Jaubert et al, 2022). Some diatoms also possess phytochromes (DPH), similar to the bacterial class, with a conserved photosensory module followed by an output domain composed of an histidine kinase/ATPase and REC (response receiver) domains (Bowler et al, 2008; Fortunato et al, 2016).

The DPH from the model diatom species *Thalassiosira pseudonana* (TpDPH), from the centric group, and *Phaeodactylum tricornutum* (PtDPH), from the pennate group, were both shown to be able to bind as chromophore biliverdin (BV), the bilin bound to bacterial and fungal phytochromes (Bhoo et al, 2001; Blumenstein et al, 2005). TpDPH and PtDPH both display absorption spectra of the Pr and Pfr maximally peaking at 686 and 700, and 764 et 750 nm, respectively, which are red-shifted in respect to canonical plant phytochromes and other algal phytochromes (Fortunato et al, 2016).

Attempts to understand the function of this photoreceptor in *P. tricornutum* showed that exposure to far-red light leads to expression changes of a set of genes. These patterns are lost in PtDPH knockout (KO) mutants. Most of these genes are of unknown function, or
encoding proteins involved in transcriptional regulation and signaling (Fortunato et al, 2016). Hypotheses have been proposed about a role of these phytochromes as depth sensors, given the light spectrum differences along the water column, or as sensors of phototrophic cells in the surroundings, during algal blooms in particular, through the sensing of chloroplast fluorescence (Fortunato et al, 2016). However, up to now, the physiological function algal phytochrome remains a mystery, and the presence of red/far-red photoreceptors in marine diatoms that are exposed to a mostly blue-green light environment is still puzzling.

In this work we went beyond the bias of DPH being activated only in the red-far red bands and explored over the whole spectrum the response of this unique photoreceptor.

We first characterized the *in vivo* sensing properties of PtDPH, using an ad hoc designed reporter system which allowed to monitor and quantify PtDPH activation in *P. tricornutum* over all the wavelength range available in the ocean surface layer. We then modeled the PtDPH activation state in various computed or real marine light fields, to infer its *in situ* response. We additionally investigated the occurrence and distribution of DPH in *Tara* Oceans expedition data, characterized photochemical properties of a suite of novel DPHs from different ecological niches and different taxa, and verified similar properties of those of the model systems previously analyzed. In brief, the interplay between the varying underwater light field and the highly specific spectral properties of DPH produces an unique vertical pattern in the photoreceptor response. Altogether, these results permit new perspectives both for DPH functioning and for its putative role in marine diatom biology.

## RESULTS

### In vivo action spectra of DPH

To monitor DPH activity *in vivo*, we introduced in wild-type (WT) and KO-PtDPH *P. tricornutum* strains the coding sequence of the enhanced Yellow Fluorescent Protein
(eYFP) under the control from a promoter of a PtDPH-regulated gene (Hsf4.6a)
(ProHSF4.6a::YFP WT and ProHSF4.6a::YFP KO, respectively) (Fortunato et al. 2016)(Fig.
1A). Reporter strains were grown in continuous green light to avoid diurnal regulation
and because green light is the least absorbed by PtDPH in vitro.

<figure><img src="image_33.png" /><figcaption>Figure 1. PtDPH activity reporter system and experimental conditions. A. Schematic illustration the PtDPH activity reporter system. B. Kinetics of the eYFP induction in different reporter strain backgrounds: P. tricornutum wild-type (WT), knockout PtDPH mutants (KO), or non mutated cells originated from the same transformed colony than the KO (WT (Tc)(Transformation control)). Cells were grown in continuous green light (22 µmoles of photon/m²/sec) and exposed for 10 min to 800-nm far-red light (15 µmoles of photon/m2/sec) at t0 then left in the dark («10min far-red») up to 9 h post-irradiation, or directly transferred from green to darkness («Dark»). eYFP signals were measured by flow cytometry and normalized to t0.</figcaption></figure>

To verify the PtDPH-dependent control of reporter gene expression, ProHSF4.6a::YFP WT, ProHSF4.6a::YFP WT (Tc) (transformation control, i.e., cells originated from the same transformed colony than its corresponding KO but not having undergone PtDPH gene editing as described in Fortunato et al, (2016)), and ProHSF4.6a::YFP KO lines were submitted or not to a far-red (800 nm) light exposure of 15 µmoles of photon/m²/sec for 10 minutes and then left in the dark. The YFP signal was measured by flow cytometry at different time points. As shown in Fig. 1B, YFP signal increased over 6h in the dark in ProHSF4.6a::YFP WT and remained constant up to 9h post-irradiation. On the contrary, it remained at a basal level when the ProHSF4.6a::YFP WT lines were not exposed to far-red light over the same period. Moreover, eYFP induction was not observed in ProHSF4.6a::YFP KO lines submitted to the same far-red light exposure (Fig. 1B). These results indicate that our reporter system specifically reports DPH- and FR- light-dependent inductions, and is thus suitable for the characterization of *in vivo* DPH activity.
Using the same experimental setup as described above, fluence rate response curves were then generated from reporter lines submitted for 10 min to a gradient of different monochromatic lights (Fig. S1A) from the far-red to violet-blue regions (850, 810, 765, 740, 730, 470, 430 and 405 nm (Fig. S1B)). As shown in Fig. 2A, for all the colors tested, the curves exhibited the same sigmoidal shape, with no response measured at the lowest light intensities, followed by a log-linear phase as intensity increases, before reaching a plateau when light intensity is saturating (Fig. 2A). Notably, photoinduction of YFP was effective in the blue band and even up to 850 nm, a wavelength at which the recombinant PtDPH protein barely absorbs. These responses were all DPH-dependent, as the reporter ProHSF4.6a::YFP KO lines showed no induction of YFP at any wavelength (Fig. S2). Furthermore, the same fluence rate response curves performed with the light-regulated promoter *LHCF2* and the constitutive promoter *H4* (Siaut et al, 2007), showed no wavelength- or intensity-dependent induction (Fig. S2). The induction of expression in saturating blue as well as in far-red light was also verified by RT-qPCR for *HSF4.6a* and other PtDPH-regulated genes previously identified in Fortunato et al (2016) (*Phatr3_J15138*, *Phatr3_J46431*, *Phatr3_J18096*, *Phatr3_J45662*) (Fig. S3). On the contrary, no induction occurred in *PtDPH* KO lines. This loss of induction by blue light in the KO lines was shown to be specifically regulated by PtDPH because blue light-mediated induction of expression of the *Phatr3_J18180* gene (encoding LHCR7 fucoxanthin chlorophyll a/c light-harvesting protein and not found regulated by PtDPH), was still observed in KO as well as in WT lines (Fig. S3).

The log linear phase of each curve occurs at different intensities depending on the light used. It extends to the far-red (765nm) from 0.1 to 10 µmol photon/m²/s, with an intensity needed to reach half the maximum YFP (I₁/₂) of 1.47 µmol photon/m²/s. Surprisingly, the log linear phase in blue (430 nm) is shifted to lower intensities, from 0.01 to 1 µmol photon/m²/s, with an I₁/₂ of 0.3 µmol photon/m²/s while PtDPH absorbs less in the blue than in the far-red light.
<figure><img src="image_34.png" /><figcaption>Figure 2. Action and inhibition spectra for PtDPH-mediated responses. A. Action spectra of PtDPH (via activation of the YFP reporter) in response to different light colors and intensities. Cells were grown in continuous green light and then exposed to different intensities of monochromatic light for 10 min and the YFP signal measured after 6h in the dark, as in Fig.1. B. YFP signal at saturation, determined from the 4 highest intensities in A, and superimposed to PtDPH absorption spectra (red lines, Pr, dark red line, Pfr (Fortunato et al, 2016)). Values correspond to normalized means +--sd from 4 independent WT strains (WT and WT (Tc) (Transformation control). C. Inhibition spectra of PtDPH, as photoreversibility of the far-red-induced response. Cells were grown in continuous green light, then treated to saturating 765-nm LED for 10 min, and then exposed to different intensities of monochromatic LED or dark. In A and C, the YFP signal was averaged from the measure of 29700 cells by flow cytometry, and normalized between YFP signal of cells unexposed to light (=0) and exposed to saturating far-red light (765 nm) (=1). Colored lines in A and C reflect the fitting of the normalized YFP for each wavelength as function of the light intensity (Supplemental method).</figcaption></figure>
The levels of YFP reached by the ProHSF4.6a::YFP WT lines in saturating light are different for each wavelength (Fig. 2A). These levels plotted as a function of wavelengths fit with the ratio of absorbance of Pfr compared to Pr (Fig. 2B). This suggests that the observed responses are primarily regulated by PtDPH under our conditions, and that the level of YFP under a given light reflects the photoequilibrium between the active and inactive forms of PtDPH *in vivo*. Fluence rate response curves for the inhibition of YPF induction were also generated, by exposing reporter strains first to 10 min of saturating far-red light (800nm) followed by 10 min of an intensity gradient of 5 different monochromatic LEDs (730, 690, 630, 520, 405 nm (Fig. S1B)) (Fig. 2C). For all these wavelengths, the same sigmoidal-shaped fluence rate response curves, in an opposite orientation to the induction experiments, were obtained, clearly showing that PtDPH-activated responses can be photoreversed by red, but also by green and violet wavelengths (Fig. 2C).

Altogether, these results indicate that, *in vivo*, PtDPH is a photoreversible photoreceptor, not only sensitive to red and far-red light but also effectively responding to blue and green wavelengths. This implies that the integration of the full absorption spectra of PtDPH as well as its photoreversibility properties are necessary to fully understand the activation of DPH in an highly variable light field strongly dependent on depth.

## *In situ* DPH activation model

Because our reporter system appeared to be a specific and quantitative marker of the PtDPH activation state *in vivo*, we then sought to construct a general model describing DPH activity *in situ*. The rate of phytochrome photoconversion depends on phytochrome photoconversion yield, extinction coefficient, and light intensity. Assuming that PtDPH acts as a dimer and that the active form is PrPr (Rockwell et al, 2006; Fortunato et al, 2016 and this study), the percentage of PrPr in a given light field can thereby be expressed by the equations previously established (Mancinelli, 1994) (detailed in Suppl methods). The ratio of photoconversion yield, η, and the photoconversion rate of phytochrome at a
wavelength $\lambda$, $k_\lambda$ (corresponding to the sum of the two kinetic constants for activation and inhibition of PtDPH), were determined by exploiting the action and inhibition fluence rate response curves generated.

To estimate the *in vivo* ratio of photoconversion yield, $\eta$, we plotted the normalized YFP signal of action curves at saturating light intensity, reflecting the %PrPr, as a function of the ratio of absorbance ($A_{Pr}/A_{Pfr}$) deduced from the PtDPH recombinant protein absorption spectra (Fig. 3A). We also enriched this dataset with normalized YFP levels of the reporter strains exposed to mixed wavelength light fields (mix of 2 LEDs) at intensities saturating the PtDPH response. By fitting the equation of photoconversion at equilibrium (Fig. 3A), we could estimate a $\eta$ of 0.98 +/- 0.092. It should be noted that, in this simple model, the photoconversion ratio integrates the photoconversion coefficient of both forms but also all the different downstream events.

<figure><img src="image_35.png" /><figcaption>Figure 3. Modelling of PtDPH activity *in vivo*. A. Estimation of the ratio of photoconversion rates from YFP signal at saturating light (from Fig. 2A) and in response to mixed wavelengths. Colors indicate the wavelength of the LED, and pie charts, the proportion the relative intensity of each LED in mixed illuminations. Line represents the fitting of the PtDPH photoequilibrium as a function of the ratio of absorption of PtDPH Pr and Pfr forms (Equation 5 in Supplementary method). Values are means +/- sd on 4 independent WT strains. B. Sensitivity of PtDPH to different wavelengths represented from the fitting done on the curves in Fig. 2A and 2C. $k_{Fit}$ is the resulting exponential constant. Mean +/- sd on the 4 independent WT strains; dashed line represents the theoretical $k_{Fit}$, calculated with the absorption spectra of the recombinant protein and the determined ratio of photoconversion yield (Supplementary method).</figcaption></figure>

With this estimation of $\eta$, we could fit exponential curves to each normalized eYFP-fluence rate response data, both for induction and inhibition experiments (Fig. 2A and 2C), and fit an exponential constant $k_{Fit}$ for each wavelength. When plotting the $k_{Fit}$ values against wavelengths (Fig. 3B), we found the inhibition and activation curves exhibit peaks in the blue and red or far-red regions, as the absorption spectra of the Pr and Pfr forms,
respectively (Fig. 3B). However, the relative height of these peaks does not fit the
theoretical ones calculated with the absorption spectra and ratio of photoconversion yield
(Fig. 3B, dashed line). In the action spectra, k at 430 nm, k<sub>430nm</sub>, was expected to be about
a third (0.3) of k<sub>730nm</sub>, but we observed it to be 4.68 times higher. The same difference is
observed in the inhibition spectra with k<sub>405nm</sub> vs k<sub>730nm</sub> (theoretical factor: 0.59, observed
factor:3.3) or k<sub>690nm</sub> vs k<sub>730nm</sub> (theoretical factor:1.4, observed factor:10.3) (Fig. 3B). A
linear relation between theoretical and observed values only reaches adjusted R<sup>2</sup>=0.4311,
but this rose to 0.6247 when ignoring the values in far-red bands (&gt;700nm). Given the
poor abundance of far-red light in the marine environment, the second relation was kept
to calculate k in a complex light field and to model DPH activity in the environment.

DPH activity in the environment

To scan DPH activation state in the ocean in a wide range of conditions, we projected the
DPH activity model established above in a large suite of modeled marine light fields, using
a relatively simplified bio-optical model. Briefly, we computed the sea surface light
spectra using the Tropospheric ultraviolet and visible radiation model (Madronich 1987).
The underwater spectrum at any depth depends on the sea surface light irradiation, depth
and the spectral diffuse attenuation coefficient of the water environment K. K can be
modeled as the sum of the absorption spectrum of the water (Kw), the attenuation
coefficient of the phytoplankton (Kph) and the attenuation coefficient of the background
turbidity (Kbg) due to non-algal particles and dissolved organic matter. The formula and
tabulated values from Morel and Maritonema (2001) have been used to calculate light
attenuation (350-700nm) of phytoplankton in the water column for different Chl a
contents. We also added absorption by dissolved organic matter and inanimate particles
(Kbg) using Kbg(λ) = Kbg(440nm)exp(-S(λ-440nm)), with S=0.017 (Babin et al, 2003, Holtrop et
al, 2021). It is worth noting that this approach neglects transpectral processes, but allows
a rapid exploration of a large suite of scenarios. We started considering that all
parameters are homogeneous throughout the vertical column water. Very different
spectra were calculated at the bottom of the photic zone (1% of surface light intensity)
(Fig. S4A and D), with particularly varying ratios of blue to red and blue to green bands
(Fig. S4B, C, E, F) with depth, and the angle the sun light hits the water surface (Fig. S4G and H).

Projection of %PrPr at saturation - using the model developed with the fluence rate response results - showed that for Chl a concentration from 0 to 1mg/m³, the percentage of active PtDPH increases with depth, rapidly in the first approx. 20 m and more slowly going deeper (Fig. 4A). Interestingly, at the same time, red and far-red bands are strongly absorbed by water and blue light becomes dominant (Fig. S4A to F). For higher, though homogeneous, values of Chl a (1 to 5 mg/m³), % of PrPr increases compared to the surface down to approx. 20m, and then begins to decrease (Fig. 4A). The depth of this subsurface maxima decreases when Chl a increases (17m for 2mg/m³, 8m for 5mg/m³) (Fig. 4A). Because Chl a absorbs mainly blue and red bands, the increase in its concentration results in a reduced intensity predominantly of blue light (Fig. S4A to C). These results indicate that DPH activation state is thus depth- and Chl a- concentration/distribution dependent (Fig. 4A and S5C). Kbg variations, mimicking coastal (Kbg from 0.005 to 0.6 m⁻¹) and clear oceanic (Kbg around 0.003 m⁻¹) environments resulted in the same effects as increasing Chl a concentration due to the absorption of blue light by organic matter (Fig. S4D, E and F), although at low but not null Kbg values, DPH activity will slightly increases (Fig. S5A and D).

To take into account the light intensity and the sensitivity of PtDPH, we used an integration time of 10 min (as in experimental data), hypothesizing 100% of PfrPfr as initial state. This showed that PtDPH can be active even below 100 m in clear waters, while it would be active only at the surface in turbid environments (Fig. 4B and S5B).
<figure><img src="image_36.png" /><figcaption>Figure 4. PtDPH activity as proportion of PrPr formed in modeled environmental fields varies with depth, water turbidity, and solar angle. Effect of chlorophyll a (Chl a) and depth on DPH photoequilibrium at saturation (A) or after 10 min of illumination (B). C. Combined effects depicted as color of depth (Chl a), background turbidity (Kbg) and sun zenith angle on PtDPH activity, calculated for 1h of illumination. Transparence levels reflect the saturation of the photoreactions in 1h.</figcaption></figure>

To include all the components of the underwater light field in a realistic scenario, we ran
the photoresponse model using profiles measured *in situ* with light wavelengths up to 800
nm compared to the modeled ones that were only going to 700 nm. The shape of the DPH
activity profiles and Chl a-dependence were clearly close to those calculated with
modeled light (Fig. 5) while reflecting the dishomogeneous distribution of bio-optically active components and transpectral processes. These *in situ* light profiles are also examples of non-homogeneous distributions of Chl a in the water column, with the presence of a deep chlorophyll maximum. Using the environmental light profiles, results matched those obtained with the modeled light (Fig. 4B and 5). Interestingly, PtDPH activity drops below the deep chlorophyll maximum (when present) (Fig. 5).

<figure><img src="image_37.png" /><figcaption>Figure 5. PtDPH activity in *in situ* measured field fields. A. DPH activity as proportion of PrPr formed (%PrPr), at saturation (dot) and after 10min illumination (triangles), in *in situ* environmental light fields measured at different stations in the Atlantic Ocean(data gathered in Bracher et al, 2015). In red, chlorophyll a (Chl a) concentrations measured by HPLC at different depths.</figcaption></figure>

We also examined the effect of solar angle. Solar zenith angle changes with time of day, but maximal zenith angle in the day also changes with the season and the geographical position. The effect of solar angle on the underwater spectral field (Fig. S4G and H), and therefore PtDPH activity, appears to be only impacted for very low solar angles (from 80°), i.e., sunrise and sunset, or at high latitudes in winter (Fig. 4C and S5E). The exposure time was critical to determine if PtDPH would sense the spectral variation
at low solar angle. If exposure time was kept to 10min, PtDPH would not perceive the
increase of blue compared to red and green lights at the end of the day. However,
increasing exposure time to 1h, hypothesizing a fluence dependent response, would allow
PtDPH activity to increase at low solar angles (Fig. 4C, Fig. S5E). In that case, the % of PrPr
can increase from 20% during the day to 27% at dawn/dusk (Fig. S5E, at 10 m). We also
used time series light data from Veedin Rajan et al (2021), and calculated an increase of
DPH activity at the beginning and end of day of about 5 % of %PrPr variation between
middle of the day (28%+/-2.8 at noon) and dawn/dusk (33%+/-0.8 at 9pm)) (Fig. S6).

In conclusion, we show here that in environmental light spectra, PtDPH activity increases at depth in clear water, but decreases when turbidity increases; to a lesser extent, DPH can sense spectral variations at low solar angles. This behavior matches the relative enrichment of blue bands compared to red and even green bands in the light fields.

DPH biogeography and its global drivers

As DPH is not conserved in all diatoms (Fortunato et al, 2016), we conducted
biogeography studies to identify environmental factors that could influence DPH
distribution. To do so, we looked directly for *DPH* genes in the Marine Atlas for *Tara*
Oceans Unigenes (MATOU, version2) (Carradec et al, 2018).

We designed DPH-specific HMM models based on known DPH protein sequences. This
search resulted in 95 *DPH* genes attributed to abundant diatom genera such as the centric
Thalassiosirales or Cymatosirales, with 2 near-full-length proteins, while only 2 pennate
DPH genes were found (Supplementary Data, online). This is consistent with DPH not
being present in the most abundant oceanic pennate diatom genera, *Fragilariopsis* and
*Pseudo-Nitzschia* (Malviya et al, 2016). Searches in the recently produced SMAGs data
(Single cell and Metagenome Assembled Genomes) from *Tara* Oceans (Delmont et al,
2020), produced similar results with only MAGs corresponding to the Thalassiosirales or
Cymatosirales phyla possessing DPH. No other diatom MAGs possessed DPH or other
types of phytochrome-derived photoreceptors (i.e., neochromes or other combinations of
domains).
<figure><img src="image_38.png" /><figcaption>Figure 6: DPH distribution is linked to latitude, temperature and optical parameters. A. Map of the presence and absence of DPH genes in Tara Oceans sampling stations. B. Centric DPH (top panels) and Aureochrome (Aureo, bottom panels) gene abundance relative to total centric diatom genes as a function of absolute latitude, from metagenomic (left panels) and metatranscriptomic (right panels) data. C. Spearman's correlation of Aureo or DPH relative abundance with different environmental parameters. D. Generalized additive models (GAM) of Aureo and DPH relative abundances with different environmental parameters ("individual gams"). E. Complex GAMs explaining DPH relative abundance with a combination of environmental parameters.</figcaption></figure>
Mapping the presence and absence of DPH in the different *Tara* Oceans stations (Fig. 6A) revealed that these genes are hardly found in tropical regions (only 2 stations with DPH) while they are present in nearly all stations in extratropical regions (absolute latitude > 30°) (Fig. 6A and Fig. S7A and B). Biogeography of the DPH-containing diatom MAGs reflected those found for DPH genes in the *Tara* Oceans metatranscriptomes, with nearly no presence detected in tropical regions (Fig. S8).

To confirm that this localization was specific to DPH, we designed HMM models specific for Aureochromes (Aureos), which are blue light photoreceptors ubiquitous in diatoms. 1305 genes were found as diatom Aureos in the *Tara* Oceans metatranscriptomes of which 817 belonging to centric species. When mapping the presence/absence of Aureo genes in *Tara* Oceans stations, Aureos were found to be ubiquitous (Fig. 6B and Fig. S7C and D).

Given the low number of pennate DPH sequences, we performed a quantitative analysis of DPH distribution only with centric genes. The abundance of centric DPH reads was normalized against total centric reads from the MATOU2 gene set. As shown in Fig. 6B (and Fig. S9), the relative abundance of DPH sequences increases with absolute latitude, while Aureo abundance did not exhibit any particular latitudinal pattern, suggesting a specific biogeography of DPH containing diatoms. To decipher the impact of environmental conditions on DPH biogeography, we used relative gene abundance to perform Spearman's correlations with a set of environmental variables (Fig. 6C and Fig. S10A). These were a mix of parameters measured *in situ* (available at PANGEA, Pesant et al., 2015), but also seasonality indexes derived from the World Ocean Atlas (nitrate concentrations and temperature annual variations) and from satellite measurements (annual variation of chlorophyll a concentration, downwelling attenuation coefficient Kd, dissolved organic matter (Ardyna et al, 2017)). In agreement with previous analyses (Fig. 6A and B), a strong positive correlation was detected between the relative centric DPH abundance and the absolute latitude, and a negative correlation with temperature, salinity and conductivity (Fig. 6C et Fig. S10A). Strong positive correlations were found with Chl a concentrations, total carbon and other related *in situ* optical parameters such as backscattering at 470 nm, fluorescence, particle backscattering coefficient or beam attenuation coefficient at 660 nm, and a negative correlation was found with depth of the
euphotic zone (Fig. 6C and Fig. S10A). DPH abundance also positively correlated with seasonality indexes related with Chl a and light. However, DPH relative abundances were not strongly linked to nutrient concentrations (no correlation with NO3, PO4, Si, small with Fe). These tendencies are common to metatranscript reads and metagenomic reads, and are conserved when separating samples by filter size (Fig. S10A). Aureo relative abundance also correlated strongly with *in situ* optical parameters, but not with their annual variations. Interestingly, Aureo relative abundance in metatranscriptomes showed positive correlations with nutrients (especially NO3 and PO4) and CO2 as well as a negative correlation with pH (Fig. 6C and Fig. S10A). Such a pattern was largely absent in metagenomes. This could suggest regulation of Aureo transcription by nutrients, contrary to what is observed for DPH.

To go further with this analysis of DPH biogeography and capture non-monotonous relations between DPH abundance and environmental parameters, we performed "individual" generalized additive models (GAMs, DPH abundance explained by one environmental variable). Individual GAMs analysis summarized the above correlation analysis, but detected also additional relations, such as with iron concentration and sunshine duration (Fig. 6D and Fig. S10B). More complex GAMs were performed by selecting environmental variables which were found with high explanatory power in the individual GAM analysis and did not correlate with each other. We ended with 4 variables: *in situ* temperature, total carbon (which correlates with angular scattering coefficient at 470nm and Chl concentration in the water), sunshine duration, and annual variability of Chl a concentration. Together, these parameters could explain 84.7% to 96.6% of the deviance in metagenomes and metatranscriptomes, respectively (Fig. 6E, Fig. S11). The analysis done with the same environmental variables using Aureo genes explained only little deviance (14.2 and 25.1% in metatranscriptomes and metagenomes), with a many non-significant terms (only temperature was found to be significant).

These analyses indicate that DPH can be found in open ocean regions, especially in temperate and polar regions where high variations of primary productivity occur.
Photochemical properties of DPH are strongly conserved

TpDPH and PtDPH absorption spectra are so far the only 2 examples of DPH absorption
spectra, and both exhibit peaks in red and in far-red bands, characteristic of
bacteriophytochromes (Fortunato et al, 2016). To determine whether these features were
widely conserved among DPH or if there can be spectral tuning in some species, as seen
in other algal groups (Rockwell et al, 2014), we produced recombinant versions of
additional DPH photosensory modules (PSM), by co-expressing in *Escherichia coli*, PSM
sequences together with the *Synechocystis* Heme Oxygenase gene, encoding the enzyme
producing biliverdin, and purified the recombinant proteins. Four sequences were cloned
from different distantly related diatom species, with distinct ecological niches, isolated at
different locations around the world (Table S1). Moreover, DPH from two synthetic genes
from sequences identified in *Tara* Oceans dataset were also produced (Table S1).

<figure><img src="image_39.png" /><figcaption>Figure 7. Spectral properties of DPH from various species are conserved. A. Absorption spectra of far-red light illuminated (red line) and red light illuminated (darker red line) recombinant photosensory domains of DPH expressed with biliverdin as chromophore. B. Differential absorption spectra between red- and far-red- illuminated DPH. *Acor*, *Arcocellulus cornucervis*; *Ccry*, *Cyclotella cryptica*; *Mini-like*, *Minidiscus-like sequence*; *Mpol*, *Minutocellus polymorphus-like sequence*; *Sbio*, *Shionodiscus bioculatus*; *Scos*, *Skeletonema costatum*; *Tpse*, *Thalassiosira pseudonana*.</figcaption></figure>
<table><caption>Table 1: Spectral and photochemical properties of different recombinant DPH</caption><thead><tr><th>Phytochrome</th><th>max ΔAin B</th><th>min ΔA in B</th><th>max ΔA in R</th><th>min ΔA in FR</th><th>η</th></tr></thead><tbody><tr><td>Acor</td><td>385</td><td>424</td><td>682</td><td>766</td><td>0.65</td></tr><tr><td>Ccry</td><td>383</td><td>424</td><td>678</td><td>767</td><td>0.76</td></tr><tr><td>Mini-like</td><td>388</td><td>427</td><td>680</td><td>769</td><td>NA</td></tr><tr><td>Mpol-like</td><td>383</td><td>424</td><td>678</td><td>766</td><td>0.82</td></tr><tr><td>Scos</td><td>388</td><td>425</td><td>680</td><td>767</td><td>0.61</td></tr><tr><td>Sbio</td><td>385</td><td>423</td><td>678</td><td>767</td><td>NA</td></tr><tr><td>Tp*</td><td>384</td><td>425</td><td>684</td><td>767</td><td>0.68</td></tr><tr><td>Pt*</td><td>380</td><td>420</td><td>To 698</td><td>751</td><td>0.66</td></tr></tbody></table>

*absorption spectra from Fortunato et al, 2016. Max and min ΔA : maximum and minimum of the differential absorption spectra in the blue (B), red (R), and far-red (FR) bands.

η : ratio of quantum yield. NA, indicates that the ratio of the quantum yield could not be determined from the absorption spectra obtained (recombinant proteins were not pure and abundant enough, which would lead to inaccurate values).

Acor, *Arcocellulus cornucernis* RCC2270; Ccry, *Cyclotella cryptica* CCMP332; Mini-like, synthetic gene related to Minidiscus species identified in MATOU; Mpol-like, synthetic gene related to *Minutocellus polymorphus* species identified in MATOU; Scos, *Skeletonema costatum* RCC1716; Tp, *Thalassiosira pseudonana* CCMP1335; Sbio, *Shionodiscus bioculatus* RCC1991; Pt, *Phaeodactylum tricornutum* CCMP2561.

All these sequences exhibit a cysteine residue in the N-terminal extremity domain, at a conserved position with the cysteine residue found to bind the biliverdin chromophore in bacterial and fungal phytochromes (Fig. S12). All purified proteins were chromophoric and exhibit similar absorption spectra, with absorption maxima in the red (682 to 686 nm) and far-red (762 to 768 nm) light, essentially the same as previously observed in Pt- and TpDPH (Fig. 7 and Fig. S13). As already observed for Pt- and TpDPH, no thermal reversion occurred over 24h with these recombinant proteins (data not shown). We used the method from Giraud et al (2010) to estimate the absorption spectra of the pure Pfr form and estimate the ratio of photoconversion reactions (Pr→ Pfr and Pfr→Pr) of these
recombinant proteins. These latter little vary, with values from 0.61 for *S. costatum* DPH to 0.82 for the synthetic *Minidiscus*-related DPH (Table 1). It is noteworthy that the value determined for the recombinant PtDPH is pretty close to the one determined *in vivo* with the reporter system (0.66 and 0.98 respectively).

The photochemical properties of phytochrome, both in terms of absorption spectra and ratio of quantum yield, thus appear to be strongly conserved in phylogenetically and ecologically distant diatom species.

Additionally, we introduced in a ProHSF4.6a::YFP KO reporter line, the *DPH* gene of *T. pseudonana*. We observed that YFP induction in response to far-red and blue light was restored in the transformed line (Fig. S14), showing that TpDPH can complement the PtDPH KO mutant. This result indicates that TpDPH under its Pr form can activate the same signaling cascade as PtDPH.

We then used the photochemical values determined for recombinant DPH from *T. pseudonana* and *S. costatum* (Table 1), to model their activity in the marine light fields. Overall, the effects of depth, turbidity and time of day were the same as we found for PtDPH, as could be expected from the very close biophysical properties of the recombinant proteins (Fig. S15). However, TpDPH in particular seems to be more responsive to Chl a variations (Fig. S15B). In summary, here we showed that the modeled DPH activity using PtDPH can be extended to DPH of ecologically relevant species, and reveals the ubiquitous property of DPH to effectively sense blue and green lights variations along the water column, in addition to red and far-red lights. This allows ewquisite sensing of depth and chlorophyll concentrations.

# DISCUSSION

## DPHs from distant diatom species display conserved spectral and photochemical properties

Prior to the current study, absorption spectra from only two diatom phytochromes were known: from the pennate *P. tricornutum* and the centric *T. pseudonana*. Additionally, the
phytochrome from the closely related multicellular brown alga, *Ectocarpus siliculosus*, showed spectral variation with a far-red/green photocycle that was hypothesized to be linked to the intertidal and shallow sublittoral habitats of this alga (Rockwell et al, 2014). Moreover, the blue-shifted dark state of several phytochromes from marine Prasinophyte and Glaucophyte living in coastal and open waters was suggested to be an adaptation to marine blue-enriched light fields (Rockwell et al, 2014). This is clearly not the case in diatoms, as spectra of different DPHs show strong conservation of spectral properties. Even DPH from species that are evolutionary more distant (*Minutocellus* sp. and *Arcocellulus cornucervis* compared to the Thalassiosirales) or synthetized from environmental sequences, and therefore likely to be chimeric, share close photophysical properties in terms of absorption and ratio of photoconversion yield (Table1).

We also found that TpDPH can complement PtDPH function in *P. tricornutum*, at least for the phenotype studied here (far-red- or blue light-induced expression of the reporter protein). This suggests that at least the first signaling step downstream of DPH activation could be shared between *T. pseudonana* and *P. tricornutum* and that Pr could be the active form for TpDPH, and possibly also that of the other DPHs characterized in this study.

Overall, these results imply that the conserved spectral features of phytochrome among diatom species/genera must confer a specific adaptive value to the life of this group of organisms, different from those provided by phytochrome spectral tuning observed for algae belonging to other phyla.

## Blue light-enriched fields favor DPH activation

Analysis of diatom phytochrome activity and distribution has revealed several novel aspects about the adaptation of this photoreceptor to work in the marine environment. We found that *P. tricornutum* DPH-dependent responses exhibit photoreversibility, characteristic of phytochrome photoreceptors. Moreover, we also found that these responses could be elicited upon blue light exposure, with an even higher efficacy than the far-red ones, which was unexpected based on the PtDPH absorption spectra.
The PtDPH-dependent blue light induction was observed for the *HSF4.6a* gene, whose expression was monitored with the eYFP reporter, but also for other genes, previously identified as far-red light- and PtDPH- regulated (Fortunato et al, 2016) (Fig. S3). Phytochrome-dependent blue light responses have also been observed in fungi, terrestrial heterotrophic organisms possessing phytochromes similar to bacterial phytochrome ones and phylogenetically close to DPH. For instance, *Aspergillus nidulans* phytochrome FphA was shown to be a dominant photoreceptor controlling both red- and blue-light mediated gene expression, with 86% of the blue-light induced genes requiring the presence of this sensor (Yu et al, 2021). A milder role of phytochrome in blue light responses has also been observed in the fungus *Ustilago maydis* (Brych et al, 2021). On the contrary, in plants, while phytochromes also exhibit a minor absorption peak in the blue band (Butler et al, 1959; Beggs, 1981), a very limited effect of blue light has been reported for phytochrome-mediated responses (Shinomura et al, 1996). This weak efficacy of blue photons can be explained by a possible distortion of light in leaves due to the presence of absorbing photosynthetic pigments as well as scattering by cell walls (Kazarinova-Fukshansky, 1985; Kusuma, 2021).

In diatoms, despite the presence of blue light-absorbing photosynthetic pigments (i.e., chlorophyll c and fucoxanthin), a higher responsiveness to blue, compared to far-red light, has been observed for PtDPH. This behavior could suggest a local increment of blue photons actually reaching DPH within the cell. It is known from diatom species with a frustule that the light field can be modified when it goes through the silica cell wall, the valves trapping and scattering photons, blue ones in particular, resulting in an enhancement of photosynthesis (Goessling et al, 2018). However, the *P. tricornutum* cell wall is very thin, and no such light field alteration has been reported so far. Because the level of the measured PtDPH-mediated response at saturating light matches the absorption ratio of the Pfr/Pr forms, we hypothesize a reduction in the number of far-red photons reaching DPH, rather than an increase of blue photons. In addition, this also suggests that in our condition, PtDPH would be the sole photoreceptor responsible for this response, but we cannot exclude that the sensitivity to blue compared to far-red light derives from the synergistic action of other blue-light photoreceptors on the DPH-
signaling pathway, as shown for FphA (Yu et Fischer, 2018; Yu et al, 2021) and in plants
(Casal et al, 1998; Usami et al, 2004; Hughes et al, 2012).

In any case, our modeling reveals that blue light is the prominent waveband
favoring DPH activation in the marine environment. Indeed, both our characterization of
the *in vivo* PtDPH features, and the rapid depletion of the long wavebands in the water
column drive this effect. Consequently, red, as well as green light, act mostly in reverting
blue light activation. This new scenario minimizes our previous efforts to find the far-red
light sources eliciting DPH activation underwater (Fortunato et al, 2016). It also makes
the contribution of the red/far-red fluorescence emitted by the chloroplast(s) negligeable
since it represents <5% of the exciting blue light, and is therefore largely overcome by the
light applied.

DPH is an extremely sensitive photosensor in marine environments

Our projection of DPH activity in different marine environments, from clear oceanic
waters to turbid coastal environments, shows that DPHs sense both the intensity and the
ratio of wavebands. The first parameter controlling DPH activity is depth. In our model,
DPH will be less active at the surface due to a higher red-to-blue ratio, but can quickly
change its photoequilibrium with the mild spectral changes occurring in the first ~ 20
meters of the water column, as the light intensity is high and not limiting. With depth, DPH
becomes more active while the red- and green- to-blue ratio get lower, but then, its
activity becomes fluence rate-dependent, as light is limiting. Because of the dynamic of
DPH equilibrium and the effect of different wavebands on its sensitivity all along the
photic zone, DPH is predicted to be an efficient depth sensor measuring green-red/blue
ratio. At depth, in a blue-enriched environment, DPH could also mediate long-term low
light acclimation responses. DPH could also have a role in perceiving changes in the light
field, both in amplitude and spectrum, associated with vertical displacements, due to
convective events or wind mixing, both being more intense at mid-high latitudes.
Considering vertical velocities in the range of 0.01 to 0.1 m.s⁻¹, which are typical of
convection or Langmuir circulation (Moum and Smyth, 2019), in one hour cells can be
displaced by a few to tens of meters producing a change in the ratio between the two DPH
forms by a factor of more than 3.

In addition to depth, our model also shows that DPH activity increases when solar zenith
angles are low, such as at dawn and dusk, during winter, or at high latitudes. Parallel
analysis of DPH biogeography in the *Tara* Oceans dataset showed that DPH-containing
diatoms are distributed with a latitudinal pattern, in areas with strong seasonal
variations. DPHs are quasi-absent from the tropics (metatranscriptomes form only 2
stations contained DPH sequences), which is consistent with a recent analysis of
metatranscriptomic data in a tropical region (north of Hawaii), indicating an absence of
*DPH* genes (Coesel et al, 2021). This is not due to diatoms being absent in these regions,
as the Thalassiosirales, one of the most represented genera in *Tara* Oceans, are
particularly abundant in the tropical Pacific Ocean (Malviya et al, 2016). In addition, we
did find the blue light Aureo photoreceptors from diatoms in these regions. The strong
latitudinal distribution pattern observed for DPH opens interesting novel hypotheses
about the role of this photoreceptor in the detection of seasonal light variations, and
photoperiodic regulation, processes that are still largely unexplored in phytoplankton.
Interestingly, we also found that DPH activity in the environment is strongly modulated
by light field changes induced by phytoplankton concentration (i.e. Chl a concentration),
and tightly follows the displacement of the deep chlorophyll maximum. This result is
evocative of the neighbor detection response triggered by plant phytochromes (Casal et
al, 2012), though strikingly different, as in diatom it would be based on the reduction of
blue/green ratio light absorbed by the photosynthetic pigments and not in a reduction in
the ratio of red/far-red light, as in plants. DPH have been found at high latitudes where
seasonal blooms are more common than in the oligotrophic ocean (Malviya, et al. 2016).
In blooms, phytoplankton concentration increases drastically, which, according to our
model, will reduce activity of DPH. Blooms of Thalassiosirales species in late winter/early
spring, followed by Chaetoceros species (a non DPH-containing genus) have been
recorded (Caracciolo et al, 2021; Arsenieff et al, 2020). It is thus appealing to speculate
that a change in DPH activity because of increased phytoplankton concentrations could
be an end-of-bloom signal, controlling species succession and bloom timing. This
modulation of DPH activity could also play a role in the germination of diatom resting
spores (Eilertsen et al, 1995) or even sexual reproduction as a signal that cell
concentration is high enough to allow gametes to meet. This last hypothesis is particularly
appealing, as red light is known to induce sexual reproduction in the centric diatom
*Stephanopyxis palmeriana*, in a far-red reversible manner (Ren et al, 2005). Also,
phytochromes are known to be primary regulators of sexual reproduction in fungi
(Blumenstein et al, 2005). Unfortunately, no genomic and genetic tools are available for
*Stephanopyxis palmeriana*, and the transcriptome data from a related species,
*Stephanopyxis turris*, does not show DPH expression. In the future, we could potentially
test this phenotype in other diatom species expressing DPH and showing sexual
reproduction.

Finally, although the function of this photoreceptor remains unestablished in diatoms, as
in other microalgae, this study identifies critical parameters influencing the activity of
DPH in an environmental context. By completely changing perspectives on phytochrome
activity in the watercolumn, it identifies DPH as a highly sensitive photoreceptor, whose
activity would be intimately tuned to the light distribution pattern and critical biological
processes in the marine environment.

MATERIAL & METHODS

Culture conditions

Wild-type *P. tricornutum* (Pt1 8.6; CCMP2561) cells, PtDPH-KO mutants (Fortunato et al, 2016), and the reporter lines derived from them, *Thalassiosira pseudonana* CCMP1335, *Cyclotella cryptica* CCMP332, and *Skeletonema costatum* RCC1617 were maintained in Enriched Seawater, Artificial Water at 19 °C under a 12L/12D regime using 50 µmol photons.m⁻².s⁻¹ white light (Philips TL-D De Luxe Pro 950). *Arcocellulus cornucervis* RCC2270 and *Shionodiscus bioculatus* RCC1991 were grown in enhanced artificial seawater at 4°C in 12hL/12hD cycles with RGB LED panel. See also Supp Table1 for details of the strains.
Experimental conditions for action and inhibition spectra

For action and inhibition spectra cells, reporter lines were maintained in continuous green light (LEDs, 520nm, 22 µmol/m²/sec) with shaking (160 rpm) for at least two weeks before the experiments. On the day of the experiments, 2 or 4L cultures at about 1.5e⁶ cells/mL were split into 20mL in plastic culture flasks (Corning). 25 flasks per LED were then irradiated one behind the other to generate a gradient of light. For the inhibition spectra flasks were exposed in an incubator to 10min of far-red light (800nm, 10 µmol m²/sec) prior to illumination with the LED to test. The LED was turned on for 10min with no other light source then turned off, and the flasks remained in the dark for an additional 5h50min time needed to have a stable amount of eYFP/cells. Shaking was maintained during the illumination and the dark period. The same experiment was repeated for all the LEDs to test (405, 430, 470, 730, 740, 765, 810, 850nm) for the action fluence responses, and 405, 520, 630, 690 and 730 nm for the inhibition fluence responses; see figure S1 for LEDs spectra). Multi-wavelength expositions were performed in incubators equipped with Blue (450nm), Green (520nm), Red (680nm) and FR (800nm) LEDs. Light mixes were chosen based on the action and inhibition spectra results (ratio of wavebands and intensities) to cover ratio of DPH absorption not covered before, and to use light intensity supposed to saturate DPH reaction in 10min. Cultures grown in the same conditions as for the spectra and split to 20mL cultures, exposed to 10min of LED mix then transferred in the dark for 5h50min before eYFP measurements. These experiments were done twice for each strain, with two technical replicates each time.

At the end of the dark period, each flask was sampled and passed through a MAQSQuant flow cytometer. eYFP signal per cell was recorded in the B1 channel (excitation 450nm, 500-550nm emission filters). Each sample was run until 29700 events from the target population were measured, and eYFP signal measures per cell were averaged on the whole diatom population. eYFP signal was normalized between the signal from cells that went directly from green (growth) light to darkness (normalized to 0) and to cells that were exposed to 10min of FR (765nm) and then darkness (normalized to 1). This allowed us to compare different strains with different basal expressions of the eYFP.
The different equations used for the modeling of DPH activity are issued from Mancinelli (1994), and are presented in detail in Supplementary method. Analysis of the normalized eYFP signal at "saturating" light intensities was done using the 4 flasks exposed to the highest intensities for monochromatic illumination and the data from the bichromatic illuminations. For each strain, equation corresponding to the normalized YFP for each wavelength (λ) at light saturation (Equation 5 in Supplementary method) was fitted with the nls function in R, with only the ratio of quantum yield as parameter to estimate. For the analysis of the eYFP-light intensity curves, for each strain and for each LED, the eYFP signal was normalized between 0 and 1. The equation corresponding to the normalized YFP for each wavelength (λ) as function of the light intensity and time (equation (7) in Supplementary methods) was fitted to the data, with only the exponential constant k as missing parameter (Supplementary method).

## Modeled and real environmental light spectra.

We used TUV v5.3 (Madronich 1987) to compute sea-levels light spectra (direct solar and diffuse) for different sun zenith angles (downwelling irradiance only). Underwater attenuation coefficients were calculated with the formula from Morel and Maritonema (2001) and for CDOM and particles (Babin 2003; Kirk, 2021; Holtrop 2021). Light spectra were then calculated down to 100m deep for different solar zenith angles, Chl a concentration and particles content (hypothesizing that Chl a, CDOM and particles concentration would be homogeneous in the water column). Measured environmental light spectra were retrieved from various studies (Bracher et al, 2015).

## Construction of Pt-DPH activity reporter lines

The 931-nt upstream region of the *Hsf4.6a* (Phatr3_J49557) and the *eYFP:FcpAt* fragment were amplified by PCR using the Phusion High-Fidelity DNA polymerase (Thermo Fisher, USA) from genomic DNA and the primer pairs : *Hsf4.6ap_Fw0* and *Hsf4.6ap-YFP_Rv*, and from the pDEST-C-HA vector **(Erreur ! Référence de lien hypertexte non valide.)** *Erreur ! Référence de lien hypertexte non valide* and the primer pairs : *YFP-Hsf4.6ap_Fw* and *FcpAT_Rv0*, respectively (Table S2). The two fragments were assembled
by PCR using the *Hsf4.6ap_Fw0* and *FcpAT_Rv0*, and the final product cloned into pGEM-T (Promega). Transgenic lines were obtained by biolistic, by co-transformation with Phleomycin resistance plasmid as described in De Riso et al. (2009), in WT, WT (Tc) (Transformation control) and PtDPH-KO mutants obtained in Fortunato et al (2016).

RNA extraction and gene expression analysis

WT and PtDPH KO mutant lines were grown as described for the action spectra experiments, and exposed for 30 min of saturating blue (450 nm, 15 µmol of photons/m²/sec) or far-red (800 nm, 15 µmol of photons/m²/sec) light, or placed in the dark. About 1.5e⁸ cells were harvested by filtration and total RNA was extracted as in Huysman et al. (2013). For qRT-PCR, 500 ng of total RNA were reverse-transcribed using the QuantiTect Reverse Transcription Kit (Qiagen, USA) and the reaction accomplished with 12.5 ng cDNA as template, following the SsoAdvanced Universal SYBR Green Supermix instructions (Bio-Rad, USA), in a CFX 96 Real-Time Detection System (Bio-Rad). Primer sequences are indicated in Table S2. *H4* were used as reference gene as in Siaut et al. (2007) and normalization was performed to the values before specific exposure (i.e. cells grown in continuous green light at 15 µmol of photons/m²/sec).

Pt-DPH KO mutant complementation :

Tp-DPH coding sequence was amplified and domesticated from *T. pseudonana* cDNA by
fusing with the primer pair UNS1FL.FW and UNSXFL.RV, the PCR products obtained with
the primer pairs: TpPHY.D.Fw and TpPHY.Sap1.Rv, TpPHY.Sap1.Fw and TpPHY.BsaI.Rv,
TpPHY.BsaI.Fw and TpPHY.Sap2.Rv, TpPHY.Sap2.Fw and TpPHY.E.Rv.

The 1041 nt upstream PtDPH start codon and the 368 nt downstream the PtDPH stop
codon were amplified from *P. tricornutum* gDNA with the primer pairs : PrPtPHY.A.Fw
and PrPtPHY.C.Rv, and TrPtPHY.E.Fw and TrPtPHY.F.Rv, respectively. The Tp-DPH
domesticated coding sequence, Pt-DPH promoter region and terminator regions were
each cloned into the pL0 vector from the modular cloning system uLoop and assembled
with the Venus fluorescent protein sequence in pL1 as in Pollack et al, (2020). The
assembled product was amplified with the primer pair UNS1FL.FW and UNSXFL.RV, and
inserted into the SmaI-linearized pUC19 vector containing the Blasticidin resistance cassette of the pPTbsr (Buck et al, 2018) in Eco53KI site.

## Expression, purification and spectral analysis of Dph proteins

Photosensory module (PSM) of DPH genes were obtained either from previous studies (Pt-DPH, Tp-DPH (Fortunato et al, 2016)), cloned from cDNAs generated as described above, from cultures of *S. costatum*, *C. cryptica*, *Minidiscus spinulatus*, *S. biocatulatus*, *A. cornucervis*, or synthetized by GenScript (USA) (for the 2 near full length sequences identified in MATOU, *Minidiscus*-like and *M. polymorphus*-like) with codons optimized for *E. coli* expression. Primer pairs used are provided in Suppl Table2). Sequences were amplified with the primers pairs indicated in the Table S2, and cloned into pET28a-HO vector generated by inserting the *Synechocystis* Heme oxygenase gene amplified from pKS270 vector (Mukougawa et al., 2006) with the primer pair HO1xpET.HindIII.Fw and HO1xpET.NotI.rv, in HinDIII/NotI in the pET28a vector (Novagen). PSM sequences were expressed as N-terminal His6 tagged proteins in *E. coli* BL21 (DE3) strain, by auto-induction system (Studier, 2005). Recombinant proteins were purified as in Fortunato et al. (2016) and their absorption spectra of the recombinant proteins were measured immediately after purification on a Varian's Cary-50 spectrophotometer. Illumination with LEDs at 810, 630 and 405 nm were performed (approx. 1min illumination) to reach pure Pr spectra (after 810 illumination) or equilibriums between the Pr and Pfr forms. These spectra were used as in Giraud et al (2010) to calculate pure Pfr spectra and ratios of quantum yields.

## Search for DPH and diatom Aureochrome genes in environmental sequence data

Known DPH sequences were aligned with mafft v7.4 to generate separate Centric and Pennate HMM profiles for each DPH domain, covering nearly the full-length protein. HMM searches (HMMER version 3.2) were performed against different set of control sequences: protein sequences of known phytochromes from streptophytes, chlorophytes, glaucophytes, cryptophytes, bacteria, cyanobacteria, fungi and Stramenopiles, the MetDB dataset, diatom genomes (P.tri, T.pse, C.cry and F.cyl) and a set of proteins specific for each domain (either the proteins used in the corresponding Pfam domain alignment or if these were too few in numbers, 1000 randomly selected proteins from proteins known to
possess the specific domain in InterPro). All domains with a e-value<1e-5 were retrieved, clustered at 90% identity with cd-hit and submitted at EFI to generate Sequence Similarity Network (SSN). Alignment scores for the SSN were chosen so that domain sequences from diatom phytochromes clustered together, not linked to other diatom proteins nor to non-diatom phytochromes (as illustrated in Fig. S8; see Table S3 for the SSN alignment score values for each domain). The same search method was then applied to the different environmental sequence dataset (MATOU gene atlas (6-frame translated) and Tara SMAGs (Carradec et al, 2018; Delmont et al, 2020)

The same method was applied to the aureochromes except for a few changes: only one HMM profile was generated from diatom aureochromes protein sequences, covering both the bZIP and the LOV domain; SNN being insufficient to distinguish diatom aureochromes from other aureochromes, environmental sequences were placed on an Aureochrome phylogenetic tree. Briefly, sequences were aligned to the Aureochrome HMM model using hmmalign and the phylogenetic tree was reconstructed with FastTree (v2.1, default settings). Only sequences branching in the diatom clades (as delimited by the sister Bolidophyceae clade) were annotated as diatom aureochromes.

Mapped reads to the MATOU sequences were retrieved from each Tara Oceans sample. Relative abundance was expressed as the sum of all reads mapping to centric DPH (or centric Aureos) divided by the sum of all the reads mapping to centric diatom genes. The analysis was performed on each filter size separately (Supplementary figures, 5-20µm, 20-180µm, 180-2000µm and 0.8-inf), or average values of the 4 filter sizes when all 4 were available. Environmental variables measured *in situ* during the Tara Oceans campaign are available at PANGEA (Pesant et al., 2015).

Influence of environmental parameters on DPH abundances were assessed with Spearman's correlations or GAMs (mgcv package in R). Only variables with high explanatory power in the individual GAM analysis (pvalue of the smooth term<0.5, explained deviance>0.35) and that did not correlate to each other (spearman's rho<0.7) were used for complex GAMs. The "proportion of explained deviance" attributed to each environmental parameter was obtained by performing the same GAM without this parameter; we then used the difference of explained deviance between full and reduced
GAM as the “proportion of explained deviance”, i.e. the weight of this parameter in the model.

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<figure><img src="image_40.png" /><figcaption>Supplementary Figure 1: Experimental setup for the action and inhibition spectra. A. Illustration of an illumination of reporter lines to a gradient of a monochromatic LED. B. Spectra of the different LEDs used for the action and inhibition spectra.</figcaption></figure>

<figure><img src="image_41.png" /><figcaption>Supplementary Figure 2: Control lines show no induction of YFP in the action spectra. Control lines correspond to: PtDPH KO background transformed with the ProHsf4.6a::YFP construct, or WT background transformed with a DPH-independent promoter controlling eYFP expression, ProLHCF2 or ProH4 (Siaut et</figcaption></figure>
al, 2007)). Cells were exposed to a gradient of monochromatic LED. YFP signal was measured by flow cytometry.

<figure><img src="image_42.png" /></figure>

Supplementary Figure 3: Expression analysis of PtDPH-regulated (HSF4.6a, Phatr3_J15138, Phatr3_J18096, Phatr3_J45662, Phatr3_J46431) and PtDPH-non regulated (Phatr3_J18180) genes upon far-red and blue irradiation. *P. tricornutum* WT, DPH KO mutant and their corresponding non mutated transformant (Tc) cells were collected in their continuous green light growth condition (G) and following a 30 min of irradiation with 470-nm (B) or 765-nm (FR) LED (15 µmol·m⁻²·s⁻¹) or kept in the dark for the same time (D). Gene expression quantifications were performed by RT-qPCR, with *H4* used as normalizer gene and relativized to the G conditions. Values are the mean +/- SD on 3 independent biological replicates.
<figure><img src="image_43.png" /><figcaption>Supplementary Figure 4. Light modeling examples. A and D, Modelled light fields at the bottom of the photic zone (1% of surface irradiance) for sun at Zenith, for different Chl a concentrations (Kbg=0) (A), and different Kgt440 (Chl a=0) (D). B,C,E,F,G and H, variation of the ratio of blue (400-450nm) to green (500-</figcaption></figure>
550nm)(C, F, H) or red (630-680nm) 'B, E, G) bands in different modelled light fields: B and C, ratios from light fields showed in A; E and F, ratios from light fields showed in D; G and H ratios from the light fields at different solar zenith angles as selected depths, for Chl a=0.5 mg/m³ and Kgt440=0.06m⁻¹

<figure><img src="image_44.png" /></figure>

Supplementary Figure 5: DPH activity as proportion of PrPr formed (%PrPr) in different modelled environmental conditions: at saturation (A) and upon 10 min of exposure (B), depending on the depth and different Kgt440; at saturation at 75-m depth depending on the Chl a concentration (C) and Kgt440 (D); upon 1h of exposure of light spectrum at different depths and depending on the sun Zenith angle (E).
<figure><img src="image_45.png" /><figcaption>Supplementary Figure 6: DPH activity as proportion of PrPr formed (%PrPr) in different *in situ* measured environmental light fields as a function of the time of the day over a year time series. Data from Veedin Rajan, 2021, with a downwelling irradiance spectra taken every 15 min at 10 m-depth near Ischia (Gulf of Naples, Italy) in 2010-2011. DPH activity at saturation was calculated.</figcaption></figure>

<figure><img src="image_46.png" /><figcaption>Supplementary Figure 7: Biogeography of DPH and Aureochromes (Aureo) from centric diatoms, per filter size, in metagenomic and metatranscriptomic data from *Tara Oceans*. Crosses indicate that no gene was</figcaption></figure>
found at the station, while circles indicate presence. Circles size indicate the abundance of the DPH or Aureo genes relative to all centric genes.

<figure><img src="image_47.png" /><figcaption>Supplementary Figure 8: Metagenome Assembled Genomes (MAGs) geographical distribution of centric diatoms. Black crosses indicate *Tara Oceans* station where the MAG was not found present; colored points indicate the photoreceptor content of the MAG, while the shape indicate the centric diatom family the MAG belongs too.</figcaption></figure>
<figure><img src="image_48.png" /><figcaption>Supplementary Figure 9: DPH relative abundance (A) compared to Aureochrome (B) relative abundance as a function of absolute latitude. Point shapes indicate the filter size range.</figcaption></figure>
<figure><img src="image_49.png" /></figure>

<figure><img src="image_50.png" /><figcaption>Supplementary Figure 10: Correlation (A) and individual generalized additive models (B) analysis for DPH and Aureochromes (Aureo) relative abundance in metagenomic and metatranscriptomic data from *Tara Oceans*, separated by filter sizes.</figcaption></figure>
<figure><img src="image_51.png" /><figcaption>Supplementary Figure 11: Complex generalized additive models explaining DPH relative abundance with a combination of environmental parameters for each filter size.</figcaption></figure>

<figure><img src="image_52.png" /><figcaption>Supplementary Figure 12: Alignment of a portion of the PAS and GAF domains of selected DPH and reference phytochrome sequences (Arabidopsis thaliana, *Atha_PhyA*; *Synechocystis*, *Syne_Cph1*; *Neurospora crassa* PHY2, tr-Q7S5Q3, and *Deinoccocus radiodurans*, *Drad_Bph*). In red is highlighted the cysteine residue conserved with the one used for chromophore binding in the PAS domain in bacterial and fungal phytochromes, and in the GAF domain, in plant and cyanobacteria phytochromes. Tp, *Thalassiosira pseudonana*; Mpol-like, *Minitocellus polymorphus*-like sequence from MATOU; Acor, *Arcocellulus cornucervis*, Mini-like, *Minidiscus*-like sequence from MATOU; Scos, *Skeletonema costatum*; Sbio, *Shionodiscus bioculatus*; Ccry, *Cyclotella cryptica*; Ptri, *Phaeodactylum tricornutum*, Acof, *Amphora coffeaeformis*.</figcaption></figure>
<figure><img src="image_53.png" /><figcaption>Supplementary Figure 13 : Purified recombinant DPH. Photosensor module of DPH genes expressed as a N-terminal 6 Histidine-tagged recombinant proteins along with *Synechocystis* Heme oxygenase gene encoding the enzyme allowing the synthesis of the biliverdin chromophore, were purified on Ni-NTA affinity chromatography. From left to right: *Minutocellus polymorphus*-like sequence; *Skeletonema costatum*; *Arcocellulus cornucervis*; *Cyclotella cryptica*; *Thalassiosira pseudonana*; *Minidiscus*-like sequence; *Shionodiscus bioculatus*.</figcaption></figure>

<figure><img src="image_54.png" /><figcaption>Supplementary Figure14: TpDPH restores blue and far-red-dependent YFP induction in PtDPH KO line. PtDPH KO reporter strain (line KO #1 in Supplementary Figure 2) was transformed with TpDPH gene under the control of PtDPH promoter and terminator, and subjected to a gradient of blue (430 nm) or far-red (765) lights, according to the same experimental setup used for the action spectra.</figcaption></figure>
<figure><img src="image_55.png" /></figure>

<figure><img src="image_56.png" /></figure>

<figure><img src="image_57.png" /></figure>
Supplementary Figure 15: Projection of DPH activity as proportion of PrPr formed (%PrPr), in modeled light fields, with variations of chlorophyll a (Chl a), Kbg440 and solar zenith angle, using properties of PtDPH (A), TpDPH (B) and ScosDPH (C), determined in vitro, from the recombinant protein absorption spectra. The color legend is common to all 3 figures.

<figure><img src="image_58.png" /></figure>

Supplementary Figure 16: Method for DPH and diatom Aureochrome gene search. A, Example of Sequence Similarity Network of the DPH REC domain, searched in Tara Oceans data showing a nice separation of the different phytochromes (centric diatom, pennate diatom, other stramenopiles) for an alignment score of 20. Environmental sequences grouping with the centric or pennate DPH will be annotated as such. B, Phylogenetic tree of diatom Aureochrome gene search. Diatom clades are identifiable (grey label), delimited by the Bolidophyceae (a sister picoplanktonic group of diatoms) Aureochromes (pink labels). Branches are colored by the different Aureochrome types: Aureochrome 1a, 1b, 1c and 2.
SUPPLEMENTARY TABLES

<table><caption>Table S1. Species from which DPH have been characterized in this study and previous one* (*Fortunato et al, 2016) and information about their isolation site.</caption><thead><tr><td>Species</td><td>Strain</td><td></td></tr></thead><tbody><tr><td>Phaeodactylum tricornutum*</td><td>CCMP2561</td><td>North Atlantic, off Blackpool, England</td></tr><tr><td>Thalassiosira pseudonana*</td><td>CCMP1335</td><td>North Atlantic, Moriches Bay, Forge River, Long Island, New York USA</td></tr><tr><td>Cyclotella cryptica</td><td>CCMP332</td><td>North Atlantic, Martha's Vineyard, Massachusetts USA</td></tr><tr><td>Minidiscus spinulatus</td><td>RCC4659</td><td>Atlantic Ocean, English Channel, Britanny coast, pelagic</td></tr><tr><td>Skeletonema costatum</td><td>RCC1617</td><td>Atlantic Ocean, English Channel, Britanny coast, pelagic</td></tr><tr><td>Arcocellulus cornucervis</td><td>RCC2270</td><td>Arctic Ocean, Beaufort Sea, at 0m depth pelagic</td></tr><tr><td>Shionodiscus bioculatus</td><td>RCC1991</td><td>Arctic Ocean, Beaufort Sea, at 65m depth, pelagic,</td></tr></tbody></table>

<table><caption>Table S2 : Sequences of primers used in this study</caption><tr><td>qPCR-Phatr3_45662FW</td><td>CGAGGGAGCTCGGTTTATGG</td></tr><tr><td>qPCR-Phatr3_45662RV</td><td>TGATGGGAACTGTTCTGCCC</td></tr><tr><td>qPCR-Phatr3_46431FW</td><td>GGTTTGCGAGTGCATTTGGT</td></tr><tr><td>qPCR-Phatr3_45662RV</td><td>TGTCAGCAACCTCATTCCCC</td></tr><tr><td>qPCR-Phatr3_18180FW</td><td>CCGGGAACGTAGGTTTGAT</td></tr><tr><td>qPCR-Phatr3_18180RV</td><td>CCGCGGCCAACATAGCAAG</td></tr><tr><td>qPCR-H4FW</td><td>AGGTCCTTCGCGACAATATC</td></tr><tr><td>qPCR-H4RV</td><td>ACGGAATCACGAATGACGTT</td></tr><tr><td>Hsf4.6ap_Fw0</td><td>TCTAGAGAGCTCGGATTTCGAATCTGTTTTGGGCA</td></tr><tr><td>Hsf4.6ap-YFP_RV</td><td>GATATCGGATCCTGTCAAAGGTTTAAGAGAATCGGC</td></tr><tr><td>YFP-Hsf4.6ap_Fw</td><td>AAACCTTTGACAGGATCCGATATCATGGTGAGCAAGGGCGAG</td></tr><tr><td>FcpAT_Rv0</td><td>TCTAGATGAAGACGAGCTAGTGTTATTCC</td></tr><tr><td>HO1xpET.HindIII.Fw</td><td>GCAAGCTTAGAAGGAGATATACATATGAG</td></tr><tr><td>HO1xpET.NotI.Rv</td><td>GCGCGGCCGCCTAGCCTTCGGAGGTGGCGAG</td></tr><tr><td>TpPHY.D.Fw</td><td>AGGCTGTCTCGTCTCGTCTCAGGTCTCAAGGTATGAGTGTCAAAAAGAGCAC</td></tr><tr><td>TpPHY.Sap1.Rv</td><td>CAATGTGTTTCGCATGAACAGCTCTCGCTTG</td></tr><tr><td>TpPHY.Sap1.Fw</td><td>CAAGCGAGAGAGCTGTTCATGCGAAACACATTG</td></tr><tr><td>TpPHY.Bsal.Rv</td><td>CAAGCTGTTTATCAGGGTCACCACCCCACGTGACAC</td></tr><tr><td>TpPHY.Bsal.Fw</td><td>GTGTCACGTGGGGTGGTGACCCTGATAAACAGCTTG</td></tr><tr><td>TpPHY.Sap2.Rv</td><td>GAGATCGAACAAAGTGCTCCTCAATCTGAAGGTCGTATG</td></tr><tr><td>TpPHY.Sap2.Fw</td><td>CATACGACCTTCAGATTGAGGAGCACTTGTTCGATCTC</td></tr><tr><td>TpPHY.E.Rv</td><td>TGGTAATCTATGTATCCTGTTGGTCTCTAAGCTCATCGTTCATTTTTGTGAT</td></tr></table>
<table><caption>Table S3. SSN parameters for DPH and diatom Aureochrome search in *Tara Oceans* data</caption><tr><td>PrPtPHY.A.Fw</td><td>GGCTGTCTCGTCTCGTCTCAGGTCTCAGGAGCCCGGGGATATCGAAGATCC</td></tr><tr><td>PrPtPHY.C.Rv</td><td>TGGTAATCTATGTATCCTGGTGGTCTCGCATTTTTAAAGGCGTGGTTCCTTG</td></tr><tr><td>TrPtPHY.E.Fw</td><td>TCGTCTCGTCTCAGGTCTCAGCTTCATGGTCGTTCATTCATAGAAG</td></tr><tr><td>TrPtPHY.F.Rv</td><td>TGGTAATCTATGTATCCTGGTGGTCTCAAGCGCGCTCTTTCCACCTCATCTC</td></tr><tr><td>UNS1FL.FW</td><td>CATTACTCGCATCCATTCTCAGGCTGTCTCGTCTCGTCTC</td></tr><tr><td>UNSXFL.RV</td><td>GGTGAAGGGCTCGGAGTTGTGGTAATCTATGTATCCTGG</td></tr><tr><td>SbioPHY.Nhel.Fw</td><td>GCGGCTAGCATGTCTGCCAGTTCCACCAC</td></tr><tr><td>SbioPHY_PCD.Sall.Rv</td><td>GCGGTCGACCTAAAGATTTTCCTTTTGATCTTTG</td></tr><tr><td>AcorPHY.Nhel.Fw</td><td>GCGGCTAGCATGTCGGCACCTGCGGCAGC</td></tr><tr><td>AcorPHY_PCD.Sacl.Rv</td><td>GCGGAGCTCCTAGTAGCTTGTTGTGTTCTCGTC</td></tr><tr><td>MspiPHY.Nhel.Fw</td><td>GCGGCTAGCATGACCTCCTCCTCAACCAAC</td></tr><tr><td>MspiPHY_PCD.Sacl.Rv</td><td>GCGGAGCTCCTACTTCTGATCGGCAATCAATTC</td></tr><tr><td>ScosPHY.Spel.Fw</td><td>GCGACTAGTATGTCGTCCACCAATAGCAC</td></tr><tr><td>ScosPHY_PCD.Sacl.Rv</td><td>GCGGAGCTCCTAGAGGTTTTTCCTTTTGATCTTG</td></tr><tr><td>CcryPHY.Nhel.Fw</td><td>GCGGCTAGCATGGCAGCACCCCACAAAAC</td></tr><tr><td>CcryPHY_PCD.Sacl.Rv</td><td>GCGGAGCTCCTACTTCTGATCTTTAATCAAATC</td></tr></table>

<table><thead><tr><td>Domain model</td><td>SSN alignment score threshold</td></tr></thead><tbody><tr><td>Centric GAF</td><td>30</td></tr><tr><td>Centric PHY</td><td>10</td></tr><tr><td>Centric HisKA</td><td>20</td></tr><tr><td>Centric HATPase</td><td>20</td></tr><tr><td>Centric REC</td><td>20</td></tr><tr><td>Pennate GAF</td><td>30</td></tr><tr><td>Pennate PHY</td><td>10</td></tr><tr><td>Pennate HisKA</td><td>20</td></tr><tr><td>Pennate HATPase</td><td>20</td></tr><tr><td>Pennate REC</td><td>20</td></tr><tr><td>Diatom Aureochrome</td><td>70</td></tr></tbody></table>

## SUPPLEMENTARY DATA

Supplementary Data: List of MATOU2 genes identified as diatom aureochromes and diatom phytochromes. Available at

https://mycore.core-cloud.net/index.php/s/hD5zBsV1yhGkvfH
**Detailed description of DPH activity model construction based on equations described in (Mancinelli, 1994)**

Phytochrome equilibrium can be theorized as the equilibrium between the $Pfr \rightarrow Pr$ transition (kinetic constant k1) and the $Pr \rightarrow Pfr$ reverse reaction (kinetic constant k2) with the following equation:

$$Pfr \underset{k1}{\overset{k2}{\rightleftharpoons}} Pr \quad (1)$$

The rate of phytochrome photoconversion depends on phytochrome extinction coefficient, photoconversion yield and on light intensity, so k1 and k2 can be expressed as:

$$k_1 = N * \sigma_{Pfr} = N * 2.3 * \varepsilon_{Pfr} * \phi_{Pfr} \quad \text{and} \quad k_2 = N * 2.3 * \varepsilon_{Pr} * \phi_{Pr}$$

where N is the fluence rate, or light intensity (mol photon/m²/s), and $\sigma_{Pfr}$ is the photoconversion cross-section of the $Pfr \rightarrow Pr$ reaction, $\varepsilon_{Pfr}$ is the extinction coefficient of Pfr and $\phi_{Pfr}$ the quantum yield of the $Pfr \rightarrow Pr$ reaction; $\sigma_{Pr}$ is the photoconversion cross-section of the $Pr \rightarrow Pfr$ reaction, $\varepsilon_{Pr}$ is the extinction coefficient of Pr and $\phi_{Pr}$ the quantum yield of the $Pr \rightarrow Pfr$ reaction.

The photoconversion rate of phytochrome can be expressed as the sum of k1 and k2 and expressed as k.

Considering that the total amount of phytochrome P stays constant, i.e. synthesis and degradation are negligible, then $P = P_r + P_{fr}$

From equation 1, the rate of Pr production can be expressed as :

$$\frac{dPr}{dt} = k_1 * Pfr - k_2 * Pr \quad (2)$$
$$ \text{At equilibrium, } \frac{dPr}{dt} = 0 \text{ and } \%Pr = \frac{Pr}{P} = \frac{k_1}{k_1+k_2} = \frac{1}{1+\frac{\varepsilon_{Pr}}{\varepsilon_{Pfr}}\frac{\phi_{Pr}}{\phi_{Pfr}}} = \frac{1}{1+\frac{\varepsilon_{Pr}}{\varepsilon_{Pfr}*\eta}} $$

$$ \text{With } \eta = \frac{\phi_{Pr}}{\phi_{Pfr}} \text{ the ratio of quatum yields.} $$

Integration of equation (2) also gives:

$$ \%Pr(t) = \left( \%Pr0 - \frac{k_1}{k_1 + k_2} \right) e^{-(k_1+k_2)t} + \frac{k_1}{k_1 + k_2} $$

with %Pr0 the percent of Pr at time t=0.

If one considers that phytochromes is in a dimeric form (Klose et al., 2015; Brockmann et al.), they exist in equilibrium between homodimer PrPr, homodimer PfrPfr and the intermediate heterodimer PfrPr. The rate of conversion of each monomer is supposed independent of the dimer state, and each monomer photoconverts as in the monomer model explained previously, and can be expressed as:

$$ PfrPfr \underset{2k_1}{\overset{k_2}{\rightleftharpoons}} PrPfr \underset{k_1}{\overset{2k_2}{\rightleftharpoons}} PrPr \quad (3) $$

We always consider that synthesis and degradation are negligible, with

$$ P = PrPr + PrPfr + PfrPfr $$

Considering PrPr as the active form and total amount of phytochrome constant, the PrPr photoconversion rate can be expressed as:

$$ \frac{dPrPr}{dt} = k_1 * PrPfr - 2k_2 * PrPr $$

At equilibrium, $\frac{dPrPr}{dt} = 0$ and:

$$ \%PrPr_{eq} = \left( \frac{k_1}{k_1 + k_2} \right)^2 = \frac{1}{\left( 1 + \frac{\varepsilon_{Pr}}{\varepsilon_{Pfr}} \frac{\phi_{Pr}}{\phi_{Pfr}} \right)^2} = \frac{1}{\left( 1 + \frac{\varepsilon_{Pr}}{\varepsilon_{Pfr}*\eta} \right)^2} \quad (4) $$
In this latter equation, $\varepsilon_{Pr}/\varepsilon_{Pfr}$ can be known from the recombinant PtDPH absorption spectra.

The $\varepsilon_{Pr}/\varepsilon_{Pfr}$ ratio, in both monochromatic and bichromatic illumination, as been determined with the LED spectra used (and not only the $\varepsilon_{Pr}$ value at the peak of intensity of the LED). More exactly, we express $\varepsilon_{Pr}/\varepsilon_{Pfr}$ as:

$$ \frac{\varepsilon_{Pr,LED}}{\varepsilon_{Pfr,LED}} = \frac{\sum_{\lambda=300}^{\lambda=900} A_{Pr,\lambda} * N_{\lambda}}{\sum_{\lambda=300}^{\lambda=900} A_{Pfr,\lambda} * N_{\lambda}} $$

where $A_{Pr,\lambda}$ is the absorbance of Pr at wavelength $\lambda$ and $N_{\lambda}$ the intensity measured at this wavelength.

%PrPr from equation (4) can be estimated from the eYFP signal at saturating light at each wavelength. Our output measure is the eYFP signal for a LED at wavelength $\lambda$ at saturating intensities (the 4 points at highest light intensity were used), normalized between the signal from cells that went from constant green light (520nm, growth condition) to darkness, and the signal of cells exposed 10 min to saturating far-red light (765nm). This is expressed as following:

$$ \text{NormalizedYFP}(\text{LED}\lambda, \text{sat}) = \frac{e^{\text{YFP}}(\text{LED}\lambda, \text{sat}) - e^{\text{YFP}}(520\text{nm}, \text{growth})}{e^{\text{YFP}}(765\text{nm}, \text{sat}) - e^{\text{YFP}}(520\text{nm}, \text{growth})} $$

If we consider that the eYFP is linearly linked to %PrPr, then :

$$ \text{NormalizedYFP}(\text{LED}\lambda, \text{sat}) = \frac{\%PrPr_{eq,LED} - \%PrPr_{eq,520nm}}{\%PrPr_{eq,765nm} - \%PrPr_{eq,520nm}} $$

And therefore :

$$ \text{NormalizedYFP}(\text{LED}\lambda, \text{sat}) = \frac{\frac{1}{(1+\frac{\varepsilon_{Pr,LED}}{\varepsilon_{Pfr,LED}}*\eta)^2} - \frac{1}{(1+\frac{\varepsilon_{Pr,520}}{\varepsilon_{Pfr,520}}*\eta)^2}}{\frac{1}{(1+\frac{\varepsilon_{Pr,765}}{\varepsilon_{Pfr,765}}*\eta)^2} - \frac{1}{(1+\frac{\varepsilon_{Pr,520}}{\varepsilon_{Pfr,520}}*\eta)^2}} (5) $$

In this equation, $\eta$ is the only unknown.
To determine it, the equation was fitted with the nls function in R with η= 1 as starting value, on the experimental values obtained from monochromatic and bichromatic illuminations.

Integration of Equation 4 gives the following expression for %PrPr as a function of time:

$$
\%PrPr(t) = \left( \frac{k_1}{k_1+k_2} * \%PfrPfr0 + \frac{k_2}{k_1+k_2} * \%PrPr0 - \frac{k_1 * k_2}{(k_1+k_2)^2} \right) e^{-2*(k_1+k_2)t} + \\ \frac{k_1}{k_1+k_2} (\%PrPr0 - \%PfrPfr0 - \frac{k_2-k_1}{k_1+k_2})e^{-(k_1+k_2)t} + \left( \frac{k_1}{k_1+k_2} \right)^2 (6)
$$

To determine k, the following equation (7) was fitted to the eYFP levels further
normalized for each LED and each strain (resulting in value between 0 and 1, and the
fitting of the exponential will be independent on the fitting of the values at equilibrium as
done above. However, we used the ratio of photoconversion η estimated above):

$$
\text{NormalizedYFP}(\lambda, N) = \frac{\left( \frac{k_1}{k_1+k_2} * \%PfrPfr0 + \frac{k_2}{k_1+k_2} * \%PrPr0 - \frac{k_1 * k_2}{(k_1+k_2)^2} \right) e^{-2 * k_{\text{Fit}} * N * t} + \frac{k_1}{k_1+k_2} (\%PrPr0 - \%PfrPfr0 - \frac{k_2-k_1}{k_1+k_2}) e^{-k_{\text{Fit}} * N * t} + \left( \frac{k_1}{k_1+k_2} \right)^2 - \%PrPr0}{\left( \frac{k_1}{k_1+k_2} \right)^2 - \%PrPr0} \quad (7)
$$

where %PrPr at time 0 is either the equilibrium in green (for the action
spectra) or in far-red (for the inhibition spectra), t, the time of illumination, i.e.
10min, and k<sub>Fit</sub> is the exponential constant to be estimated.

Further analysis of k and k<sub>Fit</sub> in Equations 6 and 7 gives:

$$
k * t = (k_1 + k_2) * t = (2.3 * \varepsilon_{Pr} * \phi_{Pr} + 2.3 * \varepsilon_{Pfr} * \phi_{Pfr}) * N * t
$$

$$
\text{so } k = \alpha * (A_{\text{Pr,LED}} * \eta + A_{\text{Pfr,LED}}) * N = k_{\text{Fit}} * N
$$

$$
\text{where } A_{\text{Pr,LED}} = \frac{\sum_{\lambda=300}^{\lambda=900} A_{\text{Pr},\lambda} * N_{\lambda}}{\sum_{\lambda=300}^{\lambda=900} N_{\lambda}}, \quad N = \sum_{\lambda=300}^{\lambda=900} N_{\lambda} \text{ and } \alpha \text{ is a constant}
$$

In theory, *k*<sub>Fit</sub> should be a linear combination of (*A*<sub>Pr</sub>*\* η + *A*<sub>Pfr</sub>) (dotted lined on Figure 3D).
Fitting linear relationship between k<sub>Fit</sub> and (A<sub>Pr</sub>* η + A<sub>Pfr</sub>) with the lm function in R gave α=0.012033+/-0.001969 (adjusted R<sup>2</sup>=0.4311) when considering all the data points (Figure 3C), but α=0.02005+/-0.002906 (adjusted R<sup>2</sup>=0.6247) when ignoring the data points above 700nm.

From the previous equations, we can calculate %PrPr(t) in a given environment.

The light environment is described with I<sub>tot</sub> = l * Σ<sub>λ=300</sub><sup>λ=900</sup> N<sub>λ</sub> with l the bandwidth and N<sub>λ</sub> the intensity at wavelength λ.

We can calculate Pr and Pfr absorption in this environment with A<sub>Pr</sub> = (Σ<sub>λ=300</sub><sup>λ=900</sup> A<sub>Pr,λ</sub>*N<sub>λ</sub>) / (Σ<sub>λ=300</sub><sup>λ=900</sup> N<sub>λ</sub>),
from which we calculate k<sub>Fit</sub> = 0.02005 * (A<sub>Pr</sub> * η + A<sub>Pfr</sub>)

The ratio of absorption is calculated with $\frac{\varepsilon_{Pr}}{\varepsilon_{Pfr}} = \frac{\sum_{\lambda=300}^{\lambda=900} A_{Pr,\lambda}*N_{\lambda}}{\sum_{\lambda=300}^{\lambda=900} A_{Pfr,\lambda}*N_{\lambda}}$, from which we can calculate the ratio of photoconversion:

$$ \frac{k_1}{k_1 + k_2} = \frac{1}{1 + \frac{\varepsilon_{Pr}}{\varepsilon_{Pfr}} * \eta} $$

($\eta$ is either the value determined in vivo or in vitro)

We have all the elements to calculate %PrPr(t)

$$ \%PrPr(t) = \left( \frac{k_1}{k_1 + k_2} * \%PfrPfr0 + \frac{k_2}{k_1 + k_2} * \%PrPr0 - \frac{k_1 * k_2}{(k_1 + k_2)^2} \right) e^{-2 * k_{Fit} * ltot * t} + \frac{k_1}{k_1 + k_2} (\%PrPr0 - \%PfrPfr0 - \frac{k_2 - k_1}{k_1 + k_2}) e^{-k_{Fit} * ltot * t} + \left( \frac{k_1}{k_1 + k_2} \right)^2 $$

with %PrPr0=0 and %PfrPfr0=1 and t=10min or t=1h.
CHAPTER 3: DPH IN BENTHIC SPECIES

Following the results from chapters 1 and 2, it was shown that amongst pennate diatom possessing DPH, most are found in benthic environments, and some have multiple DPH copies. Therefore, we wanted to explore the links between DPH in pennate species and the benthic environment. For this, we characterized the 4 DPH from a benthic diatom, *Amphora coffeaeformis*, as recombinant proteins. We also explored the physiological response of pennate species to red light, which was shown to induce chromatic acclimation in some of them (appearance of fluorescence emission at 710nm). We observed this fluorescence in several benthic species, and investigated the role of DPH in this phenomenon.

I participated in amplifying AcDPH CDS from cDNA and I purified recombinant DPH expressed in *E.coli* and characterized their absorption spectra. I was responsible of the characterization of fluorescence emission at 710nm under some conditions.
In the previous chapter, we have seen that DPH is sensitive to the whole light spectra. As a consequence, in the open ocean, DPH will sense mostly variation of the blue to green wavebands due to the absence of red and far-red lights after few meters. In addition, we showed that in the open ocean, essentially centric species possess DPH with a strong latitudinal repartition gradient. In the chapter one, we saw (as previously shown by (Fortunato et al., 2016), that centric and pennate DPH form separate clades, suggesting independent acquisition, maybe from viruses. Moreover, most of the pennate species known to possess DPH have a benthic/tychoplanktonic lifestyle, and some show multiplication of DPH copy number while centric diatoms only possess one DPH. This suggests that DPH conservation and multiplication is linked to the adaptation to benthic environments. In these environments, such as sediments and biofilms, blue light is strongly absorbed, resulting in red and far-red light enriched light field, where DPH could be sensing either sediment depth or the density of other photosynthetic organisms through the red/far-red ratio.

This led us to consider that DPH could have a different role in centric and pennate species, or at least mediate the response to different light cues. Centric species living in the open ocean sense blue to green ratio through phytochrome, which could help them to adapt to low blue light, highly mixed environments, or trigger sexual reproduction for example. Pennate species living in the sediments would use DPH to sense red/far-red light (or absence of blue) to adapt to life inside the sediments, i.e. low red light, metabolism (heterotrophy in the dark), phototaxis or sexual reproduction.

Recent analyses of the *Seminavis robusta* genome (Osuna-Cruz et al., 2020) and omics approaches (Chapter 1) suggest that an increase in photoreceptor numbers in some pennate diatoms may be related to benthic lifestyle. Indeed, DPH phytochromes are present in several copies in pennate diatoms (See Table1). This expansion appears essentially for species known to live in sediments. We therefore wanted to explore the significance of DPH in sediments by addressing several questions. Do the different
phytochromes present in the same species have the same photochemical properties? Can we associate the phytochrome to a chromatic acclimation linked to this particular light environment? Is this adaptation benthic and/or DPH specific?

<table><caption>Table 1. Pennate diatom species with phytochrome(s). In bold, species found in a tidal flat in (An et al., 2020)</caption><thead><tr><td>Diatom species</td><td>Number of phytochromes</td></tr></thead><tbody><tr><td><i>Phaeodactylum tricornutum</i></td><td>1</td></tr><tr><td><b>Amphora coffeaeformis</b></td><td><b>4</b></td></tr><tr><td><b>Seminavis robusta</b></td><td><b>4</b></td></tr><tr><td><i>Fragilaria radians</i></td><td>4</td></tr><tr><td><i>Fragilaria sp</i></td><td>2</td></tr><tr><td><b>Nitzschia punctata</b></td><td><b>2</b></td></tr><tr><td><b>Nitzschia palea</b></td><td><b>2</b></td></tr><tr><td><b>Nitzschia sp</b></td><td><b>2</b></td></tr><tr><td><b>Nitzschia inconspicua</b></td><td><b>2</b></td></tr><tr><td><b>Navicula sp</b></td><td><b>2</b></td></tr><tr><td><b>Gyrosigma sp</b></td><td><b>1</b></td></tr><tr><td><i>Craticula ambigua</i></td><td>2</td></tr><tr><td><i>Diatoma tenuis</i></td><td>1</td></tr><tr><td><i>Synedra sp</i></td><td>1</td></tr><tr><td><i>Synedropsis recta</i></td><td>1</td></tr><tr><td><i>Staurosira complex</i></td><td>1</td></tr></tbody></table>

# RESULTS

## Different DPH copies show different light-sensing abilities

Exploring the genomes and transcriptomes of diatom in the search for DPH genes unveiled that not all diatom possesses DPH, but that a lot of pennate species possess several copies of DPH genes, suggesting that these DPH can perform different functions or have some specific properties such as observed in plants (Franklin and Quail, 2010; Burgie et al., 2021) (See Table 1). Among these diatoms, the benthic diatom *Amphora*
*coffeaeformis*, possesses 4 DPH genes, and 3 of them possessing not only a putative chromophore-binding cysteine in the N-terminal region, conserved with bacterial phytochromes; but also a second one in the GAF region. The second cysteine is conserved with cyanobacteria and plant phytochromes, but also with *Ectocarpus siliculosus* phytochrome 1, which binds phycocyanobilin and show a far-red/green photocycle (Rockwell et al., 2014). This suggested that different DPH copies could bind different chromophore and have different light-sensing abilities in *A. coffeaeformis*.

To address this question, we cloned the 4 *A. coffeaeformis* phytochromes photosensory domains and expressed them in *E. coli* with the enzymes producing either biliverdin (BV), phytochromobilin (PΦB) or phycocyanobilin (PCB).

<figure><img src="image_59.png" /><figcaption>Figure 1. Absorption spectra and difference spectra between the Pr and Pfr forms for 4 *A. coffeaeformis* phytochromes. A. AcDPH1 with BV; red lines is the absorption spectra after illumination at 810nm and dark red lines correspond to illumination at 630nm. B AcDPH3 expressed with BV, red line is illumination at 740nm and dark line illumination at 66nm. C. AcDPH4 expressed with PCB, red line illumination at 740nm and dark red line is illumination 666nm. In D, AcDPH2 production (with BV) was very low, and the spectrum was only taken under white illumination. E to G, Difference spectra between far-red illuminated and red-illuminated AcDPH (E-AcDPH1, F-AcDPH3 and G-AcDPH4). In F, the comparison of the difference spectra between AcDPH3 expressed with BV (black) and PCB (red) is also presented, normalized to 1 for the maxima.</figcaption></figure>

AcDPH1 seems to bind BV, and has an absorption spectra close to the one of PtDPH and other known DPH (see Chapter2), with peaks at 687nm for Pr form (810nm illumination)
and 742nm for Pfr-enriched mix (630nm illumination) (Fig1). For reference, PtDPH
Pr/Pfr and TpDPH Pr/Pfr have peaks at 700/750nm and 686/764nm respectively.

AcPDH3 also seems to bind BV, as the absorption spectra of AcDPH3 expressed with PCB shows the same absorption peaks (Fig1F). However, AcDPH3 absorption spectrum is shifted towards shorter wavelength compared to AcDPH1 and PtDPH: the maxima and the minima of the absorbance difference are at 669 and 712nm, respectively.

AcDPH4 was binding PCB, but did not exhibit photocycle as other DPH. Absorption shows a maximum at 653nm, with only a slight bleaching at 686nm upon illumination at 740nm. This effect is photoreversible by illuminating at 666nm after irradiation at 740nm. AcDPH2 could not be produced in high amounts, but seems to show the same absorption peak as AcDPH4 (Fig1D).

With the analysis of these spectra, we can conclude that *A. coffeaeformis* DPH seem to be all red (R)/far-red (FR) phytochromes with light reversible photoconversion but have different spectral properties: AcDPH1 possesses conserved absorption spectra with known PtDPH, while AcDPH3 is shifted towards shorter wavelengths. AcDPH4 seems to have peculiar photocycle, is binding another chromophore (phycocyanobilin) and is expressed only in some conditions, suggesting that it might regulate different aspects of diatom life (Keeling et al., 2014). Indeed, AcDPH4 is more expressed in the presence of acetate in the diatom growth media, suggesting a link with the heterotrophic metabolism of *Amphora* species (Lewin and Lewin, 1960). We can notice that the wavelength difference between the absorption peak maxima of the Pr and Pfr forms seems smaller than for the other diatom phytochrome (43 nm for AcDPH3 (669/712nm) vs 78 nm for TpDPH (686/764nm) for example) and leading to a Pr/Pfr photoequilibrium rich in Pr form whatever the light used. Globally, this data suggests some spectral tuning of the different AcDPH, which might be linked to the benthic environment. This was not observed in the different oceanic and coastal centric diatoms DPH (chapter 2), and this is also different from the absorption spectra observed in *Ectocarpus siliculosus* (Rockwell et al., 2014).
DPH regulates red-light acclimation response in *P. tricornutum*

Sediments but also mats or biofilms are characterized by a strong depletion in green and blue light, resulting in red and far-red enrichment due to either wavelength-dependent light scattering or strong filtering of light by absorption by chlorophyll (Kühl and Jorgensen, 1992; Kühl et al., 1994; Kühl and Jorgensen, 1994). Some cyanobacteria have evolved a specific chromatic adaptation to enhance light harvesting in this red/far-red light environments called Far-Red Light Photoacclimation (FaRLiP) (Gan et al., 2014). Chromatic adaptation is a general mechanism observed in cyanobacteria allowing an adaptation of the photosynthesis pigment to the environmental light (Grossman, 2003). In the case of FaRLiP, this phenomenon is under the control of a phytochrome (RfpA) and two response regulators (RfpB and RfpC) to activate the expression of genes involved in this acclimation (Zhao et al., 2015). Little is known in diatoms about chromatic adaptation, but one of the responses of *P. tricornutum* to red light is the appearance of a fluorescence peak around 710nm at room temperature and slightly shifted at 714 nm at 77K (hereafter F710) (Fujita and Ohki, 2004; Herbstová et al., 2015; Oka et al., 2020). We thus tested whether F710 and this acclimation could be under DPH control. As shown in Fig2, fluorescence spectra at 77K of *P. tricornutum* showed two peaks one at 680nm characteristic of PSII, but also another one at wavelength around 710nm (F710). The fluorescence spectrum of *P. tricornutum* has been further investigated under different light conditions (wavelength) and all fluorescence spectra curves have been normalized to the peak at 680 nm for qualitative analysis. F710 was induced by red light or in high cell concentration cultures grown without agitation in accordance with previous reports (Brown, 1967; French, 1967; Shimural and Fujita, 1973; Fujita and Ohki, 2004; Herbstová et al., 2015). With agitation, in green or green plus far-red light, F710 did not increase even at high cell concentration, but in red light with agitation, the far-red fluorescence was clearly induced suggesting a specific red light induction of F710 (Fig2). Therefore, light color and "agitation" (also linked to culture age/cell concentration) are parameters controlling the red-shifted forms of chlorophyll allowing F710.
<figure><img src="image_60.png" /><figcaption>Figure 2. Fluorescence emission spectra at 77K of *P. tricornutum* grown in different conditions. A, Cells grown in white light without agitation and sampled at different two cell concentrations; B, cell grown in red light with and without agitation, sampled at ~2e⁶C/mL, C cells at ~2e⁶C/mL grown with agitation but under different lights.</figcaption></figure>

In order to test a possible role of DPH in this red light acclimation, wild-type strain, transformation control lines (Wt (Tc), cells originated from the same transformed colony than its corresponding KO but not having undergone Pt-DPH gene editing as described in Fortunato et al, (2016)) and DPH-KO lines have been grown in the presence of red light but also with an increasing gradient of FR in order to vary the FR/R ratio, FR light being a light which activate PtDPH. Interestingly, we observed a FR and DPH-dependent regulation of F710 in red+far-red light (Fig 3 and FigS1). Indeed in Wt strains, F710 increases when FR light (and FR/R ratio) increases. In the DPH KO, F710 is present, but does not change with increasing ratio of FR/R lights. This experiment was repeated once for another pair of Wt Tc and DPHKO, and not in the exact same conditions, but showed the same type of phenotype (FigS1). F710 is thus induced by red light, but further regulated by DPH in a FR-dependent manner.
<figure><img src="image_61.png" /><figcaption>Figure 3. DPH controls F710 amplitude in response to increased FR/R ratio. Fluorescence emission spectra at 77K of *P.tricornutum* grown with agitation in red light with different far-red intensities (Tc tranformation control)</figcaption></figure>

A kinetic follow-up of the appearance of F710 on cells grown at 735nm showed that this phenomenon took place after a few hours and that it was much faster and important on the wild type lines than on the KO lines (Fig4), confirming the potential role of DPH as a sensor of this light condition inducing the development of F710.

<figure><img src="image_62.png" /><figcaption>Figure 4. Kinetics of induction of F710. Cells were grown in blue light with agitation and transferred to FR light (735nm) at T0. F710 is the fluorescence emission at 77K at =711nm (normalized to fluorescence at 687nm).</figcaption></figure>
F710 response seems restricted to benthic diatom species

The presence of F710 is known in Pt and in *Nitzschia closterium* (Fujita 2004), but it was not seen in centric species (*Chaetoceros*, *Skeletonema*, *Thalassiosira*) except for *Detonula sp* (Jupin and Giraud, 1971; Fujita and Ohki, 2004; Lavaud and Lepetit, 2013), and not in all pennate species (not observed for *Nitzschia punktata*, (Fujita and Ohki, 2004)). Therefore, we wanted to explore diatoms ability to develop the F710.

<figure><img src="image_63.png" /><figcaption>Figure 5. F710 in centric species: Fluorescence emission spectra at 77K of *Thalassiosira pseudonana* and *Skeletonema costatum* grown with agitaiton under different ligth colors.</figcaption></figure>

As expected, the centric species tested did not show F710 induction (Fig5). Among the pennate diatoms tested, *Fragilariopsis cyclindrus* is a planktonic and/or ice associated diatom. In the conditions we tested, which are known to trigger F710 in *P. tricornutum*, i.e white or red light without agitation at high cell concentrations, no F710 could be observed.

The benthic diatom *A. coffeaeformis* showed a high induction of F710 in red light (peak of fluorescence at 707nm) even with agitation, and in white light without agitation, but the F710 in green light with agitation remained low. It thus seems to be regulated in the same way as F710 in *P. tricornutum*.

The other three diatom species were grown only in white light without agitation, and both *Navicula salinicola* and *Nitzschia inconspicua* show a fluorescence peak at 708nm and
712nm respectively, while *Nitzschia agnita* did not show any peak around 710nm under this conditions.

<figure><img src="image_64.png" /><figcaption>Figure 6. 77K fluorescence spectra of different pennate diatom species. A. *Fragilariopsis cylindrus* grown in red or white light without agitation, B. *A. coffeaeformis* grown in green, red or white light, with and without agitation, C to D, *Navicula salinicola*, *Nitzschia inconspicua* and *Nitzschia agnita* grown in white light without agitation.</figcaption></figure>

Therefore, different pennate species possess or not the ability to develop F710, and this
might be linked to their planktonic vs benthic lifestyle.

DISCUSSION

In this chapter, we focused on the role of phytochrome in benthic diatoms. We had previously seen that most of the pennate diatoms living in open water lack phytochrome and that on the contrary, pennate diatoms living in a benthic environment presented an
increase of their phytochrome number which could go up to 4 copies. In benthic
environment, the quality and quantity of light changes significantly from the open water,
which is dominated by blue/green light. One known red-induced phenotype in *P. tricornutum* is the adaptation of photosynthesis to red light. This is characterized by the
appearance of a peak in far-red light (around 710nm) in the cell's fluorescence emission
spectra. If the implication of DPH in such a phenotype has been suggested, it has not been
tested so far. We therefore explore the possible links between phytochromes, adaptation
to red light and life in the sediments.

F710 and life in the sediments

In our study, we reproduced known effects of light, cell concentration and agitation on the induction of F710. Interestingly, the effect of the absence of agitation and high cell concentration could mimic the conditions experienced in a biofilm. Indeed, cell concentration in a biofilm is very high, with cells stacked on each other (Cartaxana et al., 2011). This results locally in high O₂ and low dissolved inorganic carbon due to photosynthetic activity and self-shading by other cells (Kühl and Jorgensen, 1992; Cartaxana et al., 2016; Marques da Silva et al., 2017). In a lab culture without agitation, cells sediment at the bottom of the flask; cell growth results locally in very dense cell layer, probably causing self-shading and high O₂/low DIC and low nutrient availability. In the agitated cultures, we considered the gas concentration to be homogenous in the culture and equilibrated with ambient atmosphere, independent of cell concentration. At high cell concentration, one could suppose that self-shading is also occurring in agitated cultures, but this did not result in high F710, suggesting that gas equilibrium is a signal triggering F710.

Changes in light spectra due to high cell concentrations were not directly addressed in
this preliminary work where we used mainly monochromatic light but may be addressed
later. However, light is clearly an important signal for F710 development. Light intensity,
quality and fluctuation are known to control F710 in *P. tricornutum*: induction in low
white light (Brown, 1967) and intermittent light (Giovagnetti and Ruban, 2021), strong
induction in red and yellow light, and small increase in green compared to white and blue
(Oka et al., 2020). In our work, F710 is clearly induced by red light, independently of DPH. This could be an effect of low photosynthesis efficiency in red, which is also supported by the appearance of F710 in low light. Alternatively, another red-light photoreceptor could be involved, but this photoreceptor has not been identified by genomic studies (Chapter 1), and would be of a new, unknown type, or a known type with atypical light sensing properties. In sediment, the light field is enriched in red and far-red light at depth, especially in the presence of photosynthetic organisms. Developing F710, and therefore red-shifted FCP, might be a response to the red-enriched light field in sediments.

The exact nature of this long-wavelength emission F710 is still unknown. F710 is clearly linked to photosystem II (PSII) activity but is debated whether this is due to an extra Light harvesting complex (LHC) binding to PSII and dissipating energy as FR fluorescence, or if this is spill-over of PSII over PSI (Fujita and Ohki, 2004; Herbstová et al., 2015; Bína et al., 2016; Giovagnetti and Ruban, 2021). However, the first hypothesis seems more convincing. In (Herbstová et al., 2015), the author performed sucrose gradient followed by gel filtration fractioning of thylakoid membranes of cells grown in red of white light. PSI fraction shows limited FR fluorescence and only at 77K but not at room temperature, while another fraction showed both 77K and room temperature FR fluorescence. Analysis of this second fraction showed that it contains LHCs, and one of them, Lhcf15 is specific of red light growth conditions. Lchf15 seems to be a good candidate, as it was also found enriched in red-light grown cells compared to blue-acclimated cells in proteomic studies (Schellenberger Costa et al., 2013), and is transcriptionally induced upon 6 and 24 h of red light illumination (Valle et al., 2014) which is in accordance with our kinetic of induction of F710. We are currently trying to generate Lhcf15 knock out to validate (or not) the Lhcf15 hypothesis. If validated, we could explore the Lhcf family in diatoms to try to identify Lhfc15 homologs in other diatom species (database from Chapter 1) to assess if this antenna is restricted to benthic species. Lhcf phylogenetic trees exist (Herbstová et al., 2015; Bilcke et al., 2021; Kumazawa et al., 2022), but unfortunately without enough sequences from different diatom species to explore the link between Lhcf and the different environmental niches. One last comment is that the fitness advantage of developing F710 in red light is not established yet but to extend light-absorption abilities to longer wavelengths might play important roles in light adaptation of *P. tricornutum* and
benthic diatoms species. If we manage to control F710 induction, we could try to link F710 to other photosynthetic parameters and explore this question.

## DPH and the fine-tuning of F710

DPH clearly controls F710 levels in response to R and FR in *P. tricornutum*, although only two experiments were done. However, as many parameters control F710 (light, cell concentration, agitation), the experiments are difficult to reproduce with exactly the same F710 induction.

Interestingly, the pennate diatom *Fragilariopsis cyclindrus*, lacking DPH, did not produce F710 while, *A. coffeaeformis* showed F710 induction in the same conditions as *P. tricornutum*, suggesting that in this diatom, F710 could be regulated by the same mechanisms including by DPH. Interestingly, the two other pennate diatoms showing F710 are likely to possess phytochromes too. The only *Navicula* species studied so far possess two DPH, so it might also be the case in the *Navicula salinicola* strain used here. However, *Nitzschia* species do not always possess DPH: some do (*Nitzschia palea*, *sp*, *N. punctata* see Table1), while some don't (see Chapter 1, although it is only transcriptome data). Here, *Nitzschia agnita*, such as *Nitzschia punktata* in (Fujita and Ohki, 2004) did not show F710, while *Nitzschia closterium* (Fujita and Ohki, 2004) and *inconspicua* (this study) have this red shift antenna phenotype. The *Nitzschia inconspicua* strain we used is not the one for which the genome is available (Oliver et al., 2021), but the genome sequenced from a close strain contains 2 DPH genes, what make us think that it might also be the case for the strain we used. However, we cannot exclude that we did not find the conditions inducting F710 in *F.cylindrus* and *N.agnita*, suggesting different regulations in these strains.

As mentioned above, light in the sediment is enriched in red and far-red light. In addition, the FR/R ratio increases with depth in the presence of photosynthetic organisms. For example, in microbial mats, light below 700nm is quickly attenuated (Kühl and Jorgensen, 1992; Cartaxana et al., 2016), while far-red light is less affected. DPH could thus be activated when going under a layer of photosynthetic cells due to high FR/R ratio. Under
this condition, the photoequilibrium of DPH or the co-regulation of several DPH in species
such as *A. coffeaeformis* could be determinant for this chromatic adaptation. It is therefore
tempting to propose that DPH and F710 co-evolved in benthic diatoms, with DPH enabling
the fine-tuning of F710 in sediments enriched in far-red light.

If the link between F710 and Lhcf15 mentioned above is confirmed, this would enable the
exploration of diatom genomic database for co-occurrence of Lhcf15 and DPH.

DPH multiplication in benthic species

In *P. tricornutum*, only one phytochrome is present and was shown here to control F710.
However, several DPH are present in other benthic diatom species and could control
F710. In *A. coffeaeformis*, we have shown here that the different DPH have different light-
sensing abilities and are induced under different growth conditions, suggesting that they
might have different functions. The strong conservation of AcPH1 spectra with PtDPH and
centric DPH from Chapter 2, led us to propose that all benthic diatoms have such a DPH
with light sensing in the red and far-red, that could control F710. AcDPH knock out are
being generated in the lab by CRISPR-Cas9 and will allow us to address this question.

DPH could control other aspects of diatom life in the sediments. Interestingly, *A. coffeaeformis* is able to grow in the dark in the presence of glucose, and to respire nitrates in the dark (Lewin and Lewin, 1960; Kamp et al., 2011). Other benthic species also show metabolic adaptation to life in the sediments (Lewin and Lewin, 1960). In our study and (Keeling et al., 2014), we saw high expression of AcPH4 in the presence of acetate, which hints towards a link between DPH and carbon metabolism in *A.coffeaeformis*. However, growth did not seem to be higher in the presence of acetate, suggesting that acetate might be a signal rather than a carbon source for heterotrophic growth. Testing growth and photosynthesis parameters with and without acetate and red light will provide interesting answer to this question.

Phototaxis is another behavior important for diatom life in the sediments (Barnett et al., 2020). A phototaxis assay for *A. coffeaeformis* is being developed in the lab and could be used to test the effect of different wavelengths and the implication of DPH(s).
As a conclusion, some photosynthetic adaptations seem to be specific for the benthic lifestyle of pennate species, and are likely to be regulated by DPH. In addition, the multiplication of DPH in these diatoms suggests the importance of light-regulation of diatom life in this environment enriched in red and far-red lights.

# MATERIAL AND METHODS

## Diatom culture conditions

Wild-type *P. tricornutum* (Pt1 8.6; CCMP2561) cells, PtDPH-KO mutants and transformation controls (Fortunato et al., 2016) were maintained in Enriched Artificial Seawater without silica; *Amphora coffeaeformis*, *Fragilaropsis cylindrus*, *Navicula salinicola*, *Nitzschia inconspicua* IRTA_CC1 and *Nitzschia agnita* were cultured with Si. All except *Fragilariopsis cylindrus* were grown at 19 °C under a 12L/12D regime using 50 µmol photons.m⁻².s⁻¹ white light (Philips TL-D De Luxe Pro 950) (“White condition”) or in continuous light in incubators equipped with green (520nm), red (670nm) or far-red (800nm) LEDs, with agitation at 160rpm. For the FR gradient, flasks were homogeneously illuminated from above with LED at 660nm at 25 µmol photons/m²/s, and from the side with FR LED at 765nm or at 810 nm, resulting in a gradient of FR light (from 30 to 250µmol photons/m²/s for the LED at 760nm and 30 to 170 µmol photons/m²/s for 810nm). *Fragilariopsis cylindrus* was grown at 4°C with a white spot under a 12L/12D regime, and covered with a red filter for red-light growth. For F710 induction kinetics, cells were first grown in blue light (450nm, 10µmol photons/m²/s) until similar fluorescence spectra and a cell concentration around 2.10⁶ cells/mL were obtained. The cultures were then placed at 735nm (40µmol photons/m²/s) to follow the induction of F710.

## AcDPH cloning and expression

Photosensory module (PSM) of DPH genes were cloned from *A.coffeaeformis* cDNAs grown in different conditions. Sequences were amplified with the primers pairs indicated in the Table S2, and cloned into pET28a-HO vector already containing *Synechocystis* Heme
oxygenase gene or pET28a-pcyA containing SynHO1 and pcyA from *Synechocystis* PCC6803. PSM sequences were expressed as N-terminal His6 tagged proteins in *E. coli* BL21 (DE3) strain, by auto-induction system (Studier, 2005). Recombinant proteins were purified as in Fortunato et al. (2016) and absorption spectra of the recombinant proteins were measured immediately after purification on a Varian's Cary-50 spectrophotometer. Illumination with LEDs at 810, 740, 666 and 630 nm were performed (approx. 1min illumination) to reach pure Pr spectra (after far-red illumination) or equilibriums between the Pr and Pfr forms.

## 77K Fluorescence

Low temperature (77 K) fluorescence emission spectra were measured with an in housebuilt setup. Samples were immersed in liquid nitrogen and excited with a LED source (LS-450, Ocean Optics - blue LED, 450 nm). The emission spectra were recorded using a CCD spectrophotometer (QE6500, Ocean Optics).

## SUPPLEMENTARY FIGURE:

<figure><img src="image_65.png" /><figcaption>Figure S1. DPH controls F710 amplitude in response to increased FR/R ratio. Fluorescence emission spectra at 77K of *P.tricornutum* grown with agitation in red light with different far-red intensities (Tc transformation control). Compared to Fig3, the strains are an other pair of DPHKO and its transformation control, and the FR light used is a LED at 810nm.</figcaption></figure>
<table><caption>Table S1. Primers used to clone A. *coffeaeformis* DPHs</caption><thead><tr><td>Names</td><td>Sequence</td></tr></thead><tbody><tr><td>AcDPHY1_Fw_Spel</td><td>gcgACTAGTATGACCATGCCTCCTGACGAAG</td></tr><tr><td>AcDPHY1_PCD_Rv_Sacl</td><td>gcgGAGCTCCTATCCAACAAGGAAAAGTATC</td></tr><tr><td>AcDPHY2_Fw_Spel</td><td>gcgACTAGTATGACAATCTTTCAGAAAATTGG</td></tr><tr><td>AcDPHY2_PCD_Rv_Sacl</td><td>gcgGAGCTCCTAATCAATGGCCGAGTAATAC</td></tr><tr><td>AcDPHY3_Fw_Spel</td><td>gcgACTAGTATGGCTCCTCCTGACTCCATGGAAG</td></tr><tr><td>AcDPHY3_PCD_Rv_Sacl</td><td>gcgGAGCTCCTATTCGAGTGCGTCAAAGCTTCGGATATTTG</td></tr><tr><td>AcDPHY4_Fw_Spel</td><td>gcgACTAGTATGGCTCCACCTCCTAAGGATATC</td></tr><tr><td>AcDPHY4_PCD_Rv_Sacl</td><td>gcgGAGCTCCTACATCGCCCGCATTGAAGCTCTT</td></tr></tbody></table>
CHAPTER 4: DPH SIGNALIZATION

In this last part, we explored different levels of the DPH light-sensing mechanisms, from chromophore binding to the integration of several environmental cues into the DPH signaling cascade. For this part, I searched for putative chromophore synthetizing enzymes in *P. tricornutum* genome, and tested some by cloning and co-expressing them with DPH in *E. coli*. I used *P. tricornutum* transgenic lines expressing tagged version of the PtDPH to look at DPH localization. I also addressed the question of the signalization cascade by Yeast two hybrid. I cloned DPH (full length or fragments) into Y2H vectors to perform an interaction screen with an available Pt Y2H library. I finally turned to a candidate approach with the only histidine phosphotransferase found in *P. tricornutum* genome PtHpt, to test its interaction with PtDPH. Finally, I characterized the effect cell concentration on DH-mediated response in *P. tricornutum*.
In the previous chapter, I have explored DPH evolutionary history, modeled its activity *in vivo* and projected it *in situ* and started to explore potential DPH-regulated phenotypes in model species.

In this last chapter, I will explore DPH signaling-related mechanisms, i.e. the assembly of the apoprotein with its chromophore, DPH subcellular localization, and components of the signaling cascade triggered by DPH. I will also present evidence of the co-regulation of PtDPH-controlled genes by other factors. Each of these aspects aims at broadening our knowledge about DPH light-sensing in diatoms.

RESULTS

DPH chromophore

Phytochrome chromophores are all linear tetrapyrroles, also called bilins (Rockwell and
Lagarias, 2017). Bilins derive from the oxidative cleavage of heme by heme oxygenase to
form biliverdin (IXα)(BV), which can be further reduced by ferredoxin-dependent bilin
reductases (BVRs), into phycocyanobilin (PCB), phytochromobilin (PB), phycourobilin
(PUB) or phycoerythrobilin (PEB) (Fig. 1) (Dammeyer and Frankenberg-Dinkel, 2008).

Bacteriophytochromes (Bph) bind BV (Bhoo et al, 2001; Karniol et al, 2005). In bacteria, the heme oxygenase gene is often found in the same operon as the *Bph* gene (Bhoo et al, 2001, Giraud et al, 2002). BV was found to bind a conserved cysteine in the N-terminal extremity of the apoprotein Bph and fungal phytochromes (Lamparter et al, 2002; Karniol et al, 2005; Blumenstein et al, 2005)

Land plant phytochromes bind phytochromobilin (PΦB), which is produced by the
enzyme HY2 in a one-step process (Lagarias and Rapoport, 1980), while cyanobacteria
bind phycocyanobilin produced by the enzyme PcyA (Hughes t al, 1997; Frankenberg et
al, 2001). In both cases, the chromophore is covalently bound to phytochrome apoprotein to a conserved cysteine in the GAF domain (Wagner et al, 2005).

PUB has been identified as chromophore of some cyanobacteriochromes (Sun et al, 2014), and as pigments of phycobilisome light-harvesting complexes, as the phycoerythrobilin produced by the enzymes pebA and pebB, in most red microalgae and in Cyanophyceae organisms (Dammeyer and Frankenberg-Dinkel, 2008).

<figure><img src="image_66.png" /><figcaption>Figure 1. Phytochrome chromophores biosynthesis pathways (from Rochkwell and Lagarias 2017)</figcaption></figure>

In plants, the phytochrome chromophore biosynthesis enzymes are located in the chloroplast, while in fungi it has been shown that heme oxygenases were attached to the mitochondrial outer membrane (Muramoto et al, 1999; Streng et al, 2021). Diatom phytochromes display a structure similar to the ones of bacteriophytochromes, with a conserved cysteine in the N-terminal part of the protein, and Fortunato et al (2016) showed that recombinant *Phaeodactylum tricornutum* and *Thalassiosira pseudonana* DPH covalently bind BV as chromophore. PtDPH expressed with the enzymes for production of РФВ or PEB had the same absorption spectra as with BV, suggesting that BV is the preferentially bound chromophore, and therefore possibly the native one. However, this
has not been demonstrated, and it has also been reported that some DPH from the diatom *Amphora coffeaeformis* possess an additional cysteine residue in the GAF domain conserved with the one used in plant and cyanobacterial phytochromes to bind their chromophore, suggesting that different chromophores could be synthetized in diatoms (Fortunato et al, 2016)(see Chapter3).

In this first part, we will explore the possible DPH chromophore biosynthesis pathways by mining genome for HO and BVR homologous genes, and testing their activity, as well as the reconstitution of chromophoric recombinant DPH.

## Heme oxygenase

Candidate heme oxygenase genes have been previously identified in *P. tricornutum* and *T. pseudonana* genomes (Fortunato et al., 2016). Here, we confirmed the presence of the Heme oxygenase domain (PF01126.2) (Table 1), and that no other protein in Pt genome possesses this domain. We also explored the predicted localization of these enzymes with ASAFind (Gruber et al, 2015), HECTAR (Gschloessl et al, 2008) and SignalP5 (Almagro Armenteros et al, 2019) software. Two of these proteins seem to be close to plant and cyanobacterial HO1 and are predicted to be addressed to the chloroplast. One enzyme is closer to mammalian HO, while the other did not have any good hit in Swissprot database (best blast hit had an e-value of 4.2). Both the plant/cyanobacteria-like and the mammalian-like HO had homologs in *T. pseudonana* and in *Amphora coffeaeformis* (Ac) (Table 1).

<table><caption>Table 1 Characteristic of the candidate Heme oxygenase in *Phaeodactylum tricornutum*</caption><thead><tr><th>Pt V3 Id<br>Phatr3_</th><th>Best blast hit in<br>swissprot</th><th>Blast<br>e-value</th><th>Pfam<br>(PF01126.2)<br>evalue</th><th>SignalP5</th><th>HECTAR</th><th>ASAFind</th><th>Tp<br>Thaps3_</th><th>Ac<br>CAMPEP_</th></tr></thead><tbody><tr><td>HO1<br>J12588</td><td>P09601.1<br>Heme oxygenase 1<br>[Homo sapiens]</td><td>1.13E-52</td><td>4.80E-55</td><td>other</td><td>other<br>localisation</td><td>not plastid</td><td>17865</td><td>0186525012<br>0186524814</td></tr><tr><td>HO2<br>J5851</td><td>Q69XJ4.1<br>Heme oxygenase 1,<br>[Oryza sativa]</td><td>1.09E-50</td><td>1.20E-07</td><td>signalP</td><td>chloroplast</td><td>not plastid</td><td>17854</td><td>0186534672</td></tr><tr><td>HO3<br>J35647</td><td>No good hit</td><td>4.20E-10</td><td>other</td><td>other<br>localisation</td><td>other<br>localisation</td><td>not plastid</td><td></td><td></td></tr><tr><td>HO4<br>J5902</td><td>Q8YVS7.1<br>Heme oxygenase 1<br>[Nostoc sp. PCC 7120]</td><td>9.11E-58</td><td>2.00E-56</td><td>signalP</td><td>chloroplast</td><td>plastid, high<br>confidence</td><td></td><td></td></tr></tbody></table>
We successfully cloned *PtHO2*, *PtHO3* and *PtHO4* from cDNA. As PtDPH is difficult to produce, we cloned these enzymes for recombinant expression in *E. coli*, along with *T. weissflogii* DPH Photosensory module (PSM) (TweiDPH), which is easily produced in high amounts and thus more suitable for screening (Fig3).

Only the strain expressing *PtHO2* produced high amounts of functional TweiDPH. Pellets of bacteria expressing *PtHO3* or *PtHO4* with TweiDPH were uncolored, suggesting that they did not produce the suitable chromophore (or that the enzymes were not functional in *E. coli*). TweiDPH produced with *PtHO2* could reversibly photoconvert to Pr upon far-red (765nm) light illumination, and had the same absorption spectra as TweiDPH when produced with *Synechocystis PCC6803* heme oxygenase 1 (SynHO1) (Figure 2). This suggests that *PtHO2* produced the same chromophore as SynHO1, which is efficiently incorporated in DPH and enables photoconversion.

<figure><img src="image_67.png" /><figcaption>Figure 2. Spectral properties of TweiDPH expressed in *E.coli* with *Synechocystis* HO1 (SynHO1), producing biliverdin, or *PtHO2*. A Absorption spectra of TweiDPH at room (white) light (black line) or after illumination by a far-red LED (765nm) (red line), normalized to Pr peak, in red. B. Difference in absorption spectra for recombinant TweiDPH produced with SynHO1 or *PtHO2*.</figcaption></figure>

Therefore, *PtHO2* is able to produce BV for phytochrome activity *in vivo*; *PtHO2* knock-out mutants are planned to be generated by CRISPR-Cas9 in *P. tricornutum*, to test whether or not this enzyme is required for phytochrome-mediated gene expression.
<figure><img src="image_68.png" /></figure>

Figure 3 Scheme of the vector structure allowing to co-express T. weissflogii DPH photosensory module along with the putative enzymes of chromophore biosynthesis pathway identified in P. tricornutum genome. The backbone vector used was the pET28a, and the different coding sequences were differentially combined via sequential insertions according to the restriction sites indicated.

## Biliverdin Reductases

Putative bilin-reductase genes have been identified in *P. tricornutum* genome, raising the possibility of other chromophores synthetized in diatoms. Among these 4 candidates, 2 were predicted to harbor a peptide signal by the 3 software and possess as best hit counterparts in cyanobacteria and plant (Table 2).

These different BVR were amplified from cDNA and recombinantly expressed with *Synechocystis ho1* and *TweiDPH* as polycistronic sequence (Fig3).

When *TweiDPH* was expressed with *PtBVR3* and *4*, the absorption spectra were the same as with *SynHO1* (Fig3A), suggesting that these phytochromes were not binding a new chromophore, but bound BV. However, when *TweiDPH* with *PtBVR1* or *PtBVR2*, a new absorption peak appeared around 620nm, suggesting that the DPH was binding another chromophore in addition or in place of BV. However in both cases, this peak did not show any photochemistry (Fig3B and C). As this DPH is produced in high amounts, it is possible that it binds non-optimal chromophores. Zinc induced fluorescence gel will tell if this new chromophore is covalently bound to the protein, and absorption spectra of the denatured protein will give indication on the nature of this chromophore.

Overall, alternative chromophores produced by *BVR1* and *BVR2* could be possible alternative chromophore but it seems that BV is the most convincing as DPH native chromophore.
<table><caption>Table 2 Characteristic of the candidate BVR in *P. tricornutum*</caption><thead><tr><th>Accession</th><th>best blast hit in swissprot</th><th>evalue</th><th>Pfam domains</th><th>SignalP</th><th>HECTAR</th><th>ASAFind</th><th>Tp Thaps3</th></tr></thead><tbody><tr><td>BVR1 J10830</td><td>Q7NL66.2, 15,16-dihydrobiliverdin:ferredoxin oxidoreductase Gloeobacter violaceus PCC 7421]</td><td>4.00E-51</td><td>Ferredoxin-dependent bilin reductase (PF05996.15) 1.6e-53</td><td>signalP</td><td>signal peptide</td><td>Plastid, low confidence</td><td>32431</td></tr><tr><td>BVR2 J33770</td><td>Q93TL6.2, 15,16-dihydrobiliverdin:ferredoxin oxidoreductase [Nostoc punctiforme PCC 73102]</td><td>4.00E-39</td><td>Ferredoxin-dependent bilin reductase (PF05996.15) 9.2e-47</td><td>other</td><td>other localization</td><td>Not plastid, SignalP negative</td><td>32431</td></tr><tr><td>BVR3 J42659</td><td>Q8H124.1, Uncharacterized protein At2g34460, chloroplastic [Arabidopsis thaliana]</td><td>8.00E-16</td><td>NAD(P)H-binding (PF13460.9) 6.7e-39</td><td>signalP</td><td>signal peptide</td><td>Plastid, high confidence</td><td>264007</td></tr><tr><td>BVR4 J45446</td><td>Q10UW6.1, Phycoerythrobilin:ferredoxin oxidoreductase [Trichodesmium erythraeum IMS101]</td><td>1.00E-30</td><td>Ferredoxin-dependent bilin reductase (PF05996.15) 1.5e-35</td><td>other</td><td>signal peptide</td><td>Not plastid, SignalP positive</td><td>262396</td></tr></tbody></table>

<figure><img src="image_69.png" /><figcaption>Figure 4. Spectral properties of TweiDPH expressed in *E. coli* with SynHO1 and different PtBVR, in white light; B TweiDPH in R of FR with PtBVR1, C TweiDPH in R and FR with PtBVR2</figcaption></figure>

We have shown that *P. tricornutum* possesses at least one enzyme able to synthesize biliverdin as chromophore of phytochrome, and at least two enzymes are able to further
process BV. The nature and the role of this pigment in phytochrome activity need to be further investigated.

## DPH LOCALIZATION

Phytochrome localization is an important parameter for DPH signaling. In plants, phytochromes translocate to the nucleus upon light illumination, where they interact with transcription factors (Legris et al., 2019). This seems to be the case in green algae too, as *Micromonas pusilla* phytochrome is shuttled to the nucleus during the light phase of the day (Duanmu et al., 2014). In Fungi, FphA of *Aspergillus nidulans* interacts with the first component of its signaling cascade in the cytoplasm, and it is one of the downstream signaling component that is shuttled to the nucleus upon phytochrome light activation (Yu et al., 2016). Other studies also showed that FphA is interacting with blue light signalization components in the nucleus (Purschwitz et al., 2008).

We thus tested DPH subcellular localization in *P. tricornutum*. PtDPH was fused with Venus or NeonGreen fluorescent proteins either in N-terminal or C-terminal and expressed under the strong promoter of the *LHCF2* antenna gene in the episome system. Cells were grown in green light (DPH-inhibitory) and exposed to FR light to investigate eventual light-induced re-localization.

<figure><img src="image_70.png" /></figure>

Figure 5. DPH localization. Venus-PtDPH-expressing *P. tricornutum* cells grown in green light (A to F), or far-red light (G to L). Bar scale: 5µm. A and G, Bright field; B and H, Chlorophyll autofluorescence; C and I, Hoescht fluorescence (nucleus staining, also showing chlorophyll fluorescence); D and J, Hoescht and chlorophyll overlay; nucleus appears blue when chloroplast appears magenta; E and K, Venus fluorescence (PtDPH localization); F and L, overlay.
<figure><img src="image_71.png" /><figcaption>Figure 6. DPH localization. PtDPH-NeonGreen-expressing *P. tricornutum* cells grown in green light (A to F), or far-red light (G to L). Bar scale: 5µm. A and G, Bright field; B and H, Chlorophyll autofluorescence; C and I, Hoecsht fluorescence (nucleus staining, also showing chlorophyll fluorescence); D and J, Hoeschst and chlorophyll overlay, nucleus appears blue when chloroplast appears magenta; E and K, Venus fluorescence (PtDPH localization); F and L, overlay.</figcaption></figure>

PtDPH seems to be cytoplasmic (Fig4 and 5), although it is difficult to affirm with certainty that PtDPH is absent from the nucleus. No major changes were observed upon FR illumination. We thus concluded that PtDPH is present in the cytoplasm.

As control, lines expressing only the Venus fluorescent protein (Fig7), or the fusion Venus-PtCPF1 (Fig8, previously shown to be addressed to the nucleus (Coesel et al, 2009)) under the same regulatory sequences and episomal vector, were observed and display cytoplasmic and nuclear localization, respectively.

<figure><img src="image_72.png" /><figcaption>Figure 7. Venus localization. Venus-expressing *P. tricornutum* cells grown in white light (A to F). Bar scale: 2µm. A, Bright field; B, Chlorophyll autofluorescence; C, Hoecsht fluorescence (nucleus staining, also showing chlorophyll fluorescence); D, Hoeschst and chlorophyll overlay, nucleus appears blue when chloroplast appears magenta; E, Venus fluorescence; F, overlay.</figcaption></figure>
<figure><img src="image_73.png" /><figcaption>Figure 8. PtCPF1 localization. Venus-PtCPF1-expressing *P. tricornutum* cells grown in white light (Bar scale: 2 µm. A, Bright field; B, Chlorophyll autofluorescence; C, Venus fluorescence (PtCPF1 localization in the nucleus); F, overlay.</figcaption></figure>

DPH SIGNALISATION PATHWAY

Phytochrome signalization is very different in plants compared to bacteria and fungi. In plants, phytochromes are shuttled to the nucleus upon light activation, where they interact with Phytochromes Interacting Factors, which are bHLH transcription factors. It has been described for several fungi and bacteria phytochromes that signaling cascade consists in a two-component phosphorelay system (Yeh et al, 1997; Bhoo et al, 2001; Giraud et al, 2005; Brandt et al, 2008). These systems start with the activation of a receptor histidine kinase (here, phytochrome), which phosphorylates a response regulator (REC) domain. In a simple two-components system, the response regulator directly regulates the functional response (for example gene expression). In multi-step system, the receptor histidine kinase is hybrid, as it also contains a REC domain. Upon activation, there is phosphorylation of this REC domain, then the phosphate group is transmitted to a phosphotransfer protein (characterized by a Hpt domain), which will bring the phosphate group to the effector response regulator (Fig9).
<figure><img src="image_74.png" /><figcaption>Figure 9. Elements from two-component systems, from (Schaller et al., 2011)A, simple two-component system, B, Multi-step phosphorelay where the hybrid histidine kinase contains both a histidine kinase domain and a response regulator. Phosphorylated residues Histidine (H) and Aspartate (D) are indicated.</figcaption></figure>

Bacteriophytochromes show both simple and multi-steps two-component systems, and in most of the cases reported, the signalization components are encoded in the same operon as the *Bph*, which facilitates the determination of the downstream components (Fig9) (Yeh et al, 1997; Bhoo et al, 2001; Giraud et al, 2005; Vuillet et al, 2007; Jaubert et al, 2008). In fungi, FphA from *Aspergillus nidulans* interacts with the Hpt-containing protein YpdA, which itself controls the REC-containing SskA protein (Yu et al., 2016). Downstream of SskA, a MAPKinase cascade results in the phosphorylation of SakA, which is shuttled to the nucleus where it regulates the transcription factor AtfA (See also in the Introduction).

In diatoms, we know PtDPH autophosphorylates upon far-red light irradiation *in vitro* (Fortunato et al., 2016), which would be compatible with a two-component system phosphorelay as in Bacteria and Fungi.

I performed Yeast-two hybrid (Y2H) screens to find candidate interactors for PtDPH. I used a Y2H PtcDNA library from (Huysman et al., 2013), and performed the screen with PtDPH full length protein (one time) or PtDPH output module only (PtDPH_OPM, screen was performed 3 times) as baits, fused to GAL4 binding domain. Despite the identification of some candidates during the screens, we were unable to reproduce these interactions
when re-transforming yeast strains with the candidates vectors, suggesting that these
were not true interactants.

<figure><img src="image_75.png" /><figcaption>Figure 10. DPH-OPM interacts with PtHpt in Yeast Two Hybrid. Yeast spot growth on media selecting for the presence of 2 vectors (-LW) and on the protein interaction (-LWH). 3AT, which inhibits HIS3, was added to test the strength of the Hpt-DPH-OPM interaction. Last 3 columns are negative controls to test for prey or bait autoactivation.</figcaption></figure>

I then turned towards a candidate gene approach. By looking for Histidine
phosphotransferase protein in diatom genomes, we found only one protein with the Hpt
domain (PF01627) in *P. tricornutum*. This protein is present in one copy in the diatom
*Thalassiosira pseudonana*, but also in diatoms with no phytochrome (*Thalassiosira
oceanica*, *Fragilariopsis cylindrus*). I tested the interaction of PtDPH with this protein by
Y2H, using PtHpt as bait, fused to Gal4 binding domain, and PtDPH FL (full length), PCD
(Photosensory domain), OPM (Output module) or just PtDPH REC (Response regulator)
domain as preys (fused to Gal4 Activating domain, AD). I showed that PtHpt interacts with
PtDPH OPM, but not with the other PtDPH fragments nor with the FL protein (Fig10). This
is not surprising, as *Arabidopsis thaliana* PhyA and B FL protein interaction with SAP1 is
light-dependent in Y2H, while AthaPhyA and B OMP recapitulates all interactions (Lu et
al., 2015). We will test light-dependent interaction by adding BV chromophore to the
media and grow yeast strains under different lights. We did not detect the interaction with
the reverse combination (Hpt-AD with PtDPH FL, PCD, OPM or REC-BD), maybe because
of the absence of a nuclear localization signal in the GAL4BD fusion.

I also performed Y2H screen with PtHpt as bait, but once again we were unsuccessful in re-transforming with the candidate plasmids. However, Pt genome contains only a limited number of proteins with a response regulator (RR) (16 proteins, Table 3) that could be the target of Hpt, and which could be tested by direct interaction tests as done here for Hpt. Two of these proteins are especially interesting, as they combine RR and the LuxR transcriptional regulator domain, which binds DNA (Table 3). However, in fungi the response regulator element is not the transcription factor and the end of the signaling cascade, which involves also a MAPKinase cascade. The signaling cascade for PtDPH signaling could therefore be longer than 2 steps. We are currently trying to confirm the involvement of PtHpt in DPH signaling by generating PtHpt KO by CRISPR-Cas9 in *P. tricornutum*. In addition, we will test the phosphorelay between PtDPH and PtHpt *in vitro*. Alternatively, we could also start with the promoters of DPH-regulated genes, to analyze eventual binding motives, test the minimal promoter size *in vivo* with the reporter system used in Chapter2, and perform a yeast one hybrid screen to identify transcription factors binding this promoter.

We have uncovered here a first good candidate as component of the signaling cascade initiated by DPH, even if work is needed to confirm it. Interestingly, Hpt is conserved across diatoms that do not possess DPH, suggesting that it could work as a signaling hub integrating different signals.
<table><caption>Table 3. Proteins with Response regulators domains in *P. tricornutum* genome (DPH not included)</caption><thead><tr><th>Phatr3 ID</th><th>PFAM Domains</th></tr></thead><tbody><tr><td>Phatr3_EG00123</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain;Histidine kinase-, DNA gyrase B-, and HSP9NA-like ATPase</td></tr><tr><td>Phatr3_EG01983</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain;Histidine kinase-, DNA gyrase B-, and HSP9NA-like ATPase;CHASE domain</td></tr><tr><td>Phatr3_EG02384</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain</td></tr><tr><td>Phatr3_EG02387</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain</td></tr><tr><td>Phatr3_J11994</td><td>Response regulator receiver domain;Bacterial regulatory proteins, luxR family</td></tr><tr><td>Phatr3_J12051</td><td>Response regulator receiver domain;Bacterial regulatory proteins, luxR family</td></tr><tr><td>Phatr3_J13255</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain;Histidine kinase-, DNA gyrase B-, and HSP9NA-like ATPase</td></tr><tr><td>Phatr3_J16861</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain;Histidine kinase-, DNA gyrase B-, and HSP9NA-like ATPase;GAF domain</td></tr><tr><td>Phatr3_J45485</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain;Histidine kinase-, DNA gyrase B-, and HSP9NA-like ATPase;PAS domain</td></tr><tr><td>Phatr3_J46628</td><td>Response regulator receiver domain;Histidine kinase-, DNA gyrase B-, and HSP9NA-like ATPase</td></tr><tr><td>Phatr3_J47689</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain;PAS fold;Histidine kinase-, DNA gyrase B-, and HSP9NA-like ATPase;PAS fold</td></tr><tr><td>Phatr3_J47930</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain</td></tr><tr><td>Phatr3_J55037</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain;PAS fold;Histidine kinase-, DNA gyrase B-, and HSP9NA-like ATPase;PAS domain;PAS domain</td></tr><tr><td>Phatr3_J55187</td><td>Response regulator receiver domain;His Kinase A (phospho-acceptor) domain;Histidine kinase-, DNA gyrase B-, and HSP9NA-like ATPase</td></tr><tr><td>Phatr3_J8970</td><td>Response regulator receiver domain;Leucine Rich repeat</td></tr></tbody></table>

Note: I identified these proteins as interesting candidates to look at in the new database for diatom gene expression, Diatomicbase, for which a manuscript is in preparation

## CO-REGULATION OF DPH-REGULATED GENES

We used the reporter strains from Chapter 2 to explore the regulation of DPH targeted genes by other signals. While setting up the conditions for the action spectra, we found that the YFP signal varies depending on the cell concentration (Chapter2). YFP signal per cell increased when cell concentration increased in ProHsf6.4a ::YFP reporter lines, when grown in green light without agitation (Fig11A). This effect appeared specific for *Hsf4.6a* promoter, as lines containing the YFP under the control of other promoters (ProLHCF2 (light harvesting antenna protein gene) and the ProH4 (histone H4)) did not show this kind of regulation. To understand how this effect affects DPH-dependent gene regulation,
we grew cells without agitation, and under green light supplemented or not with far-red
light (10 µmol photons/m²/s). At very low cell concentration, the YFP signal was very low
in both green and green plus far-red. From 1e⁵Cells/mL, YFP signal in green-light grown
cells increased. Adding FR light increases the YFP signal, and the FR-induction itself seems
to be cell-concentration dependent (Fig11B). The effect of cell concentration on cells
grown in green light was also seen in PtDPH KO strains (Fig11C), implying that this effect
is not DPH dependent. However, when cells were agitated (under green light), the
increase in YFP with cell concentration was abolished (Fig11C; also true for the Wt
background, data not shown).

Therefore, it seems that a signal other than light also regulates *Hsf4.6a* promoter. It could
be oxygen or CO₂ concentration in the media, as this effect appears when cell
concentration is high in non-agitated cultures. Further investigations need to be done to
decipher the different signals co-acting with PtDPH.
<figure><img src="image_76.png" /><figcaption>Figure 11: Regulation of ProHsf4.6a by light, cell concentration and agitation. A. YFP expression under different promoters at different cell concentrations. Mean YFP per cell in *P. tricornutum* lines expressing the YFP under the control of different promoters during growth in constant green light. LHCF2: Light harvesting protein, H4: Histone H4. B. Effect of cell concentration of the FR induction of ProHsf4.6a ::YFP (inset, same data on a non-logarithmic scale). YFP was normalized to the maximum levels observed under green light. No agitation was applied to the cultures C. Effect of agitation on ProHsf4.6a activity. DPH KO strains were grown in green light with or without agitation, and YFP levels are normalized to the maximum observed in green light without agitation.</figcaption></figure>
## CONCLUSION

In this chapter, we have gained insights into phytochrome signaling mechanisms. We propose that DPH binds BV produced by HO2, but that the chromophore is not further reduced by a BVR. DPH seems to be localized in the cytoplasm whatever its activation state, and interacts with the histidine phosphotransferase PtHpt, probably triggering a two-component phosphorelay. As PtHpt is conserved among diatoms, it could be a signaling hub integrating other signals such as the “cell concentration” signal, probably a gas pressure in the media, that regulates DPH-regulated genes. Overall, this resembles the situation in Fungi, where BV-binding phytochromes trigger a two-component signalization system that is also activated by osmolarity changes. As DPH and Fph are evolutionary close, these organisms might have found similar ways to integrate phytochromes in their cellular signalization network.

<figure><img src="image_77.png" /><figcaption>Figure 12 Proposition of DPH signaling mechanisms in *P. tricornutum* (cell concentration could act on Hpt, but also on independent signalisation pathways, and directly on DPH too; Hpt can shuttle to the nucleus, but is could also be one of the downstream components).</figcaption></figure>

## MATERIAL AND METHODS

### Diatom culture conditions

Reporter lines were the same as in Chapter 2. Cells were grown at 19°C under green light (530nm, 25µmol photon/m2/s) with or without agitation and far-red light (740nm at 10
µmol photon/m2/s) was added for 24h. YFP fluorescence was measured with a MACSQuant® Analyser (Myltenyi Biotech) flow cytometer with excitation by a 488nm laser and detection at 500-550nm emission filters.

## Phytochrome expression, purification and spectral analysis

Photosensory modules (PSM) of DPH genes were obtained from previous studies (PtDPH (Fortunato et al, 2016), TweiDPH from chapter 2). Sequences were cloned as indicated in the Table 3, into pET28a-HO vector generated by inserting the Synechocystis Heme oxygenase gene amplified from pKS270 vector (Mukougawa et al., 2006) with the primer pair SynHO1xpET.HindIII.Fw and SynHO1xpET.NotI.rv, in HinDIII/NotI in the pET28a vector. PtHO were cloned in SalI/NotI in the pET28a vector (see fig x and table 3 for the primers used, PtHO2, PtHO3 and PtHO4 were cloned without their peptide signal); PtBVR were cloned in NotI/XhoI in the pET28a vector (see Table S1).

PSM sequences were expressed as N-terminal His6 tagged proteins in E. coli BL21 (DE3) strain, by auto-induction system (Studier, 2005). Recombinant proteins were purified as in Fortunato et al. (2016). Absorption spectra of the recombinant proteins were measured after purification on a Varian's Cary-50 spectrophotometer. Illumination with LEDs at 810 and 630 nm were performed (approx. 1min illumination) to reach pure Pr spectra (after 810 illumination) or equilibriums between the Pr and Pfr forms.

## Phytochrome localization

Episomal vectors carrying fusion of PtDPH with fluorescent proteins were generated with the modular cloning system uLoop kit containing *P. tricornutum* regulatory sequences level 0 cloning block along with fluorescent reporter proteins and conjugative element (Pollak et al, 2019; 2020). PtDPH and PtCPF1 coding sequences were amplified by PCR from cDNA and using the primer pairs : PtDPH.D.Fw an PtDPH.E.Rv, PtDPH.C.Fw an PtDPH.D.Rv, and PtCPF1.D.Fw and PtCPF1.E.Rv, respectively, and cloned into pL0 receiver vector (Pollak et al, 2019). Each of these pL0 blocks were assembled with the FcpB promoter and terminator sequences, and the Venus or NeonGreen fluorescent proteins as 5' or 3' fusion, to generate level 1 expressing constructs. Finally, these expressing units were assembled with conjugative elements to express them from an episomal vector and the Bleomycin resistance cassette. These final constructs were transformed into *E. coli*
DH5alpha containing the pTA-MOB (plasmid encoding the machinery required for conjugal transfer), and delivered into *P. tricornutum* cells by conjugation according to the the protocol described in Karas et al, 2015). Zeomycin 100µg/ml was used to select and grow conjugants.

Epifluorescence images were taken on an Axio Observer.Z1 inverted microscope (Zeiss) equipped with an ORCA-flash4.0 digital camera (Hamamatsu) and a Colibri.2 LED system (Zeiss) for excitation at 505 nm for Venus (520-550 nm emission) and 470 nm for chlorophyll autofluorescence (665-715 nm emission). Hoechst staining was performed by adding Hoeschst at a final concentration of 20µg/mL. To study eventual DPH re-localization, we illuminated the cells with FR light (765nm) while under the microscope.

## Yeast two hybrid

Phytochrome full length or domain (Photosensory Core Domain PCD, Output Domain (HK-ATPase-REC) or REC domain) sequences were amplified by PCR from plasmid containing the Pt-DPHPro:Pt-DPH-HA produced previously (Fortunato et al., 2016) and PtHpt was cloned from PtcDNAs with primers containing attB1 and attB2 GATEWAY recombination sites (see Table2). PCR products were introduced in the pDONR221 entry vector by BP GATEWAY recombination (Invitrogen). ENTRY clones were recombined in the pDEST32 vector (bait, for GAL4 DNA-binding domain N-terminal fusion, Invitrogen) or pDEST22 vector (prey, for GAL4 Activating domain N-terminal fusion, Invitrogen) by LR recombination. *Saccharomyces cerevisae* PJ69-4a cells were transformed using the LiAc method (Gietz and Schiestl, 2007). Co-transformed yeast were selected on YNB media (recipe ?) lacking tryptophane and leucine, and interaction between the proteins was assessed by spotting on plates lacking tryptophane, leucine and histidine. 3-amino-1,2,4-triazole, which inhibits the HIS3 enzyme (that enables growth without histidine) was added at 10mM to test the strength of the interaction. Screens were performed by transforming yeast bait strain (ie transformed with just the bait vector) with the prey library obtained from (Huysman et al., 2013) and plated on selective -LWH media.
<table><caption>Table S1 Primers data used in this study</caption><thead><tr><td>Name</td><td>Sequence</td></tr></thead><tbody><tr><td>PtHpt_J33969.Gateway.Fw</td><td>GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGAATCCACCCGATGTGATCAATTG</td></tr><tr><td>PtHpt_J33969.Gateway.Rv</td><td>GGGGACCACTTTGTACAAGAAAGCTGGGTCTCAACCGTAACAAAAAATTGTTTTC</td></tr><tr><td>PtDPH FL.Gateway.Fw</td><td>GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGAGCGGGGCAAATTATAGAG</td></tr><tr><td>PtDPH PCD.Gateway.Rv</td><td>GGGGACCACTTTGTACAAGAAAGCTGGGTCTCATGCCGCCAAATATTTCCGGTTT</td></tr><tr><td>PtDPH OMP.Gateway.Fw</td><td>GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGGGATTACTGAGAGGTGATA</td></tr><tr><td>PtDPH REC.Gateway.Fw</td><td>GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGTCGGGTCCGATTTTGATCG</td></tr><tr><td>PtDPH.Gateway.Rv</td><td>GGGGACCACTTTGTACAAGAAAGCTGGGTCTCATTCCGGAACCCGCAGCCTC</td></tr><tr><td>PtHO2_J5851.Sall.fw</td><td>gcgGTCGACAGAAGGAGATATACATATGTTCGCACCGGCCTTTTCGGTG</td></tr><tr><td>PtHO2_J5851.NotI.Rv</td><td>gcgGCGGCCGCTTAATTGTGGGGATGCTGAC</td></tr><tr><td>PtHO3_J35647.Sall.fw</td><td>gcgGTCGACAGAAGGAGATATACATATGCACTCGAACGACCCCCATG</td></tr><tr><td>PtHO3_J35647..NotI.Rv</td><td>gcgGCGGCCGCTCAAGATTTAATCATTCGTC</td></tr><tr><td>PtHO4_J5902.Sall.fw</td><td>gcgGTCGACAGAAGGAGATATACATATGTTTGCTCCTTCCGTCTCGAG</td></tr><tr><td>PtHO4_J5902.NotI.Rv</td><td>gcgGCGGCCGCTTACAATGCAACTCCAAGTTTTAG</td></tr><tr><td>J45446NotI.fw:</td><td>cgcggccgcagaaggagatataccatggccgaaccaacacagcttttg</td></tr><tr><td>J45446.Xhol.rv:</td><td>ggcgctcgagctatggcgatgggaatagcac</td></tr><tr><td>J10830.NotI.Fw:</td><td>cgcggccgcagaaggagatataccatgtacactaccggcaacagcggag</td></tr><tr><td>J10830.Xhol.rv</td><td>ggcgctcgagctaccgttgcaagctgctac</td></tr><tr><td>J33770.NotI.Fw:</td><td>cgcggccgcagaaggagatataccatggtccgcgcctaccag</td></tr><tr><td>J33770.Xhol.Rv:</td><td>ggcgctcgagctacttttgcgagagtggaaag</td></tr><tr><td>J42659.NotI.Fw:</td><td>cgcggccgcagaaggagatataccatgttccagcagtcctcggtttttgg</td></tr><tr><td>J42659.Xhol.Rv:</td><td>ggcgcctcgagttaaagcttcttgttggccac</td></tr><tr><td>PtDPH.D.Fw</td><td>aggctgtctcgtctcgtctcaggtctcaaggtatgagcggggcaaattatagag</td></tr><tr><td>PtDPH.E.Rv</td><td>tggtaatctatgtgtcctggtggtctctaagcctattccggaaccgcagcctc</td></tr><tr><td>PtDPH.C.Fw</td><td>aggctgtctcgtctcgtctcaggtctcaaatgagcggggcaaattatagag</td></tr><tr><td>PtDPH.D.Rv</td><td>tggtaatctatgtgtcctggtggtctctacctttccggaaccgcagcctctg</td></tr><tr><td>PtCPF1.D.Fw</td><td>aggctgtctcgtctcgtctcaggtctcaaggtatggctaaatcggaagag</td></tr><tr><td>PtCPF1.E.Rv</td><td>tggtaatctatgtatcctggtggtctctaagcttagttgcaggttgtcgc</td></tr></tbody></table>
DISCUSSION

During this work, I have tried to get insights into diatom photoperception, and more specifically on diatom phytochromes.

Despite the availability of genomic data, there was no complete view of photoreceptors in diatoms. The genomes of the two diatom models, *T. pseudonana* and *P. tricornutum*, had been exploited to find photoreceptors, but the other diatom genomes had not been systematically mined for photoreceptors.

In the first chapter, I started to fill this gap by mining diatom genomic data for photoreceptors. I extended the search to all Ochrophyta, and included also other Stramenopiles to try to get insights into the conservation of these photoreceptors. We looked for known light-sensing domains like the LOV domain, cryptochrome photolyase-related domain, rhodopsins and phytochromes. Overall, it seems that diatoms, (but also more widely, Ochrophyta), possess a common set of blue light photoreceptors (aureochromes, cryptochromes) while some possess additional photoreceptors such as rhodopsins and phytochromes. The photoreceptor repertoire appears specific for an algal group or even down to a species, suggesting an adaptive value of the additional photoreceptors in specific environmental niches.

Some of the results confirmed already known photoreceptors distribution, such as the fact that all diatoms have 4 aureochromes. We also uncovered a new group of putative LOV-based photoreceptors that is quite conserved amongst Ochrophyta, including in diatoms, but is not found in *P. tricornutum*, showing the importance of looking in all available data. It was found here in *T. pseudonana*, but was not reported before.

Another example of new insights from the study described in chapter 1 is the CryP family of diatoms. CryP was reported recently in *P. tricornutum* as a photo-sensitive
protein with roles in both DNA repair and gene regulation. We highlighted here a new
family, CryP2, which was already described in *T. pseudonana* (Oliveri et al., 2014;
Fortunato et al., 2015), but it was not known if CryP2 was a species-specific light sensor,
or something more conserved across diatoms. Based on the analysis in Chapter 1, it
appears now as a broadly conserved CryP subgroup, probably representing a new type of
photoreceptor with distinct properties.

We extended here the catalog for diatom photoreceptors, but we could also benefit
from the environmental data to complete this view. Of course, the identified proteins are
only candidate photoreceptors that would need to be biologically characterized.

Concerning phytochromes, we confirmed previous work (Fortunato et al., 2016), and
better define the distribution and occurrence of DPH within centric and pennate diatoms.
We did not resolve the origin of DPH, but mining other databases such as viral or bacterial
sequences in addition to Ochrophyta and other eukaryotes might give new insights.
Regarding centric group, investigations in genomic/transcriptomic resources (mostly
from coastal species) indicate that DPH is found across nearly all Thalassiosirales species,
suggesting that this is an ancient acquisition in all Thalassiosirales, with a few losses (for
example in *Thalassiosira oceanica*). Thalassiosirales DPH was also found in the oceanic
regions in the *Tara Oceans* dataset. Thalassiosirales are sister to a group of diatom
containing *Minutocellus* species, which possess DPH, and *Chaetoceros* species, which do
not. The absence of DPH in *Chaetoceros* species is well supported by the *Tara Oceans*
data: centric DPH genes found in the environmental dataset were very close to
*Thalassiosirales* or *Minutocellus* genes, and even in MAGs belonging to *Chaetoceros*, we did
not find phytochrome genes. Of course MAGs are incomplete, and the other genomic data
from Chapter 1 are mostly transcriptomes, but given the amount of transcriptomes in
different species and different growth conditions, we could have expected DPH to be
expressed in at least one of them. Looking for phytochrome in the *Chaetoceros* genomes
recently published (Hongo et al., 2021; Kumazawa et al., 2022), and the new *Chaetoceros*
genomes to come will definitely resolve the question of the absence of phytochromes in
*Chaetoceros* species.
The more basal centric diatom species *Coscinodiscus* also possesses DPH, however this gene is incomplete and it is difficult to assess its exact position compared to other DPH. No DPH from this species was found in the environmental data.

Pennate DPH form a clade separated from centric DPH in phylogenetic studies, suggesting different origins, or an ancient origin in the common diatom ancestor followed by extensive gene loss, but also diversification and gene duplication. DPH was not found in the planktonic pennate genera *Pseudo-nitzschia* and *Fragilariospsis*. This is consistent with the quasi absence of pennate DPH in *Tara* oceans data, while pennate species are abundant in this dataset. Interestingly, the only 2 pennate genes found in the *Tara* oceans data are close to the two DPH copies of *Navicula sp*, (each *Tara* DPH is close to one copy of *Navicula* DPH, suggesting that these are the 2 copies from the same diatom species) and are found only in a few arctic stations. Some *Navicula* are known to thrive in sea ice, it could thus be considered that the DPH found in the *Tara* Oceans are from a sea-ice species released into the water column. The Dark Edge sampling cruise in October 2021 (coordinated by Prof. M. Babin), during which we sampled sea-ice and surrounding waters in the Artic, might bring answer to this question.

Among pennate species with DPH there are well-known benthic species, such as the *Seminavis*, *Gyrosigma*, *Nitzschia*, *Navicula*, and *Amphora* genera. The araphid pennates *Fragilaria radians*, *Synedropsis recta* and *Synedra sp* also possessed DPH, but we did not investigate their environmental niche yet. We also noticed that the benthic species often had several DPH copies (4 for *A. coffeaeformis* and *S. robsuta*, 2 for *Nitzschia* and *Navicula* species). *A. coffeaeformis* was reported before to have 2 putative chromophore-binding cysteines, one in the PAS and one in the GAF domain. However, we saw in chapter 1 that this seems to be a unique feature found in this species.

In the Ocean regions, the distribution of DPH showed a very strong latitudinal gradient, with absence in the tropics and presence in temperate and polar region of both Northern and Southern Hemispheres. In Tropical regions, diatoms, including Thalassiosirales, are found but no DPH could be detected.

It would be interesting to know if this is a feature specific for diatom phytochrome or if other marine organisms with phytochromes show the same trends. For example, the
cyanobacteria *Prochlorococcus* are abundant in Tropical regions and adapted to stratified waters, but do not possess phytochromes, while the *Synechococcus* cyanobacteria, which are more abundant in temperate regions, possess phytochromes. This could suggest that in cyanobacteria too, there is a latitudinal trend of phytochromes, maybe linked to adaptation to mixed waters such as the ones encountered in polar and temperate regions. An impressive set of data (Tara Oceans expedition, Global Ocean Sampling) are available for prokaryotes that could help to address this question. In addition, other Eukaryotic phytochromes found in the *Tara* Oceans data could extend this question to other eukaryotic microalgae, such as Cryptochytes or Glaucophytes.

This strong latitudinal gradient suggests that DPH function is linked to the high seasonal variability in these regions. This was also supported by the statistical analysis of the centric DPH abundance, which is linked to the variation of chlorophyll concentration in temperate regions. The variations of the light field and photoperiod in these regions are important, and in winter and early spring the phytoplankton experiences strong variations of light, from the surface to the bottom of the mixed layer, which can extend below the photic zone. In these conditions, the cells will spend most of their time in low blue light. The *Thalassiosirales* species seem to be early bloomers in the temperate species succession, which could be linked to an efficient adaptation to generally low but fluctuating light thanks to DPH. One important question raised here is the interplay with other photoreceptors. Indeed, we showed that in the marine environment, DPH activity is sensitive to blue, green and red light. Diatoms possess a plethora of blue-light photoreceptors, with 4 aureochromes and several light-sensing cryptochromes (Chapter 1), which makes it surprising that some diatoms have an additional blue-activated photoreceptor. The subtlety of DPH activity is that it is not only sensing the light intensity, but also measuring the ratio between blue and green to red wavebands. Some blue-light photoreceptors such as some aureochromes are thought to be involved in high light adaptation in diatoms, which diatoms experience only close to the surface. Thus, we can imagine reverse gradients of activity for blue light photoreceptors compared to phytochrome in the water column. The first ones would be active at the surface to shallow depth where (blue) light intensity is high, and their activity would decrease when going deeper due to decreasing light intensity. On the contrary, DPH is inactive at the surface,
and its activity increases when going deeper. The integration of DPH signal with blue light sensors signal could bring very precise information on the position in the water column in terms of light quality and intensity. This could also cross-talk with photosynthetic signals from the plastid and together mediate photosynthetic acclimation, such as low light acclimation at depth and photoprotection at surface. DPH KO strains in *Thalassiosira pseudonana* are available, and we could test if DPH mediates acclimation to light mimicking different environmental conditions, such as low blue light or fluctuating light.

We modeled that DPH activity was affected by phytoplankton concentration, such as the ones occurring during a bloom. DPH activity variation could therefore be a signal that a bloom is occurring, and mediate different physiological outputs such as nutrient uptake, resting spore formation or sexual reproduction (discussed in Chapter 2). The model diatom *Thalassiosira pseudonana* is sometimes used to study sexual reproduction, and we could also use another species which possesses DPH, such as *Skeletonema costatum*, to test the induction of sexual reproduction or the formation of resting spores under different light regimes, cell concentrations and nutrient status.

Finally, we showed that DPH activity increases when solar zenith angles are low, such as at dawn and dusk. DPH could thus be a sensor of the end (or beginning) of the day, but we miss critical information on the DPH state in the dark, i.e. its dark reversion rate. *In vitro*, no dark reversion could be observed for the DPH produced here nor in Fortunato et al., 2016. *In vivo*, our reporter system does not allow detection of inactivation, as the eYFP reporter is very stable in the cells. We made the hypothesis that PfrPfr is the dark state (and the form in which DPH is synthetized) and that dark reversion is negligible in the light. However, under photoperiodic conditions (i.e during the night or at sunrise/sunset), we do not know dynamics of the DPH equilibirum. On time scales such as 12h of darkness, dark reversion, synthesis and degradation of DPH are not negligible anymore. We know from tracking of tagged DPH that DPH synthesis is not stable during the day, with degradation of the protein during the light phase and re-accumulation during the night, and a maximum at the dark-light transition. This could point towards a role of DPH at dawn, where very sensitive DPH could detect the first photons and trigger acclimation to the incoming light. However, the tagged DPH might not completely reflect the native protein behaviour, and mRNA profiles have the reverse tendency: the transcription is at