(Fig. 3C), and thus experiencing during the 24 h a total irradiance equivalent to that of a 16L:8D cycle, the fluorescence oscillations showed a rhythmic pattern. Oscillation persisted for at least 4 days in free run, although with a longer period, in accordance with what already reported in Annunziata et al., 2019. Similar oscillation features were observed in cells exposed in L:L to the same light of 25 µmol photons m⁻² s⁻¹, as used for entrainment (Fig. 3D), suggesting that the total increased irradiance due to the absence of the dark period did not affect global growth and synchronization capacity (Table S2 and 4). On the contrary, rhythmicity appeared perturbated in cells experiencing higher light intensity. In particular, in cells pre-adapted in 16L:8D of 75 µmol photons m⁻² s⁻¹ and then exposed to continuous light of 50 µmol photons m⁻² s⁻¹ (Fig. 3E), clear oscillations were visible during the first two days of free-run, with a first peak of fluorescence occurring at L:L14 followed by the second peak at L:L42-44, and estimated period of about 28-30 hour. After the second day, we reported a rapid damping of the fluorescence profiles that made a reliable calculation of the period impossible. In the case of cells exposed to the 75 µmol photons m⁻² s⁻¹ in both L:D and after release to L:L (Fig. 3E), the cultures lost a consistent rhythmic profile following the end of the first subjective day. All together these data support a strong regulation of biological rhythms in diatoms by an endogenous oscillator, whose activity is potentially influenced by other light-intensity dependent metabolic and cellular processes.

We then investigated the effect of the RITMO1 mutation on free running oscillation. Differently from what reported in cells exposed to periodic L:D cycles (Fig. 2), significant effect on cellular rhythmicity was observed in all the transgenic lines, when exposed to conditions in which Wt cells exhibited strong circadian rhythms (Fig. 4). In particular, when cells were adapted to 16L:8D 25 µmol photons m⁻² s⁻¹ and subsequently

exposed to free running light of 17 photons m⁻² s⁻¹ (Fig 4A-E), both KO lines and OE lines showed alteration of rhythmicity in free running, if compared to Wt and CTR lines. In several independent biological replica (Table S4) these lines were considered arhythmic by using EPR algorithms estimating periodicity. When a rhythm could be calculated (Table S4), both KO1 and KO3 lines and the OE1 exhibited a significant reduction in oscillation amplitude and a trend of increase in RAE when compared to the Wt and CTR (Fig. 4 G and H and Table S4), indicating a strongly affected synchronization of the population in free running condition. In addition, both KO lines manifested a major and significant reduction in period when compared to Wt and CTR lines. Parallel time resolved analysis of cell growth in L:L (Fig. S4) confirmed an alteration of circadian period in KO3 compared to Wt. OE1 (Fig. 4E and Table S4) showed strongly altered oscillations, as for the KOs lines. Analysis of rhythmicity in cells exposed to free running condition of higher light intensity of 25 µmol photons m⁻² s⁻¹ (Fig. S3), showed similar alteration of rhythmicity in *RITMO1* mutants compared to wild-type cells to that observed at lower irradiance (Fig 4A-E). While the increased light intensity did not have any effect on the period of the Wt and CTR lines, a small, but still significant modification was observed in the free running period of KO and OE lines (Fig. S3).

We further inquired whether obliteration of RITMO1 in KO lines or alteration of its expression in OE might have an effect on the capacity of *P. tricornutum* to dynamically respond to environmental inputs. To do so, we first desynchronized the cells by growing them to continuous light of 17 µmol photons m⁻² s⁻¹ for two weeks and we re-exposed them to 16L:8D photoperiod of 25 µmol photons m⁻² s⁻¹ (Fig. 5), in a mirror treatment to that reported in Fig 4. This analysis indicated that all the lines are capable of recovering rhythmic changes in
<figure><img src="image_54.png" /></figure>

<figure><img src="image_55.png" /></figure>

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

<figure><img src="image_57.png" /></figure>

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

<figure><img src="image_59.png" /><figcaption>F</figcaption></figure>

<figure><img src="image_60.png" /><figcaption>G</figcaption></figure>

<figure><img src="image_61.png" /><figcaption>H</figcaption></figure>

<figure><img src="image_62.png" /><figcaption>I</figcaption></figure>

<figure><img src="image_63.png" /><figcaption>J</figcaption></figure>

<figure><img src="image_64.png" /></figure>
<caption>Figure 4. Effects of RITMO1 mutation on cellular fluorescence in free running.</caption>

(A-E) Cellular fluorescence profiles after transition from 16L:8D 25µmol photons m⁻² s⁻¹ to L:L 17µmol photons m⁻² s⁻¹ in Wt, CTR, KO1, KO3 and OE1 cells (n≥8). Dots represent mean B3-A values, coloured ribbons represent SD. White and grey regions represent light and dark periods, grey dashed regions represent subjective nights in free-run conditions. (F-J) Period (F), amplitude (G) and R.A.E. (H) estimations of B3-A fluorescence traces represented in panels A-E that passed the EPR rhythmicity test (mean ±s.e.m., *P<0.05, **P <0.01, ***P < 0.001, One-way analysis of variance against Wt with Bonferroni post-test correction). (I) Plot of period versus R.A.E. estimates. (J) Plot of amplitude versus R.A.E. estimates.

fluorescence, visible at the population level already after the first night, with a period that is attested to 24 hours. The fact that the amplitude of these oscillations is comparable between the first light-dark cycle and the two subsequent ones analysed in the course of the experiment suggests that a treatment of 8 hours of darkness is sufficient to ensure the re-synchronization of the population. Both *RITMO1* KOs and the OE lines did not show differences compared to the Wt and CTR, hinting that the alterations of RITMO1 activity does not affect the capacity of cell cycle to re-synchronise to diel environmental cues.

RITMO1 affects circadian
fluorescence rhythmicity, without
affecting growth and cell cycle
progression

The alteration of cellular fluorescence
rhythmicity observed in free running
conditions in the *RITMO1* KO mutants
compared to Wt and CTR lines opens the
question of whether this effect is due to a
possible role of RITMO1 in the regulation of
cell growth and cell cycle progression. We
therefore monitored the growth of Wt, CTR,
OE1 and KO lines under L:D cycles and
following L:D to L:L shift. The analysis in Fig.
6A and Table S4 indicated that mutants lines
had a growth capacity comparable to that of

<figure><img src="image_65.png" /><figcaption>Figure 5. Re-entrainment of Wt, KO1, KO3 and OE1 cultures adapted to continuous light (L:L) to L:D cycles.</figcaption></figure>

Cellular fluorescence profiles after transition from L:L 17µmol photons m⁻² s⁻¹ to 16L:8D 25µmol photons m⁻² s⁻¹ in Wt and CTR lines (A) and in KO1, KO3 and OE1 lines (B). Cells were pre-adapted to L:L for two weeks before re-entrainment to photoperiod. Dots represent mean B3-A values, coloured dashed lines represent SD (n=3). White and grey regions represent light and dark periods.
the Wt and CTR lines in both 16L:8D and following a shift to L:L (17 µmol photons m⁻² s⁻¹). Moreover, we also tested effect on cell cycle progression by analysing the cell cycle phases of cells first synchronized with a dark treatment of 40 hours and subsequently exposed to continuous light, as previously done (Annunziata et al., 2019; Huysman et al., 2010). As shown in Fig. 6B, the 2 KO lines showed the same percentage of cells in G1 (around 80%) at the end of the dark treatment. A similar reduction of the G1 phases, corresponding to an increase of dividing cells (S/G2), was also observed following light exposure inducing cell cycle progression. The data indicate that the alterations of cell fluorescence rhythmicity observed in *P. tricornutum* in free-run in *RITMO1* KO and

OE1 are attributable to a loss of synchronization of the population associated with an alteration of a clock component, and not to a deregulation of the cell cycle.

RITMO1 is implicated in the circadian regulation of photosynthesis in *P. tricornutum*

Adaptation of the photosynthetic apparatus to external stimuli is vital for photoautotrophic organisms and evidences in plants and algal species, including diatoms, indicate a regulation of this process by an endogenous oscillator (Brand, 1982; Dodd et al., 2014). To evaluate the involvement of RITMO1 in the rhythmic activity of the plastid, we compared the photosynthetic capacity of the Wt, KO1, KO3, and OE1 lines both in cells adapted to 16L:8D cells and following the 16L:8D to L:L transition (Fig. 7A). Samples were collected every 4 hours over 36 hours. In free running, analysis started at the second subjective day, from LL32. Different photosynthetic parameters were measured by performing a Rapid Light response Curve (RLC) (Materials and Methods, Ralph and Gademann, 2005). In 16L:8D, the four lines showed a twofold variation of the Maximal relative Electron Transfer Rate (ETRmax) (Fig. 7B) and photoprotection capacity here measured via the NPQ (Fig. 7D) over the 24 hours. Both parameters started to increase during the dark, peaked in the middle of the light and reached a minimum towards the final hours of the light period. Notably, ETRmax and NPQ peak during the light period, while the maximal

<caption>Figure 6. Analysis of the effect of RITMO1 obliteration and overexpression on growth capacity and cell cycle progression.</caption>

(A) Growth curves of Wt, CTR, KO1, KO3 and OE1 cells entrained under 16L:8D 25µmol photons m⁻² s⁻¹ and released to L:L 17µmol photons m⁻² s⁻¹. Dots and error bars represent the mean ±s.e.m. Log2 of cell density expressed as 10⁶ cells mL⁻¹ (n=3). (B) Percentage of cells in G1 phase in cultures synchronized for 48 hours in dark and subsequently exposed to 25 µmol photons m⁻² s⁻¹ continuous light. Cell cycle was determined by flow cytometry after DAPI staining of the cellular DNA content. Columns represent mean percentage values of cells in G1 at each time point ± s.e.m. (n=3). Two-way analysis of variance with Bonferroni correction was used to compare cell cycle progression among lines, but no significant difference was detected.

<figure><img src="image_66.png" /></figure>
<figure><img src="image_67.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_68.png" /><figcaption>C</figcaption></figure>

<figure><img src="image_69.png" /><figcaption>D</figcaption></figure>

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

<figure><img src="image_71.png" /><figcaption>F</figcaption></figure>

<figure><img src="image_72.png" /><figcaption>G</figcaption></figure>

<figure><img src="image_73.png" /></figure>

Figure 7. Analysis of the effect of RITMO1 mutation and overexpression on different photosynthetic parameters under periodic L:D cycles and free running conditions.

A) Schematic representation of sampling points for the analysis from Wt, KO1, KO3 and OE1 cells grown under 16L:8D 25µmol photons m⁻² s⁻¹ and released to L:L 17µmol photons m⁻² s⁻¹. Samples were collected every 4 hours over 36 hours in L:D and L:L. In free running, samples were collected starting at L:L32. (B) Maximal relative Electron Transfer Rate (rETRmax), (D) non photochemical quencing (NPQ) and (F) photosynthetic rate under light limitation (α) in L:D. (C) rETRmax, (D) NPQ and (G) α in cells transferred to L:L.. Dots represent mean values ± s.e.m. (n=3).
value is attained between ZT4 and ZT8 in both wild-type and the two KO lines. The light-limited initial slope (α), which is an indication of the maximum quantum yield of PSII, manifested oscillations of minor magnitude (25% of its maximum value over the course of the day), with a minimum after 4-8 hours after the beginning of the light period, followed by a progressive increase until the end of the dark phase (Fig. 7F). In L:L, the Wt lines retained an oscillation with a period of 24 and 28 hours for both ETRmax and NPQ respectively, although the rhythms were attenuated compared with the 16L:8D (showing approximately a 30% variation relative to the maximum value) (Fig. 7C and E). As for the cell fluorescence, a clear shift in the phase was observed compared to the L:D, with a peak around the transition from the subjective light to the subjective dark phase. The KO mutants and the OE1 line did not show significant oscillations already after the second day in free run condition, supporting an involvement of RITMO1 in the circadian regulation of these processes. Considering the α value, no major variations during the 36 hours analysed can be observed for any of the four lines considered (Fig. 7G).

RITMO1 plays a central role in the
circadian control of gene expression
in diatoms

RITMO1 belongs to the bHLH-PAS
family of the *P. tricornutum* transcription
factors. The deregulation of its expression by
ectopic overexpression generates a
perturbation of gene expression rhythmicity in
L:D but also in continuous dark (Annunziata
et al., 2019), indicating that its activity is
independent on direct light inputs. Here we
further investigated the role of RITMO1 in
the circadian clock system, by analysing the
expression of a set of genes of interest by RT-
qPCR in cells adapted to 16L:8D (25 µmol
photons m⁻² s⁻¹) and following a 16L:8D to L:L
(17 µmol photons m⁻² s⁻¹). In order to uncover

rhythms controlled by the circadian clock
only, persistence in gene expression oscillation
in free-run condition was analysed every 4
hours between LL32 to LL68, at the second
and third subjective night and day, as done for
the photosynthesis experiment (Fig. 7A). Wt,
KO3 and OE1 lines were chosen for the
analysis in order to compare the effect of both
obliteration and overexpression of *RITMO1*
on expression patterns. First, the expression
level of *RITMO1* was analysed to check how
the deregulation of the protein affects its own
transcription. Because we also analysed
expression in the ectopic OE line, we used two
different primer sets recognized the
endogenous *RITMO1* mRNAs (Fig. 8A left)
and also the total transcript (endogenous plus
transgene) in OE lines (Fig. 8A right). As
shown in Fig. 8A (left), KO3 and OE1 cells
showed rhythmic profile of the endogenous
*RITMO1* expression in L:D comparable to
that of Wt. OE1 showed a ~60 % lower
expression level at the peak of expression
(ZT12), as already shown in (Annunziata et al.,
2019). Although less pronounced, a reduction
in gene expression was observed in KO3 too,
indicating that *RITMO1* transcription was
maintained despite the mutation of the gene
by CRISPR/Cas9 and total obliteration of the
full-length protein (Fig. 1). In Wt cells in free-
run, *RITMO1* transcript showed a clear
oscillation with an increase in transcript levels
before the start of the second subjective night,
followed by a decline in the following hours, a
minimum at LL52 and a new increase in the
expression in the following hours. The peak of
expression is delayed from the subjective L:D
day by approximately four hours. As for the
OE1 and KO lines, they both lost the
oscillation observed in the Wt, with a flattened
expression profile and an average transcript
value in the range of that of the control line.
By using a pair of primers designed to
recognise the total *RITMO1* mRNA in OE1
(Fig 8A right), in L:D a rhythmic pattern was
still observed in OE1, but with an earlier peak
of expression compared to Wt and KO lines,
due to the over-expression of *RITMO1* driven
by the FcpB promoter. In L:L, OE1 showed higher expression level than Wt and distinct rhythmic pattern, with an acrophase during the subjective light period. As expected, both the Wt and KO1 showed no difference in the expression pattern compared to the analysis done with the other set of primers. In addition to *RITMO1*, we also analysed other TFs transcripts (*bHLH1b*, *bHLH3* and *bZIP7*) exhibiting strong rhythmic expression in L:D cycles (Annunziata et al., 2019). All these TFs showed persistence of gene expression rhythmicity in L:L in Wt cells (Fig. 8B), which was strongly altered in both OE1 and KO3 for the *bHLH1b* and *bZIP7*. For the *bHLH3*, a minimal residual rhythmicity was still detected in KO3, although with an altered phase. In addition, we also analysed the expression of photoreceptors, potential actors in the input pathway of the diatom circadian clock (Fig. 8C). The analysis of the *aureochrome1* (*Aureo1a*) was particularly relevant. This blue light receptor also acts as light-dependent TFs controlling the expression of an important number of genes in *P. tricornutum*, including *RITMO1* (Mann et al., 2020). In our study, *Aureo1a* expression appeared rhythmic under diurnal (L:D) and free run (L:L) conditions in Wt, and this rhythmic pattern was strongly altered by *RITMO1* mutation in KO3 or OE1 in L:L. Rhythmic expression was also observed for the Red/Far-red light phytochrome photoreceptor (*DPH*) in L:D. Similarly, DPH showed oscillating expression at the second and third day in L:L in Wt. Minor residual oscillations were observed in OE1 and KO3 only at the second subjective day, which appeared anticipated compared to the Wt. Rhythmic patterns in L:D and at the second and third L:L were detected for the blue-light receptor CPF1 too. KO3 and OE1 showed no alteration in L:D compared to Wt, but greatly reduced oscillation in amplitude and temporally offset by 4-8 h when compared with Wt in L:L.

We also analysed the expression of genes involved in the control of the cell cycle

(Huysman et al., 2010, Fig. 8 D). The *CYCP6* gene, whose expression has been shown to be associated to G1 phase in synchronized culture, showed a rhythmic expression in L:D, with a pick at the end of the night period. In the OE1 and KO3, we detected some effect in the expression levels but not in the rhythmic profile in L:D compared to the Wt. Minor rhythmicity was detected in Wt in the second free running day, further reduced at the third day. Rhythmic patterns were not observed in the transgenic lines for *CYCP6*. The *CycB1* gene, considered as a marker of the G2/M phase, showed a clear rhythmic pattern in L:D, peaking at the end of the light period, which was overall unaltered in KO3 and OE1. However, the expression of *CycB1* became arrhythmic in L:L in all the lines. Finally, we also monitored expression of *dsCYC2*, a key regulator of the cell cycle onset following dark to light transition. It has been previously reported that the strong blue-light dependent regulation of *dsCYC2* is directly controlled by *Aureo1* (Huysman et al., 2013). Our study also measured a rhythmic expression of *dsCYC2* in the Wt, with a clear anticipation of its expression before the light onset (Fig. 8 D). This pattern was also confirmed by reanalysing public diurnal transcriptomic data (Chauton et al., 2013). The strong light induction, which is mediated by *Aureo1a* just after the light onset, is not detected in our analysis because of the selected sampling points. Remarkably, *dsCYC2* maintains robust oscillation also in Wt in free running. Both, the OE1 and KO3 showed a strongly reduced expression of this gene at the end of the dark period, and loss of rhythmicity in L:L, supporting an implication of *RITMO1* in both the light anticipation of *dsCYC2* expression in L:D and circadian rhythmicity in L:L conditions. Finally, we also analyzed the expression of two genes linked to chloroplast biogenesis and activity: one of the *P. tricornutum* *FtsZ* genes, putatively encoding for a key component of the chloroplast division machinery (Gillard et al., 2008; TerBush et al., 2013) and the protochlorophyllide oxidoreductase 2, *POR2*.
involved in chlorophyll biosynthesis (Hunsperger et al., 2016). For *POR2*, we observed a strong rhythmicity of the expression in both L:D and L:L, which is affected by KO deregulation in free run. On the contrary, for *FtsZ* we observed rhythmic

oscillation in L:D conditions, but not in LL, and the expression pattern is not altered in RITMO OE or KO lines, ruling out the possible regulation of *FtsZ* by an endogenous oscillator.

Figure 8. Analysis of the expression of selected genes by qRT-PCR in Wt, OE1 and KO3 lines under periodic L:D cycles and free running conditions.

For the analysis the samples were collected every 4 hours over 36 hours in cells adapted to 16L:8D cycles 25µmol photons m⁻² s⁻¹ and released to L:L 25µmol photons m⁻² s⁻¹. In free running, samples were collected starting at LL32. Dots represents separate replicates cultures grown in independent experiments (n = 2, bold line represent the average value of gene expression per each line). (A) RITMO1 expression profiles. RITMO1-endo refers to endogenous transcripts, while RITMO1-total includes endogenous RITMO1 and RITMO-HA transgene transcripts for the OE1 line; (B) Transcription factors bHLH1b, bHLH3 and bZIP7; (C) Photoreceptors aureochrome 1a (AUREO1a), cryptochrome photolyase family (CPF1) and diatom phytochrome (DPH); (D) Expression profiles of the cyclins CYCP6, CYCB1 and dsCYC2 and (E) of the FtsZ and protochlorophyllide oxidoreductases 2 (POR2). For each line and each time point, the expression value of the genes is normalized over the geometrical average of the TBP and RPS control genes expression. Over the time course, the expression of each gene is normalised to its maximum expression over the three lines, where ‘1’ represents the highest expression value of the time series.
<figure><img src="image_74.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_75.png" /><figcaption>C</figcaption></figure>

<figure><img src="image_76.png" /><figcaption>D</figcaption></figure>

<figure><img src="image_77.png" /><figcaption>E</figcaption></figure>

<figure><img src="image_78.png" /></figure>
Discussion

This study provides evidences for a
strong temporal regulation of biological
processes by a circadian clock system in an
important group of marine phytoplankton,
diatoms, which are at the crossroads of several
evolutionary lineages (Keeling, 2013).

Our investigation of cell fluorescence
rhythmicity (Fig. 2 and Fig.3) supports the
existence of a self-sustained oscillator in
diatoms able to synchronize processes
associated to cell division. The increase in *P. tricornutum* cell number after the fluorescence
peak, begins to be observed around the
transition to darkness and parallels a decrease
in cell fluorescence caused by plastid splitting
between daughter cells (Huysman et al., 2010).
The adjustment of cell fluorescence phase
according to day length as well as the
anticipation of darkness suggest day length is a
main driver of cell division timing. Under short
days (8L:16D), cell fluorescence in *P. tricornutum* cultures still increases before the
end of the light period, but the acrophase is
reached in the dark, suggesting that a
timekeeper restricts cell division to a window
of time, more than to a specific time, as already
observed in *Ostreococcus* tauri (Moulager et
al., 2007). This short photoperiod effect is
observed in cells showing a lower division rate,
compared to that adapted to longer
photoperiods, but also in cells showing a
higher division rate in 8L:16D, when exposed
to higher irradiance. In most organisms, cell
division cycle and the circadian cycle co-exist
and are somehow intertwined (Masri et al.,
2013). We can therefore assume that under
short photoperiods, cell separation is delayed
to after the dark onset, but the activity of the
circadian clock is still visible because cells
maintain a 24-hour period. The circadian
regulation of processes monitored by cell
fluorescence is further supported by the
persistence of rhythmicity under free running
conditions, as previously described

(Annunziata et al., 2019). We here found that
this effect is observable as long as the light
intensity in free running is not limiting for the
growth and, potentially, for other metabolic
activities necessary to sustain cellular activities
in a broad sense. During free run, *P. tricornutum* rhythms of cell fluorescence shift
into a new phase and have periods longer than
24 hours at the light conditions tested,
indicating that the circadian clock adjusts to its
own time in the absence of periodic
environmental signals.

In *P. tricornutum*, photosynthesis also
appears regulated by a circadian clock (Fig. 7),
supporting previous observation (Palmer et al.,
1964) and also *in situ* studies reporting strong
oscillations in optical properties related to
photosynthetic activity and growth in natural
populations along the seasons (Kheireddine
and Antoine, 2014). In our study, rhythms of
photosynthesis are clearly visible in light-dark
and persist at least until the third day under
free running conditions, although with smaller
oscillations. On the other hand, we also
observed that cell fluorescence oscillations,
proxy of cell division and pigment synthesis, in
free running conditions are strongly affected or
even lost when cells experience higher light
intensities (Fig. 3). Possibly, changes in the
photosynthetic activity due to the absence of
dark and/or higher light intensity may affect
circadian clock system controlling cell
fluorescence rhythmicity. Increasing evidences
in multicellular plants and animals indicate that
metabolic processes not only are regulated by
the circadian clock, but also in turn regulate its
function (Asher and Sassone-Corsi, 2015;
Farré and Weise, 2012; Haydon et al., 2017;
Mora-García et al., 2017), by acting in the input
pathways of the clock system. Our study
suggests that a regulatory cross-talk between
photosynthesis and an endogenous
timekeeper likely also exists in unicellular algae
such as diatoms. This cross-talk deserves to be
studied in more detail as this may be
particularly relevant for enabling rapid
acclimatisation responses of diatoms under
changing environmental condition and for
regulating their growth capacity. Specifically, it
will be relevant to attest the persistence of other
clock-regulated processes, notably
photosynthesis and gene expression, under
increasing free-run light intensity. Such an
analysis would allow to determine whether the
masking of cellular fluorescence persistence
under high light is peculiar of this phenotype
or a generalised effect on clock activity. This
complex network of regulation may indeed
mask circadian clock activity under particular
conditions, and could explain why the step
wise increase of cell density under continuous
light, proposed as a strong indication of cell
cycle gating by a circadian clock, has been
detected in some (Ostgaard and Jensen, 1982),
but not all diatom species (Chisholm and
Brand, 1981). A weak free-running rhythmicity
of cell cycle comparable to what is shown in
Fig. S4 has also been reported in
*Nannochloropsis*, another *Stramenopila*, like
diatoms (Poliner et al., 2019).

Our work clearly defines RITMO1 as
a component of the diatom circadian clock
system. Thanks to the generation of knock-out
mutants in which this protein has been
genetically eliminated (Fig. 1), we observed
that three circadian processes described in this
study are directly affected by RITMO1
mutation (Fig. 4, Fig. 7 and Fig. 8). The fact
that mutants in the *RITMO1* gene are not
affected in growth capacity after L:D to L:L
transitions nor in the progression of cell cycle
after dark synchronisation (Fig. 6), allows us to
exclude that the altered rhythmicity of cell
fluorescence is due to a direct implication of
this protein in the regulation of the cell cycle
machinery. Moreover, the fact that the period
of cell fluorescence is independent of the rate
of division in wild-type cells, but it is altered in
the *RITMO1* KO lines, supports the idea that
the circadian clock is a different process from
the cell division cycle in diatoms. *RITMO1*-
like bHLH-PAS genes have been found in all

diatoms and other *Stramenopila* for which
omic sequences are available (Annunziata et
al., 2019; Bilcke et al., 2021; Farré, 2020),
suggesting a possible conservation of its role
across this clade. Of note, an expansion of
bHLH-PAS TFs showing different rhythmic
expression times has been recently described
in the benthic diatom *Seminavis robusta*, likely
reflecting an adaptations and functional
diversification of this protein family to both
diurnal and tidal cycles (Bilcke et al., 2021).

The transcription analysis performed
in this study support a function of RITMO1 in
the diatom central circadian transcriptional
oscillator (Fig. 8). First of all, robust circadian
oscillations in gene expression are detected for
most, but not all, of the rhythmic genes
analysed at the second and third free running
day, supporting a circadian regulation of
diatom transcription, as seen in most
eukaryotes. The phases of these rhythms are
shifted compared to the L:D, as observed for
the cell fluorescence and photosynthesis and
as photosynthesis they show an apparent
period of 24h, indicating that the
transcriptional rhythms are less affected by the
physiological changes of the cells following the
L:L transitions and maintain an apparent
period close to the one of L:D entrainment.
As for other physiological rhythms
characterized in this study, the effect of
RITMO1 obliteration on rhythmic gene
expression is visible in free running conditions
but it's masked under diel light-dark cycles. An
equally likely alternative is that the rhythmic
light cues used in the study are sufficient to
ensure complete masking of endogenous
clock effects and rhythmic physiology is just a
direct response to environmental stimuli. This
suggests that other, still unidentified clock
components, may perform a redundant
function with respect to RITMO1,
compensating for its mutation as long as there
are periodic light cues. The redundancy of
RITMO1 function could also explain some
residual cell fluorescence oscillation in KO
mutants, although with shorter periods and
reduced amplitudes, observed in some replicates. A parallel can be made with the central clock component CCA1 and LHY of *Arabidopsis thaliana*, in which a single mutation results in a shortening of the circadian period in free-run, whereas a double mutation is required to induce arrhythmicity (Alabadi et al., 2002; Mizoguchi et al., 2002).

By identifying RITMO1 as a transcriptional regulator of the diatom circadian clock, this study paves the way for the molecular identification of other RITMO1-interacting components in the as yet unresolved TTFL system. Between the transcription factor analysed, other members of the bHLH family (bHLH1b and bHLH3), but also bZIP7, represent direct or indirect RITMO1-target genes and may participate in circadian clock function.

The discovery of circadian regulation of the diatom Red/Far-Red light Phytochrome DPH photoreceptor, the blue light Cryptochrome Photolyase 1 CPF1 and aureochrome 1a (Aureo1), make these proteins interesting new candidate regulators of the light-dependent rhythmic processes and possibly of the clock input pathway of diatoms. Interestingly, the absence of RITMO1 or deregulation of its expression by ectopic overexpression does not affect the ability of cells to readapt to periodic light conditions after acclimation to constant light for several weeks (Fig. 5). This suggests that RITMO1 is not directly implicated in the input pathways needed for the clock entrainment, but rather in the core clock system. Still, an effect on the free running period is observed in the mutant KO3 for RITMO1 exposed to higher light intensity in continuous light (Fig. 4 and Fig. S3), which shows a longer period length than under L:L 17 µmol photons m⁻²s⁻¹. Further investigations under different fluence rate and quality of light would be necessary to further assess the primary role of RITMO1 in the circadian clock system and its possible

interaction with the photic control of circadian rhythms.

Regulation by RITMO1 is also observed for the protochlorophyllide oxidoreductase 2 (POR2), putatively involved in chlorophyll biosynthesis (Hunsperger et al., 2016), further supporting a circadian regulation of processes of relevance for plastid function. Between the analysed genes involved in cell cycle progression or plastid division machinery (Gillard et al., 2008; Huysman et al., 2010, 2013), a clear circadian regulation has been seen only for *dsCYC2*. This gene has been described as a critical regulator of cell cycle progression of diatoms because its down-regulation clearly alter the onset of cell division (Huysman et al., 2013). *dsCYC2* has been characterized as a blue light regulated gene. Its expression is strongly and transiently induced after switching from darkness to light, thanks to the activity of the blue light-receptor Aureo1a, which also act as transcription factor on *dsCYC2* promoter. Our analysis here identifies another important level of regulation of this critical process by the circadian clock component RITMO1. This is clearly visible from the anticipation of *dsCYC2* expression before the light onset in L:D cycles, the circadian rhythmic expression in L:L and the loss of *dsCYC2* and *Aureo1a* rhythmicity in RITMO1 mutants. Interestingly, KO and OE lines also show a deregulated and lower expression of *dsCYC2* in L:D at the transition from light to dark (Fig. 8). We still do not know if this effect corresponds to a real reduction in the maximal expression of *dsCYC2* in mutants compared to Wt or only into a shift in its peak of expression, which is not detectable in our analyses because of the selected time points.

Finally, the initial generation of clock mutants also offers new opportunities to assess the importance of the circadian clock for diatom life. Based on the initial characterisation, the RITMO1 mutation does not appear to affect *P. tricornutum* growth.
However, further research with multiple clock
mutants will be needed to clearly assess the
role of the diatom clock on fitness. In
cyanobacteria, the effect of clock mutation on
fitness is only clearly seen in competition
experiments between circadian period
mutants and wild-type cells and when their
internal rhythms match those of the
environmental cycle (Ouyang et al., 1998;
Woelfle et al., 2004). Similar observations are

also made with a variety of clock mutants in
plants (Dodd et al., 2005; Yerushalmi et al.,
2011). Further analysis of the responses to
widely varying photoperiodic conditions will
also be of interest, as diatoms are the
dominant photosynthetic organisms in polar
regions and the circadian clock is central to
photoperiodic regulation in terrestrial plants
and animals.

Acknowledgements

This work has been supported by funding from the Fondation Bettencourt-Schueller (Coups d'élan pour la recherche francoise-2018), the "Initiative d'Excellence" program (Grant "DYNAMO," ANR-11-LABX-0011-01), the ANR-DFG Grant "DiaRhythm (KR1661/20-1) to A.F. and ANR Grant ClimaClock (ANR-20-CE20-0024) to A.F and F-Y.B.
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B

<figure><img src="image_79.png" /><figcaption>Figure S1. Sequence analysis of the KO mutants in the *RITMO1* gene.</figcaption></figure>

(A) Alignment of the sequence of the RITMO1 in Pt1 Wt strain (allele 1) with that of the *RITMO1* KOs 1-4; (B) Alignment of the sequence of the RITMO1 in Pt1 Wt strain (allele 2) with that of the *RITMO1* KOs 1-4. The alignments were generated via EBI Clustal Omega webtool and visualized via Mview. Bases are colored by identity to the Wt reference sequence. gHLH1, gHLH2 and gHLH3 gRNA target sites are highlighted in red, orange and yellow respectively
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<figure><img src="image_83.png" /><figcaption>Figure S2. Characterisation of growth profile in *P. tricornutum* grown under 16L:8D cycles.</figcaption></figure>

Characterisation of growth (A and B), cellular fluorescence (C) and cell cycle synchronization (D) in Wt cells entrained under 16L:8D 25µmol photons m⁻² s⁻¹. Samples were collected every three hours for the analysis. (A) Cell concentration plotted as Log2 of cell density expressed as 10⁶ cells mL⁻¹. (B) Residuals after baseline detrending of the data shown in A, to extract oscillatory components. (C) Cellular fluorescence (B3-A parameter) profile of the samples reported in panel (A) and (B). Solid lines represent the average, coloured ribbons represent the SD (n=3). (D) Percentage of cells in G1 phase in the highlighted time window. Cell cycle was determined by flow cytometry after DAPI staining of the cellular DNA content. Columns represent mean ± s.e.m. (n=3).
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<figure><img src="image_93.png" /><figcaption>Figure S3. Effects of RITMO1 mutation on cellular fluorescence in L:L 25 µmol photons m⁻² s⁻¹.</figcaption></figure>

(A-E) Cellular fluorescence profiles after transition from 16L:8D 25µmol photons m⁻² s⁻¹ to L:L 25µmol photons m⁻² s⁻¹ in Wt, CTR, KO1, KO3 and OE1 cells (n≥6). Dots represent mean B3-A values, coloured ribbons represent SD. Gray dashed regions represent subjective nights in free-run conditions. (F-H) Period (F), amplitude (G) and R.A.E. (H) estimations of B3-A fluorescence traces represented in panels A-E that passed the EPR rhythmicity test (mean ±s.e.m., *P<0.05, **P <0.01, ***P < 0.001, t test). (I) Plot of period versus R.A.E. estimates. (J) Plot of amplitude versus R.A.E. estimates.
<figure><img src="image_94.png" /></figure>

<figure><img src="image_95.png" /></figure>

<figure><img src="image_96.png" /></figure>

<figure><img src="image_97.png" /><figcaption>Figure S4. Time-resolved analysis of growth in free-run cultures.</figcaption></figure>

Characterisation of growth and cellular fluorescence in Wt and KO3 cells entrained under 16L:8D 25µmol photons m⁻² s⁻¹ and released to L:L 17µmol photons m⁻² s⁻¹. Samples were collected every three hours starting from LL0. (A) Concentration of Wt cultures plotted as Log2 of cell density expressed as 10⁶ cells mL⁻¹. (B) Residuals of the curve in panel (A) after baseline detrending. Panels (C, D) report the same parameters for KO3 cultures. Solid lines represent the average, coloured ribbons represent the SD (n=3). All replicates (both for Wt and KO3) are rhythmic according to the EPR test.
<table><caption>Table S1. List of the sgRNAs used for CRISPR-Cas9 mutagenesis of the *RITMO1*.</caption><thead><tr><th>Gene</th><th>Accession</th><th>Name</th><th>Strand</th><th>gRNA sequence</th><th>3' PAM</th></tr></thead><tbody><tr><td rowspan="2">APT</td><td rowspan="2">Phatr3_J6834</td><td>APTg1</td><td>Rv</td><td>AAGCGTGGAATGCCTTTGAA</td><td>GGG</td></tr><tr><td>APTg3</td><td>Fw</td><td>CCTGGGTCCACCAATTGCCC</td><td>TGG</td></tr><tr><td rowspan="3">RITMO1</td><td rowspan="3">Phatr3_J44962</td><td>gHLH1</td><td>Rv</td><td>TCAAGATTCGCGAAAGCAGG</td><td>CGG</td></tr><tr><td>gHLH2</td><td>Fw</td><td>ATGCCGAGTGGGGCTAGAAA</td><td>CGG</td></tr><tr><td>gHLH3</td><td>Fw</td><td>ATACAGTACATTGGTTACGG</td><td>TGG</td></tr></tbody></table>

Target gene, sgRNA name, target strand, gRNA sequence and the associated protospacer adjacent motif (PAM) are reported.

<table><caption>Table S2: Growth capacity of *P. tricornutum* Wt cells in the different conditions used throughout this study.</caption><thead><tr><th>Wt divisions per day</th><th>Mean</th><th>s.e.m.</th></tr></thead><tbody><tr><td>16L:8D (25 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>0.97</td><td>0.02</td></tr><tr><td>16L:8D (75 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>1.49</td><td>0.04</td></tr><tr><td>12L:12D (25 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>1.05</td><td>0.05</td></tr><tr><td>8L:16D (25 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>0.64</td><td>0.03</td></tr><tr><td>8L:16D (50 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>1.02</td><td>0.01</td></tr><tr><td>16L:8D (25 µmol*m<sup>-2</sup>s<sup>-1</sup>) to D:D</td><td>N.A.</td><td>N.A.</td></tr><tr><td>16L:8D (25 µmol*m<sup>-2</sup>s<sup>-1</sup>) to L:L (5 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>0.27</td><td>0.03</td></tr><tr><td>16L:8D (25 µmol*m<sup>-2</sup>s<sup>-1</sup>) to L:L (17 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>1.18</td><td>0.02</td></tr><tr><td>16L:8D (25 µmol*m<sup>-2</sup>s<sup>-1</sup>) to L:L (25 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>1.22</td><td>0.02</td></tr><tr><td>16L:8D (75 µmol*m<sup>-2</sup>s<sup>-1</sup>) to L:L (50 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>1.15</td><td>0.03</td></tr><tr><td>16L:8D (75 µmol*m<sup>-2</sup>s<sup>-1</sup>) to L:L (75 µmol*m<sup>-2</sup>s<sup>-1</sup>)</td><td>1.4</td><td>0.02</td></tr></tbody></table>

First column indicates the irradiation and photoperiodic conditions considered in the analyses. For each condition, the cell division/day was calculated in cells in exponential phase over the course of four days (n=3). In the circadian experiments performed in cells preadapted to L:D cycles and then exposed either to darkness or light of different intensities, the data refers to the growth analysis in free running conditions, after the L:D to L:L shift.
<table><caption>Table S3. Characterisation of cellular fluorescence rhythmicity in Wt, CTR, KO1, KO3 and OE1 lines in L:D.</caption><thead><tr><th rowspan="2">LD</th><th colspan="2">Wt</th><th colspan="3">CTR</th><th colspan="3">KO1</th><th colspan="3">KO3</th><th colspan="3">OE1</th></tr><tr><th>Mean</th><th>s.e.m</th><th>Mean</th><th>s.e.m</th><th>p value</th><th>Mean</th><th>s.e.m</th><th>p value</th><th>Mean</th><th>s.e.m</th><th>p value</th><th>Mean</th><th>s.e.m</th><th>p value</th></tr></thead><tbody><tr><td colspan="15" style="text-align: left; font-weight: bold;">16L:8D (25 µmol*m-2*s-1)</td></tr><tr><td>n =</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Phase</td><td>-2.27</td><td>0.39</td><td>-1.81</td><td>1.01</td><td>N.S.</td><td>-2.73</td><td>0.87</td><td>N.S.</td><td>-2.53</td><td>1.05</td><td>N.S.</td><td>-2.52</td><td>0.62</td><td>N.S.</td></tr><tr><td>R.A.E.</td><td>0.14</td><td>0.01</td><td>0.15</td><td>0.02</td><td>N.S.</td><td>0.18</td><td>0.01</td><td>N.S.</td><td>0.19</td><td>0.01</td><td>N.S.</td><td>0.19</td><td>0.00</td><td>N.S.</td></tr><tr><td>Amplitude</td><td>1.31</td><td>0.02</td><td>1.26</td><td>0.17</td><td>N.S.</td><td>1.26</td><td>0.10</td><td>N.S.</td><td>1.28</td><td>0.05</td><td>N.S.</td><td>0.94</td><td>0.02</td><td>**</td></tr><tr><td>Period</td><td>24.10</td><td>0.08</td><td>24.27</td><td>0.16</td><td>N.S.</td><td>24.08</td><td>0.06</td><td>N.S.</td><td>24.35</td><td>0.07</td><td>N.S.</td><td>24.27</td><td>0.14</td><td>N.S.</td></tr><tr><td colspan="15" style="text-align: left; font-weight: bold;">16L:8D (75 µmol*m-2*s-1)</td></tr><tr><td>n =</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Phase</td><td>-1.07</td><td>0.54</td><td>-0.99</td><td>0.09</td><td>N.S.</td><td>0.21</td><td>0.50</td><td>N.S.</td><td>-0.45</td><td>0.80</td><td>N.S.</td><td>-2.88</td><td>1.65</td><td>N.S.</td></tr><tr><td>R.A.E.</td><td>0.26</td><td>0.03</td><td>0.31</td><td>0.03</td><td>N.S.</td><td>0.35</td><td>0.02</td><td>N.S.</td><td>0.29</td><td>0.02</td><td>N.S.</td><td>0.33</td><td>0.02</td><td>N.S.</td></tr><tr><td>Amplitude</td><td>0.78</td><td>0.08</td><td>0.68</td><td>0.09</td><td>N.S.</td><td>0.62</td><td>0.02</td><td>N.S.</td><td>0.58</td><td>0.04</td><td>N.S.</td><td>0.81</td><td>0.06</td><td>N.S.</td></tr><tr><td>Period</td><td>23.82</td><td>0.49</td><td>23.40</td><td>0.13</td><td>N.S.</td><td>23.00</td><td>0.24</td><td>N.S.</td><td>23.92</td><td>0.79</td><td>N.S.</td><td>25.54</td><td>0.98</td><td>N.S.</td></tr><tr><td colspan="15" style="text-align: left; font-weight: bold;">12L:12D (25 µmol*m-2s-1)</td></tr><tr><td>n =</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Phase</td><td>-1.35</td><td>0.51</td><td>-0.15</td><td>0.08</td><td>N.S.</td><td>-0.75</td><td>0.51</td><td>N.S.</td><td>-1.45</td><td>0.62</td><td>N.S.</td><td>-1.18</td><td>0.67</td><td>N.S.</td></tr><tr><td>R.A.E.</td><td>0.15</td><td>0.01</td><td>0.12</td><td>0.01</td><td>N.S.</td><td>0.15</td><td>0.03</td><td>N.S.</td><td>0.19</td><td>0.02</td><td>N.S.</td><td>0.14</td><td>0.01</td><td>N.S.</td></tr><tr><td>Amplitude</td><td>1.16</td><td>0.08</td><td>1.08</td><td>0.07</td><td>N.S.</td><td>1.09</td><td>0.03</td><td>N.S.</td><td>1.21</td><td>0.09</td><td>N.S.</td><td>0.93</td><td>0.05</td><td>N.S.</td></tr><tr><td>Period</td><td>24.47</td><td>0.19</td><td>23.84</td><td>0.16</td><td>N.S.</td><td>24.13</td><td>0.16</td><td>N.S.</td><td>25.08</td><td>0.60</td><td>N.S.</td><td>24.47</td><td>0.33</td><td>N.S.</td></tr><tr><td colspan="15" style="text-align: left; font-weight: bold;">8L:16D (25 µmol*m-2*s-1)</td></tr><tr><td>n =</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Phase</td><td>2.47</td><td>0.13</td><td>3.28</td><td>0.48</td><td>N.S.</td><td>2.51</td><td>0.41</td><td>N.S.</td><td>1.68</td><td>0.19</td><td>N.S.</td><td>2.71</td><td>0.38</td><td>N.S.</td></tr><tr><td>R.A.E.</td><td>0.12</td><td>0.01</td><td>0.16</td><td>0.01</td><td>N.S.</td><td>0.17</td><td>0.02</td><td>N.S.</td><td>0.18</td><td>0.02</td><td>N.S.</td><td>0.13</td><td>0.00</td><td>N.S.</td></tr><tr><td>Amplitude</td><td>1.45</td><td>0.07</td><td>1.31</td><td>0.03</td><td>N.S.</td><td>1.28</td><td>0.10</td><td>N.S.</td><td>1.23</td><td>0.05</td><td>N.S.</td><td>1.10</td><td>0.03</td><td>**</td></tr><tr><td>Period</td><td>23.47</td><td>0.05</td><td>23.34</td><td>0.14</td><td>N.S.</td><td>23.60</td><td>0.23</td><td>N.S.</td><td>23.79</td><td>0.37</td><td>N.S.</td><td>23.79</td><td>0.16</td><td>N.S.</td></tr><tr><td colspan="15" style="text-align: left; font-weight: bold;">8L:16D (50 µmol*m-2*s-1)</td></tr><tr><td>n =</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Phase</td><td>2.32</td><td>0.06</td><td>1.59</td><td>0.28</td><td>N.S.</td><td>2.68</td><td>0.17</td><td>N.S.</td><td>2.57</td><td>0.24</td><td>N.S.</td><td>3.27</td><td>0.38</td><td>N.S.</td></tr><tr><td>R.A.E.</td><td>0.19</td><td>0.03</td><td>0.22</td><td>0.01</td><td>N.S.</td><td>0.20</td><td>0.02</td><td>N.S.</td><td>0.24</td><td>0.03</td><td>N.S.</td><td>0.23</td><td>0.03</td><td>N.S.</td></tr><tr><td>Amplitude</td><td>1.40</td><td>0.07</td><td>1.40</td><td>0.09</td><td>N.S.</td><td>1.30</td><td>0.09</td><td>N.S.</td><td>1.23</td><td>0.09</td><td>N.S.</td><td>1.08</td><td>0.06</td><td>*</td></tr><tr><td>Period</td><td>23.75</td><td>0.22</td><td>24.40</td><td>0.24</td><td>N.S.</td><td>23.50</td><td>0.03</td><td>N.S.</td><td>24.22</td><td>0.12</td><td>N.S.</td><td>23.52</td><td>0.07</td><td>N.S.</td></tr></tbody></table>

Calculated phases, amplitudes, relative amplitude errors and periods of the different lines grown in different L:D cycles, as indicated also in Fig. 3, panels (F-H). The fraction of rhythmic replicates/total of replicates is reported for each line in all conditions. P-values obtained via two tailed t-test against Wt are shown. N.S.: non significant, P>0.05.

<table><caption>Table S4. Characterisation of cellular fluorescence rhythmicity in Wt, CTR, KO1, KO3 and OE1 lines in L:L.</caption><thead><tr><th rowspan="2">L:L (17 µmol*m-2s-1)</th><th colspan="2">Wt (10/10)</th><th colspan="3">CTR (8/8)</th><th colspan="3">KO1 (5/8)</th><th colspan="3">KO3 (7/8)</th><th colspan="3">OE1 (3/5)</th></tr><tr><th>Mean</th><th>s.e.m</th><th>Mean</th><th>s.e.m</th><th>p value</th><th>Mean</th><th>s.e.m</th><th>p value</th><th>Mean</th><th>s.e.m</th><th>p value</th><th>Mean</th><th>s.e.m</th><th>p value</th></tr></thead><tbody><tr><td>Period</td><td>28.36</td><td>0.82</td><td>29.11</td><td>0.85</td><td>N.S.</td><td>24.21</td><td>0.91</td><td>*</td><td>21.97</td><td>0.82</td><td>***</td><td>27.03</td><td>1.73</td><td>N.S.</td></tr><tr><td>R.A.E.</td><td>0.31</td><td>0.05</td><td>0.20</td><td>0.03</td><td>N.S.</td><td>0.42</td><td>0.03</td><td>N.S.</td><td>0.46</td><td>0.03</td><td>N.S.</td><td>0.57</td><td>0.12</td><td>*</td></tr><tr><td>Amplitude</td><td>0.62</td><td>0.06</td><td>0.60</td><td>0.05</td><td>N.S.</td><td>0.41</td><td>0.04</td><td>*</td><td>0.34</td><td>0.02</td><td>**</td><td>0.28</td><td>0.04</td><td>**</td></tr><tr><td>Division/Day (n:3)</td><td>1.18</td><td>0.02</td><td>1.26</td><td>0.07</td><td>N.S.</td><td>1.18</td><td>0.03</td><td>N.S.</td><td>1.23</td><td>0.02</td><td>N.S.</td><td>1.23</td><td>0.06</td><td>N.S.</td></tr><tr><td colspan="15" style="text-align: left; font-weight: bold;">L:L (25 µmol*m-2s-1)</td></tr><tr><td>n =</td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>Phase</td><td>28.54</td><td>0.62</td><td>28.99</td><td>0.71</td><td>N.S.</td><td>24.68</td><td>2.40</td><td>N.S.</td><td>27.31</td><td>1.54</td><td>N.S.</td><td>24.86</td><td>0.02</td><td>N.S.</td></tr><tr><td>R.A.E.</td><td>0.42</td><td>0.04</td><td>0.50</td><td>0.03</td><td>N.S.</td><td>0.68</td><td>0.05</td><td>**</td><td>0.48</td><td>0.04</td><td>N.S.</td><td>0.47</td><td>0.04</td><td>N.S.</td></tr><tr><td>Amplitude</td><td>0.47</td><td>0.01</td><td>0.42</td><td>0.04</td><td>N.S.</td><td>0.24</td><td>0.02</td><td>***</td><td>0.34</td><td>0.03</td><td>*</td><td>0.39</td><td>0.02</td><td>N.S.</td></tr><tr><td>Divisions/Day (n:3)</td><td>1.22</td><td>0.02</td><td>1.18</td><td>0.06</td><td>N.S.</td><td>1.16</td><td>0.01</td><td>N.S.</td><td>1.17</td><td>0.03</td><td>N.S.</td><td>1.14</td><td>0.09</td><td>N.S.</td></tr></tbody></table>

Calculated periods, amplitudes and relative amplitude errors of the different lines after transition from 16L:8D 25 µmol photons m⁻² s⁻¹ to L:L 15µmol photons m⁻² s⁻¹ or to L:L 25µmol photons m⁻² s⁻¹. Average and s.e.m. values were calculated on the replicates considered rhythmic according to the EPR test. The fraction of rhythmic replicates/total of replicates is reported for each line in both conditions. Values correspond to what is shown in Fig.3, panels (N-P). P-values obtained via t-test against Wt are shown. N.S.: non significant, P>0.05.
<table><caption>Table S5. List of the oligonucleotides used in this work.</caption><thead><tr><th>Gene</th><th>Accession</th><th>Name</th><th>Strand</th><th>Sequence</th><th>Type</th></tr></thead><tbody><tr><td rowspan="2">TBP</td><td rowspan="2">Phatr3_J10199</td><td>TBP_Fw</td><td>Fw</td><td>ACCGGAGTCAAGAGCACACAC</td><td>RT-qPCR</td></tr><tr><td>TBP_Rv</td><td>Rv</td><td>CGGAATGCGCGTATACCAGT</td><td>RT-qPCR</td></tr><tr><td rowspan="2">RPS</td><td rowspan="2">Phatr3_J10847</td><td>RPS_Fw</td><td>Fw</td><td>CGAAGTCAACCAGGAAACCAA</td><td>RT-qPCR</td></tr><tr><td>RPS_Rv</td><td>Rv</td><td>GTGCAAGAGACCGGACATACC</td><td>RT-qPCR</td></tr><tr><td rowspan="8">RITMO1</td><td rowspan="8">Phatr3_J44962</td><td>RITMO1_total_Fw</td><td>Fw</td><td>ATTCTTGGTCCCACCCGGTA</td><td>RT-qPCR</td></tr><tr><td>RITMO1_total_Rv</td><td>Rv</td><td>ACGCCACATTGAAAAACCGAG</td><td>RT-qPCR</td></tr><tr><td>RITMO1_endogenous_Fw</td><td>Fw</td><td>TTATGTCTTTCGGCGACGGG</td><td>RT-qPCR</td></tr><tr><td>RITMO1_endogenous_Rv</td><td>Rv</td><td>AGCAACGAATGCATGCAAGG</td><td>RT-qPCR</td></tr><tr><td>RITMO1_allele1_seq_Fw</td><td>Fw</td><td>TCACCATCAGCAAGGGAAG</td><td>sequencing</td></tr><tr><td>RITMO1_allele2_seq_Fw</td><td>Fw</td><td>TCCCATCACCATCAGCAAC</td><td>sequencing</td></tr><tr><td>RITMO1_allele1_seq_Rv</td><td>Rv</td><td>GAACCGTAGCGATTGTCTT</td><td>sequencing</td></tr><tr><td>RITMO1_allele2_seq_Rv</td><td>Rv</td><td>GAACCGTAGCGATTTTCTG</td><td>sequencing</td></tr><tr><td rowspan="2">bHLH1b</td><td rowspan="2">Phatr3_J44963</td><td>bHIH1b_Fw</td><td>Fw</td><td>CAAAAGCAGCCAACGACGAA</td><td>RT-qPCR</td></tr><tr><td>bHIH1b_Rv</td><td>Rv</td><td>GATATGAGACCCGACCGCTG</td><td>RT-qPCR</td></tr><tr><td rowspan="2">bHLH3</td><td rowspan="2">Phatr3_J42586</td><td>bHLH3_Fw</td><td>Fw</td><td>CACTCTCATCATGCGGGAAT</td><td>RT-qPCR</td></tr><tr><td>bHLH3_Rv</td><td>Rv</td><td>GCGCGTTGTCTTCCTCTATC</td><td>RT-qPCR</td></tr><tr><td rowspan="2">bZIP7</td><td rowspan="2">Phatr3_J48800</td><td>bZIP7_Fw</td><td>Fw</td><td>CCTTATTGATATTCAAGATTCCAAGG</td><td>RT-qPCR</td></tr><tr><td>bZIP7_Rv</td><td>Rv</td><td>GTTTCGGAACCTGCATAGGA</td><td>RT-qPCR</td></tr><tr><td rowspan="2">Aureochrome 1a</td><td rowspan="2">Phatr3_J49116</td><td>Aureochrome1a_Fw</td><td>Fw</td><td>ATGTCCGAACAGCAAAAGGT</td><td>RT-qPCR</td></tr><tr><td>Aureochrome1a_Rv</td><td>Rv</td><td>CTGGAGGGATTCCAACAAGA</td><td>RT-qPCR</td></tr><tr><td rowspan="2">Cpf1</td><td rowspan="2">Phatr3_J27429</td><td>Cpf1_Fw</td><td>Fw</td><td>CCAATTGTTGACCACAAGTTGG</td><td>RT-qPCR</td></tr><tr><td>Cpf1_Rv</td><td>Rv</td><td>CGATTCTTTGCACTTTCTGTTAG</td><td>RT-qPCR</td></tr><tr><td rowspan="2">CYCP6</td><td rowspan="2">Phatr3_J6231</td><td>CYCP6_Fw</td><td>Fw</td><td>AGGTGCTTGCTGCTGTTC</td><td>RT-qPCR</td></tr><tr><td>CYCP6_Rv</td><td>Rv</td><td>ACGAGGCATACTTGTGAATCC</td><td>RT-qPCR</td></tr><tr><td rowspan="2">dsCYC2</td><td rowspan="2">Phatr3_J34956</td><td>dsCYC2_Fw</td><td>Fw</td><td>CTATCATCGCACTCGTCATCAAC</td><td>RT-qPCR</td></tr><tr><td>dsCYC2_Rv</td><td>Rv</td><td>TGTCCACCAAAGCCTCCAAAC</td><td>RT-qPCR</td></tr><tr><td rowspan="2">CYCB1</td><td rowspan="2">Phatr3_J46095</td><td>CYCB1_Fw</td><td>Fw</td><td>TCCTGGTCCGCTACTTGAAAG</td><td>RT-qPCR</td></tr><tr><td>CYCB1_Rv</td><td>Rv</td><td>GCTGGCTGGGAAGATAACGC</td><td>RT-qPCR</td></tr><tr><td rowspan="2">FtsZ</td><td rowspan="2">Phatr3_J42361</td><td>FtsZ_Fw</td><td>Fw</td><td>GTGATTTTCGGGGCTTTGGT</td><td>RT-qPCR</td></tr><tr><td>FtsZ_Rv</td><td>Rv</td><td>TTCATTTCCTGTTCGCGTCC</td><td>RT-qPCR</td></tr><tr><td rowspan="2">POR2</td><td rowspan="2">Phatr3_J13001</td><td>POR2_Fw</td><td>Fw</td><td>CCTGGTTGCATTGCCGAATC</td><td>RT-qPCR</td></tr><tr><td>POR2_Rv</td><td>Rv</td><td>TCTCCAACGTATCCTCCCGT</td><td>RT-qPCR</td></tr></tbody></table>

Target gene, name, pairing strand, sequence and use of the oligonucleotides used in this work. Sequencing primers for *RITMO1* are allele specific.
Chapter II.d:

bHLH1b as a candidate
partner of RITMO1 in diatom
circadian clock

Contributions:

Alessandro Manzotti, Andrés Ritter, Jean-Pierre Bouly, Angela Falciatore

Keywords: endogenous oscillators, circadian, protein-protein interaction, Yeast two Hybrid, cellular fluorescence rhythmicity

I was fully responsible for the sequence characterization of *bHLH1b* KO mutants, cellular fluorescence rhythmicity analysis and I contributed to the realization and analysis of the Yeast two Hybrid assay
Abstract

In eukaryotes, the mechanisms responsible for generating endogenous biological rhythms of gene expression require the action of multiple molecular players involved in the positive and negative branches of a transcriptional-translational feedback loop. In the case of the circadian clock of diatoms, to date only the *bHLH-PAS* transcription factor *RITMO1* has been identified as a component of the circadian clock of the model species *Phaeodactylum tricornutum*. The characterization of the timekeeper of these unicellular algae consequently necessitates the identification of the partners that interact with *RITMO1* in the generation of cellular rhythms. In the case of *P. tricornutum*, a first candidate is the protein *bHLH1b*, also belonging to the *bHLH-PAS* family encoded by a gene located near the *RITMO1* locus and sharing with it a non-overlapping head-to-head arrangement. By means of a Yeast two Hybrid assay strategy we were able to demonstrate that *bHLH1b* and *RITMO1* physically interact via their *bHLH* domain in yeast cells, suggesting their co-implication in the *P. tricornutum* clock. Also, KO lines for *bHLH1b* show an alteration in the rhythmicity of cellular fluorescence, with in particular a shortening of the oscillation period when released in continuous light. These results, similar to those already observed in *RITMO1* mutants, confirm the involvement of *bHLH1b* in the diatom timekeeper and pave the way for further decryption of its architecture.
<figure><img src="image_98.png" /><figcaption>Figure 1. Genomic arrangement of *RITMO1* and *bHLH1b* loci</figcaption></figure>

Schematic representation of the genomic arrangement of the loci of *RITMO1* (Phatr3_J44962, position chr5:806,933-808,518 reverse strand) and *bHLH1b* (Phatr3_J44963, position chr5: 809,453-811,854 forward strand) on the chromosome 5 of *P. tricornutum*. Exons are represented as black boxes, the introns as white boxes. The head-to-head orientation of the two genes is highlighted by the directional arrows.

Introduction

The functional and molecular
analyses reported in the previous two chapters
clearly demonstrate an implication of the
bHLH-PAS protein RITMO1 in the
regulation of circadian rhythmicity in *P.
tricornutum*. Its activity as a transcription
factor still needs to be determined but given
the strong deregulation of rhythmic gene
expression in KO mutants, an implication of
RITMO1 at the level of a TTFL-like core
clock is plausible. However, a transcriptional-
feedback system typical of endogenous
timekeepers requires the implication of at least
a minimal oscillator involving one
transcription factor as the positive element and
one factor as the negative element of the loop.
Such a minimal oscillator can be found in the
green unicellular alga *Ostreococcus tauri*
(Corellou et al., 2009; Morant et al., 2010). An
alternative possibility is that RITMO1
interacts with other proteins to form
multiprotein complexes in the realization of its
regulatory function, as in the case of more
complex clocks found for example in animals
or plants (Creux and Harmer, 2019; Peschel
and Helfrich-Förster, 2011). The search for
other proteins involved in the regulation of
cellular rhythmicity in *P. tricornutum* is
therefore essential for a complete dissection of
the clock of this diatom.

A candidate-gene approach was
performed to find potential partners of
RITMO1. In order to select different
candidates, we first focused on transcription
factors with a strong daily or circadian

regulation and tested them via a Yeast two
Hybrid approach with RITMO 1 as bait. In
particular, a second *P. tricornutum* bHLH-
PAS transcription factor, bHLH1b
(Phatr3_J44963) represents a prominent
prime target for decrypting the *P. tricornutum*
clock. The gene encoding this protein is in fact
located adjacent to *RITMO1* on *P.
tricornutum* chromosome 5. The two genes
are arranged in a nonoverlapping 'head-to-
head' orientation (Fig.1). The start codons of
the two genes are located about 1,000bp apart
and the 5' upstream sequences of RITMO1
and bHLH1b therefore overlap, suggesting a
possible co-regulation of their expression. The
two transcription factors both exhibit rhythmic
and coordinated expression in light and dark
cycles, showing a maximum transcription peak
that anticipates the onset of darkness
regardless of the photoperiod of growth
(Annunziata et al., 2019). In addition, as
reported in chapter II.c, *bHLH1b* expression
is regulated in a circadian way, with a
persistence of its oscillations after transition to
continuous light. Also, *bHLH1b* transcription
is affected in continuous light in both
RITMO1a KO and OE lines, further stressing
its circadian regulation. As described before,
bHLH-PAS transcription factors
homodimerize or heterodimerize in order to
recognise and bind their target DNA
sequences (Jones, 2004), the possibility that
these two proteins cooperate in generating a
negative transcriptional/translational feedback
loop as partners of a heterodimer is therefore
plausible. The results reported in the
following paragraph represent the first
functional characterisation of bHLH1b
protein and hint for its plausible co-

implication in the regulation of *P. tricornutum*
circadian clock with RITMO1.
Materials and methods

Yeast two Hybrid (Y2H) Assay

Y2H assays were carried out
according to previously published protocols
(Cuéllar et al., 2013). Coding sequences of
target genes were cloned from ecotype Pt1
cDNA via Gateway recombination into the
pGADT7 and pGBKT7 yeast destination
vectors (Clontech). Gene IDs and primers
used for cloning are reported in Table 1. The
pGADT7 (Prey) vector allows the expression
of the protein of interest fused N-terminally to
the Activation Domain of the yeast GAL4
transcription factor, while the pGBKT7 (bait)
vector fuses the protein to the GAL4 DNA
Binding Domain. Transcription was
constitutively driven by the yeast *ADH1*
promoter in both vectors. *LEU2* and *TRP1*
auxotrophic markers were used for yeast
selection pGADT7 and pGBKT7
respectively. Prey and bait vectors were co-
transformed in *S. cerevisiae* strain PJ69-4
(*trp1-901*, *leu2-3*, *ura3-52* and *his3-200*)
following (Cuéllar et al., 2013). Positive
transformants were selected by auxotrophy on
selective plates of Complete Supplement
Mixture medium *-leu-trp* (CSM-LW). Positive
bait-prey interaction was assessed by growth
on CSM *-leu-trp-his* (CSM-LWH). In case of
interaction between bait and prey hybrid
proteins, the reconstitution of the GAL4
transcription factor drives the expression of
the *HIS3* reporter under the control of the
UAS promoter, allowing the growth in
selective media lacking *his*. Baits were tested
for autoactivation of the *HIS3* reporter by co-
transforming the bait vector construct and an
empty prey. Three technical replicates were
realized for each prey-bait combination.

Culture and irradiation conditions

Transgenic and control lines of *P. tricornutum* used for bHLH1b characterization share a Pt1 8.6; CCMP2561 genomic background. Cell cultures were

grown in batch culture in F/2 Guillard media
(Guillard, 1975). Before performing
experiments, cells were pre-adapted for two
weeks to 16L:8D at 25 µmol photons m⁻² s⁻¹.
Final refresh of culture was performed 3 days
before the release to free-run conditions (17
µmol photons m⁻² s⁻¹) to ensure minimal
perturbation of physiology due to culture
refreshment. For all experiments the
temperature was set constant to 18°C and
illumination was performed with white light
(Bridgelux BXRV-DR-1830H-3000-A-13
LEDs). Change in light intensity upon transfer
to L:L was performed at the moment of dark
to continuous light transition.

CRISPR/Cas9 Mutagenesis

*bHLH1b* KO mutants were
generated following (Nymark et al., 2016) by
biolistic bombardment transformation of a
pKS diaCAS9 sgRNA vector (Addgene ID:
74923). This vector contains a hSpCAS9 gene
designed using the *P. tricornutum* codon
usage and under the control of LHCF2
promoter and LHCF1 terminator sequences,
plus a guide RNA sequence under the control
of the *P. tricornutum* U6 snRNA promoter
and U6 terminator. The adapters for targeting
the *bHLH1b* gene were generated by
annealing complementary oligos with TCGA
and AAAC overhangs and cloned via BsaI
digestion at the 5' -end of the guide RNA
region (Nymark et al., 2016). Two sequences
for *bHLH1b* adapters (named gHLH4 and
gHLH5, Fig.3A) were designed via CRISPOR
web tool (Concordet and Haeussler, 2018)
and cloned in two independent pKS
diaCAS9 sgRNA. Target sequences of
sgRNA are reported in Table 2 The two
CRISPR/CAS9-sgRNA vectors were co-
transformed with the pAF3 plasmid (Falciatore
et al., 1999) containing the Sh ble gene
conferring resistance to phleomycin. Full
<table><caption>Table 1. List of the oligonucleotides used in this work.</caption><thead><tr><th>Gene</th><th>Accession</th><th>Name</th><th>Strand</th><th>Sequence</th><th>Type</th></tr></thead><tbody><tr><td rowspan="4">RITMO1</td><td rowspan="4">Phatr3_J4496</td><td>FL_bHIH1a_GW_FW</td><td>Fw</td><td>GGGGACAAGTTTGTACAAAAAGCAGGCTCCATGAATAAGCCAGGACAGCG</td><td>Full length Gateway Cloning</td></tr><tr><td>bHIH1a_GW_RV_bHIHd</td><td>Rv</td><td>GGGGACCACCTTTGTACAAGAAAGCTGGGTCTCAGTCTTCGGGTGTCGGTTCTG</td><td>bHLH domain Gateway Cloning</td></tr><tr><td>bHIH1a_GW_FW_PASd</td><td>Rv</td><td>GGGGACAAGTTTTGTACAAAAAAGCAGGCTCCATGGAAACTCTAGTTTTTGTCCG</td><td>PAS domain Gateway Cloning</td></tr><tr><td>FL_bHIH1a_GW_RV</td><td>Rv</td><td>GGGGACCACCTTTGTACAAGAAAGCTGGGTCTCACAGCTTCGCTGCATCGT</td><td>Full length Gateway Cloning</td></tr><tr><td rowspan="6">bHLH1b</td><td rowspan="6">Phatr3_J44963</td><td>FL_bHLH1b_GW_Fw</td><td>Fw</td><td>GGGGACAAGTTTGTACAAAAAAGCAGGCTCCATGAATTCTTTAAGCGCAGCGG</td><td>Full length Gateway Cloning</td></tr><tr><td>bHIH1b_GW_RV_bHIHd</td><td>Rv</td><td>GGGGACCACCTTTGTACAAGAAAGCTGGGTCTCACAGCAATCGCTTGCTTCTCGA</td><td>bHLH domain Gateway Cloning</td></tr><tr><td>FL_bHLH1b_GW_Rv</td><td>Rv</td><td>GGGGACCACCTTTGTACAAGAAAGCTGGGTCTCAACAGTAGTCGTGACTACCTTCA</td><td>Full length Gateway Cloning</td></tr><tr><td>bHIH1b_sequencing_Rv</td><td>Fw</td><td>CCAATGTTCACGTTCCCT</td><td>sequencing</td></tr><tr><td>bHIH1b_sequencing_Fw</td><td>Rv</td><td>CACTCTTTGTCTTCTTCACT</td><td>sequencing</td></tr><tr><td>CPF1</td><td>Phatr2_27429</td><td>FL_CPF1_GW_FW</td><td>Fw</td><td>GGGGACAAGTTTTGTACAAAAAGCAGGCTCCATGGCTAAATCGGAAGAGAAA</td><td>Full length Gateway Cloning</td></tr><tr><td rowspan="3">bZIP7</td><td rowspan="3">Phatr3_J48800</td><td>FL_CPF1_GW_RV</td><td>Rv</td><td>GGGGACCACCTTTGTACAAGAAAGCTGGGTCTCAGTTGCGACGTTGTCGCTTTGGT</td><td>Full length Gateway Cloning</td></tr><tr><td>FL_bZIP7_GW_FW</td><td>Fw</td><td>GGGGACAAGTTTTGTACAAAAAAGCAGGCTCCATGCGTAATCAAGGATTAGAGGCAT</td><td>Full length Gateway Cloning</td></tr><tr><td>FL_bZIP7_GW_RV</td><td>Rv</td><td>GGGGACCACCTTTGTACAAGAAAGCTGGGTCTCAGGCGGTTCCATTGAGCCCATCA</td><td>Full length Gateway Cloning</td></tr></tbody></table>

Target gene, name, pairing strand, sequence and use of the oligonucleotides used in this work.

<table><caption>Table 2. sgRNA used for bHLH1b KOs generation</caption><thead><tr><th>Target gene</th><th>Accession</th><th>gRNA name</th><th>Strand</th><th>SEQUENCE</th><th>3' PAM</th></tr></thead><tbody><tr><td rowspan="2">bHLH1b</td><td rowspan="2">Phatr3_J44963</td><td>bHLHg4</td><td>Fw</td><td rowspan="2">AGCGCATCACACCGAAGCT</td><td>TGG</td></tr><tr><td>bHLHg5</td><td>Fw</td><td>CGG</td></tr></tbody></table>

List of the sgRNA used for CRISPR-Cas9 mutagenesis. Target gene, sgRNA name, target strand, gRNA sequence and the associated protospacer adjacent motif (PAM) are reported.
length sequence of the bHLH1b gene was
amplified by PCR on the phleomycin-resistant
colonies obtained (primers reported in Table
2). PCR fragments were purified from agarose
gel with QIAquick Gel Extraction Kit
(QIAGEN) and analyzed via Sanger
sequencing using bHLH1b specific primers
reported in Table 1 in order to retrieviallelic
mutants.

Flow cytometry and
fluorescence rhythmicity analysis

Cellular fluorescence rhythmicity
assays were performed using a MACSQuant
Analyser flow cytometer (Miltenyi Biotec). B3-
A parameter (488 nm excitation, 655-730 nm
detection) was used to track chlorophyll
fluorescence (Annunziata et al., 2019). 30.000
events were analyzed per each time point per
culture replicate and median B3-A value was
used for following rhythmicity analysis.
Culture samples were collected every 2 hours
via a BVP standard ISM444 peristaltic pump

(Ismatec) controlled by a FC204 fraction
collector (Gilson). Cells were stored at 4°C in
the dark for at least 2 hours to a maximum of
24hours previous to flow cytometry. The
analysis of B3-A fluorescence dataset
rhythmicity was performed using the
BioDare2 webtool
(https://biodare2.ed.ac.uk/, (Zielinski et al.,
2014). For graphical representation, data
traces were baseline detrended, normalized to
the maximum value and aligned to the mean
values. Data traces were first analyzed using
the Enright Periodogram (EPR) algorithm,
which was used as an arrhythmicity test
(Zielinski et al., 2014). Period, amplitude and
RAE were calculated via Fast Fourier
nonlinear least square algorithm (FFT-NLLS),
within a period range of 18 and 34 hours. At
least three full oscillations were considered for
the analysis. Statistical differences in
rhythmicity parameters among lines were
evaluated via One-way analysis of variance
against Wt with Bonferroni post-test
correction.
Results

bHLH1b and RITMO1 physically
interact via their bHLH fold

In order to investigate the regulation
network in which RITMO1 is involved, we
performed a series of Y2H assays aimed to
identify interacting proteins potentially
involved in *P. tricornutum* clock machinery. A
first round of analysis was carried out by
assessing the possible autoactivation of
RITMO1 protein in a Y2H system.
Autoactivation experiments are aimed to
assess whether the protein fused to the GAL4
BD is able to recruit the transcription
machinery and express *HIS3* on its own
without a prey hybrid (Cuéllar et al., 2013). As
shown in Fig.4A, RITMO1 is able to
autonomously recruit the transcription
machinery and to induce the expression of
*HIS3* reporter. The autoinduction of
RITMO1 in the yeast model made it
impossible to perform a broad-spectrum
screen of its possible protein interactors.
Nevertheless, we decided to test the possible
interaction of RITMO1 with a restricted set of
proteins of interest: bHLH1b, BZIP7 and
CPF1. bHLH1b and bZIP7 are transcription
factor belonging to bHLH-PAS and the basic-
leucine zipper (bZIP) family respectively
which have a temporal expression profile
similar to that of RITMO1 (Annunziata et al.,
2019). CPF1 (cryptochrome photolyase family
1) was chosen for Y2H analysis since it was
shown to repress the activity of
CLOCK:BMAL1-induced reporter genes in a
heterologous mammalian cell system (COS7
cell line from *Cercopithecus aethiops*), thus
phenocopying the function of the endogenous
animal cryptochrome (CRY, Coesel et al.,
2009). Since mammalian CRY is known to
physically bind the bHLH-PAS protein
BMAL1-1 (Tamaru et al., 2015), we wanted to
assess whether CPF1 can display similar

dimerization properties with RITMO1 and/or
bHLH1b.

Coding sequences of the four proteins
were fused both to the BD (bait) and the AD
(prey) of GAL4, for a total of eight constructs.
First, the bait hybrids of bHLH1b, bZIP7 and
CPF1 were tested for autoactivation by co-
transforming them with empty pGADT7
plasmid. None of the other baits showed the
same autoactivation outcome showed by
RITMO1, thus excluding that the remaining
proteins under investigation can activate gene
expression on their own. As an additional
negative control, the four prey hybrids were
co-transformed with empty pGBKT7
(Fig.2A). None of these controls gave rise to
colonies on the CSM-LWH selective
medium.

bHLH1b, bZIP7 and CPF1 bait
hybrids were subsequently co-transformed
with RITMO1 prey in order to test their
possible physical interaction in yeast cells (Fig.
2B). In addition, the bHLH1b bait hybrid was
also co-transformed with the bHLH1b prey
hybrid to test homodimerization. This round
of analysis demonstrated the interaction
between RITMO1 and its paralogous
bHLH1b and the homo-dimerization of
bHLH1b, confirmed by the growth on
selective CSM-LWH media (Fig.2B). None of
the colonies bearing other combinations of
preys and baits grew on selective plates,
excluding other interactions, at least in the
yeast model.

Finally, to refine our understanding of
the mutual interaction between RITMO1 and
bHLH1b, a set of Y2H was carried out by
splitting RITMO1 gene in its bHLH and PAS
coding sequences and cloning them into
pGADT7 and pGBKT7. Similar construction
was performed with the bHLH fold of
bHLH1b (Fig. 2C). First, the bait hybrids of
bHLH and PAS domains of RITMO1 were
<figure><img src="image_99.png" /></figure>

<figure><img src="image_100.png" /></figure>

<figure><img src="image_101.png" /></figure>

<figure><img src="image_102.png" /><figcaption>Figure 2. Results of the Yeast two Hybrid assay (Y2H)</figcaption></figure>

(A) Results of the autoactivation test of the RITMO1, bHLH1b, bZIP7 and CPF1 full-length proteins cloned into bait and prey vectors. (B) Results of the Y2H interactions between the selected proteins co-transformed in yeast cells in different combinations of bait and prey hybrids. (C) Autoactivation tests of the RITMO1 bHLH and PAS folds in bait vector and analysis of the interaction between RITMO1 and bHLH1b bHLH and PAS folds. Top photos refer to cells grown in control medium (CSM-LW), bottom photos refer to cells grown in selective medium (CSM-LWH). (D) Schematic representation of the results of the Y2H assays showing the dimerization of bHLH1b with RITMO1 via the bHLH fold and the homodimerization of bHLH1b. The possible homodimerization of RITMO1 could not be assessed due to autoactivation of *His3* reported by RITMO1 bait constructs
co-transformed with the empty pGADT7 to assess which of the two regions is responsible for autoactivation (Fig.2C). Growth on selective media was observed only in yeast bearing the bHLH fold bait, thus hinting that this domain is accountable for recruiting the transcription machinery in yeast cells. Co-transformation of bHLH1b-bHLH bait hybrid with RITMO1-bHLH prey hybrid was also performed as well as full length bHLH1b bait with RITMO1 PAS prey hybrids co-transformation. Results indicate that RITMO1 and bHLH1b interact via their bHLH regions, which are known to represent the primary dimerization surface of bHLH-PAS family members (Kewley et al., 2004) (Fig.2C). A schematic summary of the interactions emerged in the Y2H assay is reported in Fig.2D.

## Generation of bHLH1b KO mutants

Following the results of the Y2H analysis, CRISPR/Cas9 mutagenesis of

bHLH1b locus was performed to test the obliteration of this protein on cellular rhythmicity (Fig.3). The transformation of *P. tricornutum* cells with the pKS diaCAS9 sgRNA vector lead to the generation of two biallelic KO lines for bHLH1b (named KO 1b#1 and KO 1b#2 respectively) identified by Sanger sequencing. Sequencing analysis of the *bHLH1b* locus highlighted a 85 bp insertion at the level of gHLH5 target region in the allele 2 of KO 1b#1, a 128 bp deletion in allele 1 of KO 1b#2 and a 848 bp deletion in allele 2 (Fig. 3B and Fig.4). All mutagenic events induce a frameshift mutation that disrupts downstream protein sequence. Allele 1 of KO 1b#1 was impossible to amplify via PCR and was not found in Sanger sequencing chromatograms, suggesting that large insertions or deletions were induced by CRISPR/Cas9 mutagenesis. This phenomenon is defined as Loss of Heterozygosity and is not uncommon in CRISPR/Cas9 targeted mutagenesis (Serif et al., 2018).

<figure><img src="image_103.png" /></figure>

<table><caption>Figure 3. Generation of bHLH1b KO mutants</caption><thead><tr><th>Strain name</th><th>Type of mutation</th></tr></thead><tbody><tr><td>KO 1b#1</td><td>Allele 1: impossible to amplify, possible Loss of Heterozygosity<br>Allele2: 85bp insertion at position nt-917 in the gHLH5 region</td></tr><tr><td>KO 1b #2</td><td>Allele1: deletion of 128bp from position nt-917 in the gHLH5 region<br>Allele2: deletion of 848-bp from position nt-82</td></tr></tbody></table>

(A) Schema of the *bHLH1b* locus with the sgRNA target sites highlighted. Blue and light blue boxes represent the relative location of the bHLH and PAS domains respectively. Light and dark green boxes highlight the localisation of the target sequences (underlined) of the three gRNAs used for bHLH1b CRISPR-Cas9 mutagenesis. Letters in bold represent the protospacer adjacent motif associated with each target site. (B) Description of the mutations induced on each allele of the *bHLH1b* locus on the two KO mutants as identified by Sanger sequencing.
A

Reference sequence (1): *Wt_Allele_1*
Identities normalised by reference length.
Colored by: Identity

<table><thead><tr><th></th><th>cov</th><th>pid</th><th></th><th></th><th></th><th></th></tr></thead><tbody><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>1</td><td>ATGAATTCTTTAAGCGCAGCGGCTGCAGAAGGGGCCAGTATGGCAGCTCGCTCAAATAACGCTAGCTACAGGAAGGCGGAGACAACGAGGTGGTTTCGCCTCCTTCGAGCTCTTGAT</td><td>120</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>ATGAATTCTTTAAGCGCAGCGGCTGCAGAAGGGCCAGTATGGCAGCTCGCTCAAATAACGCTAGCTACAGGAAGGCGGAGACAACGAGGTGGTTTCGCCTCCTTCGAGCTCTTGAT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>321</td><td>ACGAAGGCGATGAAAATTTACAGAGGGACATCAGAAGTCCGCATATCGCTGCTCTTGGAGTGAATGCTTGCTGTAAACACGCTAACTGTGTCAAATGACTTACCAACGTTATTATCGCAG</td><td>240</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>ACGAAGGCGATGAAAATTTACAGAGGGACATCAGAAGTCCGCATATCGCTGCTCTTGGAGTGAATGCTTGCTGTAAACACGCTAACTGTGTCAAATGACTTACCAACGTTATTATCGCAG</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>241</td><td>ATGACAGCAGCACCTGTACCCTCAGCATCAACGCAAGCAGGACCATCAAGTCAACAACGAAGTCTTTGGCAGCAAGGTCGACGACACTCGCTGGGAGTACTTTT</td><td>360</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>ATGACAGCAGCACCTGTACCCTCAGCATCAACGCAAGCAGGACCATCAAGTCAACAACGAAGTCTTTGGCAGCAAGGTCGACGACACTCGCTGGGAGTACTTTT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>361</td><td>CCTATGCCTACGCAGCCACTGGCAACGACATGGGCAGCAACCATGTGCCAATGTTCACGTTCCTTAGTCTATGCAACAATTTCAGTCAATACATTCGGTTCTGGAACAGGAAT</td><td>480</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>CCTATGCCTACGCAGCCACTGGCAACGACATGGGCAGCAACCATGTGCCAATGTTCACGTTCCTTAGTCTATGCAACAATTTCAGTCAATACATTCGGTTCTGGAACAGGAAT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>481</td><td>AGCGATGTTAACGCGGGGGGGCTGCTGCCTATGGTGAATGCAACAAATGTTGTTTCAGCAGCGCAACCCGCAACTATACATCAAGGTCACGACCACCGCTTCCTGCTAACACGGTT</td><td>600</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>AGCGATGTTAACGCGGGGGGGCTGCTGCCTATGGTGAATGCAACAAATGTTGTTTCAGCAGCGCAACCCGCAACTATACATCAAGGTCACGACCACCGCTTCCTGCTAACACGGTT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>601</td><td>CAAACGGGAAGCGCATCACAAACCGAAGCTTGGCCAGTACCCGTAAACAGAAGCAAACAAGGAATGACCGAAAAGTGATACAAGGAAGTGAAGAAAGCAAAGGCTCAGAAGTTGGAACCA</td><td>720</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>CAAACGGGAAGCGCATCACAAACCGAAGCTTGGCCAGTACCCGTAAACAGAAGCAAACAAGGAATGACCGAAAAGTGATACAAGGAAGTGAAGAAAGCAAAGGCTCAGAAGTTGGAACCA</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>721</td><td>TCTGTTGTGTCTTTCTCCAGTTACATTTCAGGGGACTCTTCTTCAAAGACAGCCAACGACGAAGCTCTTTCATTGGCTCAAAGAACCCCGATTGGGCAAGATGACCCCACCGCAGGT</td><td>840</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>TCTGTTGTGTCTTTCTCCAGTTACATTTCAGGGGACTCTTCTTCAAAGACAGCCAACGACGAAGCTCTTTCATTGGCTCAAAGAACCCCGATTGGGCAAGATGACCCCACCGCAGGT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>841</td><td>CGTCGACATGAGCGAAATATGAGGGAGCAACAGCGTCGGGTCTCATATCGCAACAGATCAAGGAGCTACAGAGGAGAGGTTCTGACGAGTAAATATCCTTTCAAACCAACAAGTTTTCC</td><td>960</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>CGTCGACATGAGCGAAATATGAGGGAGCAACAGCGTCGGGTCTCATATCGCAACAGATCAAGGAGCTACAGAGAG</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>961</td><td>ATCCTTGTCCTGTAGTTGACTACATCAATCATTGCAAGCCAGAGCCATCATGGCTGACTCAGAACAATCAAAACGCTGGCAGACTTATCGAGAAGCAACGATTGGTTCACCAACGGG</td><td>1080</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>ATCCTTGTCCTGTAGTTGACTACATCAATCATTGCAAGCCAGAGCCATCATGGCTGACTCAGAACAATCAAAACGCTGGCAGACTTATCGAGAAGCAACGATTGGTTCACCAACGGG</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>1081</td><td>TTAACTAGTGAAGAAGACAAAGAGTCCATGTCTCCACTTCCGACGAGCGCAAAAGTGAGCTGCTTTTCGTACAAGGAATTGACTACCGTGCTGTTTTCAATCATTGCCATTTGCCATG</td><td>1200</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>TTAACTAGTGAAGAAGACAAAGAGTCCATGTCTCCACTTCCGACGAGCGCAAAAGTGAGCTGCTTTTCGTACAAGGAATTGACTACCGTGCTGTTTTTCGTACAATCATTGCCATTTGCCATG</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>1201</td><td>GGGGTGGCTTCGTTGGACGGAAGAAATACTGGTCTGCAACAGCTCTTTCGAATCTCTGTGGGCCTAAATAAGGACGGGCTGCAGCAGAGCTTTTCGGTTGTTATCCCGAATCCCTGAAACACGGTT</td><td>1320</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>GGGGTGGCTTCGTTGGACGGAAGAAATACTGGTCTGCAACAGCTCTTTCGAATCTCTGTGGGCCTAAATAAGGACGGGCTGCAGCAGAGCTTTTCGGTTGTTATCCCGAATCCCTGAAACACGGTT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>1321</td><td>GATAATATTTGACGCCATGGCAGACCTGCTAAAACGCTCAAGTGTGCCAGTGAACAGGGGAAGGAGTCGTAGTAAAGGAAGAACATTTGTTGTACTGGTGGACAGGTAGTGACTTTG</td><td>4</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>GATAATATTTGACGCCATGGCAGACCTGCTAAAACGCTCAAGTGTGCCAGTGAACAGGGGAAGGAGTCGTAGTAAAGGAAGAACATTTGTTGTACTGGTGGACAGGTAGTGACTTTG</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_1</td><td>100.0%</td><td>100.0%</td><td>1441</td><td>CGATCTAACGGCTAATATTCAATCACAGCTGACTAGTTCTTCGAACGGAGAGCCAAGTTTTTCGGTTCTTCAGCTGCTGAAGGTAGTCACACTGACTTAGT</td><td>5</td></tr><tr><td>2</td><td>KO2_Allele_1</td><td>91.7%</td><td>91.7%</td><td></td><td>CGATCTAACGGCTAATATTCAATCACAGCTGACTAGTTCTTCGAACGGAGAGCCAAGTTTTTCGGTTCTTCAGCTGCTGAAGGTAGTCACACTGACTTAGT</td><td></td></tr></tbody></table>

B

Reference sequence (1): *Wt_Allele_2*
Identities normalised by reference length.
Colored by: Identity

<table><thead><tr><th></th><th>cov</th><th>pid</th><th></th><th></th><th></th><th></th></tr></thead><tbody><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>1</td><td>ATGAATTCTTTAAGCGCAGCGGCTGCAGAAGGGGCCAGTATGGCAGCTCGCTCAAATAACGCTAGCTACAGGAAGGCGGAGACAACGACGCGGTGGTTTCGCCTCCTCCTGCAGCTCTTGAT</td><td>120</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>ATGAATTCTTTAAGCGCAGCGGCTGCAGAAGGGCCAGTATGGCAGCTCGCTCAAATAACGCTAGCTACAGGAAGGCGGAGACAACGACGCGGTGGTTTCGCCTCCTCCTGCAGCTCTTGAT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>321</td><td>ACGAAGGCGATGAAAATTTACAGAGGGACATCAGAAGTCCGCATATCGCTGCTCTTGGAGTGAATGCTTGTAACACGCTAAATGACTTCCACGTTATTATCGCAG</td><td>240</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>ACGAAGGCGATGAAAATTTACAGAGGGACATCAGAAGTCCGCATATCGCTGCTCTTGGAGTGAATGCTTGTAACACGCTAAATGACTTCCACGTTATTATCGCAG</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>241</td><td>ATGACAGCAGCACCTGTACCCTCAGCATCAACGCAAGCAGAGACATCAAGTCAACAACGAAGTCTTTGGCAGCAAGGTCGACACTCGCTGGGAGTACTTTT</td><td>360</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>ATGACAGCAGCACCTGTACCCTCAGCATCAACGCAAGCAGAGACATCAAGTCAACAACGAAGTCTTTGGCAGCAAGGTCGACACTCGCTGGGAGTACTTTT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>361</td><td>CCCATGCCTACGCAGCCACTGGCAACGACATGGGCAGCAACCATGTGCCAATGTTCACGTTCCTTAGTCTATGCAACAATTTCAGTCAATACATTCGGTTCTGGAACAGGAAT</td><td>480</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>CCCATGCCTACGCAGCCACTGGCAACGACATGGGCAGCAACCATGTGCCAATGTTCACGTTCCTTAGTCTATGCAACAATTTCAGTCAATACATTCGGTTCTGGAACAGGAAT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>481</td><td>AGCGATGTTAACGCGGGGGGAGCTGCTGCTATGGTGAATGCAACAAATGTTGTTTCAGCAGCGCAACCCGCAACTACACATCAAGGTCACGACCACCGCTTCCTGCTAACACGGTT</td><td>600</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>AGCGATGTTAACGCGGGGGGAGCTGCTGCTATGGTGAATGCAACAAATGTTGTTTCAGCAGCGCAACCCGCAACTACACATCAAGGTCACGACCACCGCTTCCTGCTAACACGGTT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>601</td><td>CAAACGGGAAGCGCATCACAAACCGAAGCTTGGCCAGTACCCGTAAACAAGAAGCAAACAAGGACGTACCAGGAAAAGTGATTACAAAGAAGTGAAGAAAGCAAAGGCTCAGAAGTTGGAACCA</td><td>720</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>CAAACGGGAAGCGCATCACAAACCGAAGCTTGGCCAGTACCCGTAAACAAGAAGCAAACAAGGACGTACCAGGAAAAGTGATTACAAAGAAGTGAAGAAAGCAAAGGCTCAGAAGTTGGAACCA</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>721</td><td>TCTGTTGTGTCTTTCTCCAGTTACATTTCAGGGGACTCTTCTTCAAAGACAGCCAACGACGAAGCTCTTTCATTGGCTGCAAGAAGACCCCGATTGGGCAAGATGACCCCACCGCAGGT</td><td>840</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>TCTGTTGTGTCTTTCTCCAGTTACATTTCAGGGGACTCTTCTTCAAAGACAGCCAACGACGAAGCTCTTTCATTGGCTGCAAGAAGACCCCGATTGGGCAAGATGACCCCACCGCAGGT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>841</td><td>CGTCGACATGAGCGAAATATGAGGGAGCAACAGCGTCGGGTCTCATATCGCAACAGATCAAGGAGCTACAGAGAGGAGAGGTTCTGACGAGTAAATATCCTTTCAAACCAACAAGTTTTCC</td><td>960</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>CGTCGACATGAGCGAAATATGAGGGAGCAACAGCGTCGGGTCTCATATCGCAACAGATCAAGGAGCTACAGAGAGGAGAGGTTCTGACGAGTAAATATCCTTTCAAACCAACAAGTTTTCC</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>961</td><td>AGGTGACGCTCAATCATTTGTTGTTTCTTCAATCTTTTACTGACGAGGTAACAAATATCCTTTCAAACCAACAAGTTTTCCATCTTGTCTCTGTAGTTGACTACATCAACAAT</td><td>1080</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>AGGTGACGCTCAATCATTTGTTGTTTCTTCAATCTTTTACTGACGAGGTAACAAATATCCTTTCAAACCAACAAGTTTTCCATCTTGTCTCTGTAGTTGACTACATCAACAAT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>1081</td><td>TGCAAGCCAGAGCCATCATGCTGGACTCAGACAATCAAAACTGGCGACACTATTGAGAAGCAAGCGATTGTTGTCACCAACGGGTTAACTAGTGAAGAAGACAAGAGTGCATGTCTC</td><td>120</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>TGCAAGCCAGAGCCATCATGCTGGACTCAGACAATCAAAACTGGCGACACTATTGAGAAGCAAGCGATTGTTGTCACCAACGGGTTAACTAGTGAAGAAGACAAGAGTGCATGTCTC</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>1201</td><td>CACTTTCCGACGAGCCGCAAAGTGTGACTGCTGTTTTCGTACAAGGAATTGACTACCGTGCTGTTTTCAATCATTGCCATATGCCATGGGGTGGCTTCATTGGACGGAAGAATACTGGTCT</td><td>3</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>CACTTTCCGACGAGCCGCAAAGTGTGACTGCTGTTTTCGTACAAGGAATTGACTACCGTGCTGTTTTCAATCATTGCCATATGCCATGGGGTGGCTTCATTGGACGGAAGAATACTGGTCT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>1321</td><td>GCAACAGCTCTTTCGAATCTCTTTGGGCCTAAATAAGACCGGCTCGACAACAGAGCTTTTCGTGTTTATCCGGAACATCAAGATATTTGACGCCATGGCAGACCTGCTTAAAC</td><td>4</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>GCAACAGCTCTTTCGAATCTCTTTGGGCCTAAATAAGACCGGCTCGACAACAGAGCTTTTCGTGTTTATCCGGAACATCAAGATATTTGACGCCATGGCAGACCTGCTTAAAC</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>1441</td><td>GCTAAGTGTGTTCCAGTGAAACAGGGGAAGGGGTCGTAGTAAGAAAGGAACACATTTGTTGTACTGGTGGACAGGTAGTGACTTTGCGATCTCAACGCGCTAATATTCAATCACACGCTGA</td><td>1560</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>GCTAAGTGTGTTCCAGTGAAACAGGGGAAGGGGTCGTAGTAAGAAAGGAACACATTTGTTGTACTGGTGGACAGGTAGTGACTTTGCGATCTCAACGCGCTAATATTCAATCACACGCTGA</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>1</td><td>Wt_Allele_2</td><td>100.0%</td><td>100.0%</td><td>1561</td><td>CTAGTTCTTCGAACGGAGAGCCAAGTTTTTCGGTTCTTCAGCTGCTGAAGGTAGTCACACTGACTTAGT</td><td>6</td></tr><tr><td>2</td><td>KO2_Allele_2</td><td>40.6%</td><td>40.6%</td><td></td><td>CTAGTTCTTCGAACGGAGAGCCAAGTTTTTCGGTTCTTCAGCTGCTGAAGGTAGTCACACTGACTTAGT</td><td></td></tr><tr><td></td><td></td><td></td><td></td><td></td><td></td><td></td></tr></tbody></table>
Figure 4. Characterisation of bHLH1b KO mutants

Previous page: (A) Sequence alignment of bHLH1b allele 1 in Pt1 Wt strain and KO 1b#2. Allele 1 of KO 1b#1 was not possible to amplify, possibly due to Loss of Heterozigosity (B) Sequence alignment of RITMO1 allele 2 in Pt1 Wt strain and RITMO1 and KO 1b#1 KO 1b#2. Sequence alignments were generated via EBI Clustal Omega webtool and visualized via Mview tool. Bases are colored by identity to the Wt reference sequence. gHLH4 and gHLH5 sgRNA target sites are highlighted in light and dark green respectively.

bHLH1b mutations affect free-run fluorescence oscillations

KO 1b#1 and #2 lines were characterized by flow cytometry in order to assess possible alteration of cellular rhythmicity, as previously done for the RITMO1 mutants. Transgenic cultures were pre-adapted in a 16L:8D photoperiod (25 µmol photons m⁻² s⁻¹) and subsequently released under constant light at 17 µmol photons m⁻² s⁻¹, choosing the experimental setting already optimized in Chapter II.c. As reported in Fig.4, all replicates of bHLH1b KOs were shown to maintain a rhythmicity in fluorescence oscillations in free-run condition. Nevertheless, both KO mutants show a shortening of the oscillation period when

compared to Wt lines, although to a minor extent to what was previously shown on RITMO1 KO mutants. Free run period is indeed equal to 27.13±0.62 and 27.33±0.44 hours in KO 1b#1 and KO 1b#2 respectively, versus the 29.62±0.63 period shown by Wt replicates in the same experiments (Fig.5 and Table 3). In distinct experiments with the same experimental set up, RITMO1 KO1 and KO3 showed a period of 24.21±0.91 and 21.97±0.82 (mean±s.e.m.) respectively, as reported in Chapter II.c. In addition, amplitude and R.A.E. of free run oscillations in bHLH1b KO lines show no alteration when compared to Wt line (Fig.5 and Table 3), differently to what was observed in RITMO1 mutants.

<table><thead><tr><th rowspan="2"></th><th colspan="2">Wt (6/6)</th><th colspan="3">KO 1b#1 (10/10)</th><th colspan="3">KO 1b#1 (10/10)</th></tr><tr><th>Mean</th><th>s.e.m</th><th>Mean</th><th>s.e.m</th><th>p value</th><th>Mean</th><th>s.e.m</th><th>p value</th></tr></thead><tbody><tr><td>Period</td><td>29.62</td><td>0.6348</td><td>27.15</td><td>0.6163</td><td>0.02</td><td>27.33</td><td>0.4353</td><td>0.0084</td></tr><tr><td>R.A.E.</td><td>0.36</td><td>0.04733</td><td>0.423</td><td>0.04626</td><td>N.S.</td><td>0.391</td><td>0.04086</td><td>N.S.</td></tr><tr><td>Amplitude</td><td>0.7117</td><td>0.04764</td><td>0.754</td><td>0.05177</td><td>N.S.</td><td>0.752</td><td>0.03747</td><td>N.S.</td></tr></tbody></table>

Table 3. Characterisation of cellular fluorescence rhythmicity of bHLH1b mutants in L:L

Calculated periods, relative amplitude errors and amplitudes of the Wt, KO 1b#1 and KO 1b#2 lines after transition from 16L:8D 25µmol m⁻² s⁻¹ to L:L 17µmol m⁻² s⁻¹. Average and s.e.m. values were calculated on the replicates considered rhythmic according to the EPR test. The fraction of rhythmic replicates/total of replicates is reported and correspond to the whole set of replicates. Values correspond to what is shown in Fig., panels (A-C). P-values obtained via One-way analysis of variance against Wt with Bonferroni post-test correction are shown. N.S.: non significant, P>0.05
<figure><img src="image_104.png" /></figure>

<figure><img src="image_105.png" /></figure>

<figure><img src="image_106.png" /></figure>

<figure><img src="image_107.png" /></figure>

<figure><img src="image_108.png" /></figure>

D

<figure><img src="image_109.png" /></figure>

E

<figure><img src="image_110.png" /></figure>

F

<figure><img src="image_111.png" /></figure>

H

<figure><img src="image_112.png" /><figcaption>Figure 5. Effects of bHLH1b mutation on cellular fluorescence</figcaption></figure>

(A-C) Oscillations of cellular red fluorescence (B3-A parameter) in Wt, KO 1b#1 and KO 1b#2 after transition from 16L:8D 25µmol m⁻² s⁻¹ to L:L 17µmol m⁻² s⁻¹ (n=6 for Wt, n=10 for KO lines). Dots represent mean B3-A values, coloured dashed lines represent SD. (D-F) Period (D), amplitude (E) and R.A.E. (F) estimations of B3-A fluorescence traces represented in panels A-C (mean ±s.e.m., *P<0.05, **P <0.01, One-way analysis of variance against Wt with Bonferroni post-test correction). (G) Plot of period versus R.A.E. estimates and (H) plot of amplitude versus R.A.E. estimates of the replicates represented in panels A-C.
Conclusions

Although preliminary, the results
presented here suggest a possible co-
implication of bHLH1b in the regulation of
the circadian clock of *P. tricornutum* together
with RITMO1. The data obtained through a
limited range of Y2H analyses allowed us to
demonstrate a direct physical interaction
between RITMO1 and bHLH1b, an
interaction that is achieved through their
bHLH fold. Similarly, it was possible to
highlight the homodimerization of bHLH1b,
while homodimerization of RITMO1 was not
possible to assess due to the autoactivation of
the *His3* reporter in *S. cerevisiae* cells
transformed with the RITMO1-bait hybrid. In
parallel, no direct protein-protein association
in the yeast model was observed between
RITMO1 and other proteins presenting a
similar rhythmic expression profile in *P.
tricornutum*, the transcription factor bZIP7
and the photoreceptor CPF1. It is crucial to
consider that the results obtained do not
exclude that post-translational modifications
(e.g. acetylation or phosphorylation of amino
acid residues) may be required for the
interaction of RITMO1 with CPF1 and
bZIP7, since the Y2H setup does not allow to
control the post-translational modifications of
the protein under study. In mammals, CRY
interaction with BMAL1 via the bHLH fold is
enhanced by post translational modifications
of BMAL1, specifically CK2α-mediated
phosphorylation and CLOCK-mediated
acetylation (Tamaru et al., 2015). The homo-
dimerization of RITMO1 remains an open
possibility which could not be assessed in this
experiment due to the autoactivation of the
RITMO1-bait hybrid. Cellular fluorescence
data obtained by flow cytometer analysis
confirm a statistically significant alteration in
the rhythmicity of bHLH1b KO mutants
released in free running, with a shortening of
the period, of about 2h compared to Wt. In
contrast to the KO mutants of RITMO1 the
quality of the oscillations is not altered
(chapter II.c), as demonstrated by the fact that

the R.A.E. values are comparable between
mutants and Wt and by the fact that the totality
of the replicates analyzed passed the EPR test
of rhythmicity (Zielinski et al., 2014). The
presence of such differences in the phenotype
of mutants for partially redundant clock
proteins (as RITMO1 and bHLH1b are,
plausibly) is not uncommon. An example is
given by mutants of the MYB proteins CCA1
and LHY in *Arabidopsis thaliana*, both of
which exhibit a shortening of the free-running
period relative to Wt, albeit of different
magnitude (Alabadı et al., 2002; Mizoguchi et
al., 2002). The data accumulated to date do
not allow advanced hypotheses as to why
RITMO1 mutants exhibit a more pronounced
phenotype than KOs for bHLH1b. An
analysis of rhythmic free-run gene expression
profiles in bHLH1 KOs (such as that reported
for RITMO1 mutants in Chapter II.c) is
essential to verify the presence of similar
phenotypes in these lines. This analysis is
currently in planning. One possibility is that
the RITMO1-bHLH1b dimer and the
bHLH1b-bHLH1b homodimer play a
different and not perfectly complementary
role in the regulation of gene transcription in
*P. tricornutum*, as may be the case for the
possible RITMO1-RITMO1 homodimer.
This could happen by a recognition of
different DNA target sites by the different
dimers. Further investigations are needed to
test this hypothesis. A future in-depth analysis
of the interaction partners of RITMO1 and
bHLH1b, e.g. by co-immunoprecipitation
assay (Lin and Lai, 2017), will allow to
elucidate the interactome of these proteins
and to identify new potential components of
the endogenous clock of *P. tricornutum*.
Generation of RITMO1 KO lines
complemented with an exogenous version of
the protein modified with HA and fluorescent
protein tags is currently underway. These lines
will allow the realization of
immunoprecipitation assays with commercial
antibodies directed against the tag associated
with the protein, thus allowing the
identification of RITMO1 interactors. The
same lines will also provide the material for ChIP-Seq analysis for the characterization of DNA binding sites recognized by the protein. Similar complemented lines are also planned for bHLH1b KOs.

The generation of RITMO1 and bHLH1b double mutants is also planned in

order to attest the effect of double obliteration on the rhythmicity of this alga. In case the role is partially redundant, such co-mutation is expected to induce a major disruption of cellular rhythmicity, larger than those observed in single mutants (Alabadı et al., 2002; Mizoguchi et al., 2002).
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L'altezza è profondità, la luce inaccessa è abisso, la chiarezza è tenebra, il magno è parvo, il distinto è confuso, la lite è amicizia, l'individuo è dividuo, l'immenso è atomo.

adapted from Giordano Bruno - Epistole italiane
Chapter III:

bHLH2 is a light-intensity
dependent modulator of
photosynthesis and growth in the
marine diatom *Phaeodactylum*
*tricornutum*

Contributions:

Alessandro Manzotti, Soizic Cheminant-Navarro, Antonio Emidio Fortunato, Andres
Ritter, Alessandra Bellan, Marianne Jaubert, Benjamin Bailleul, Tomas Morosinotto, Jean-Pierre
Bouly, Angela Falciatore

Keywords: photosynthesis, plastid physiology, plastid retrograde signaling, light stress, chlorophyll fluorescence

In this project I designed and realized all the physiological characterization of photosynthetic
parameters in RNAi transgenic lines grown in liquid medium, as well as the analysis of gene
expression and growth.
In a parallel line of research, I led along the characterization of the diatom circadian clock, my PhD work identified bHLH2, another bHLH-PAS putative transcription factor, as a possible modulator of plastid physiology in *P. tricornutum*. This research integrated a project of the EMBRC-France (European Marine Biological Resource Centre) network, which was already in progress when I joined the laboratory for my Ph.D. program. More specifically, the project aimed to identify new diatom strains showing novel traits of interest (e.g., altered photosynthesis and biomass productivity), by using a Transcription Factor (TF)-based genetic engineering strategy. TFs are master regulators of gene expression and excellent candidates both for fundamental research approaches and for applied research. The foundation for this project was a collection of transgenic lines with a deregulated expression of each of the 220 TFs of *P. tricornutum*, which has been generated in the laboratory using the RNA interference strategy (RNAi) by Soizic Cheminant Navarro (Navarro et al., in preparation). Established by the host laboratory in 2009 (De Riso et al., 2009), RNAi has been widely used by the diatom community before the advent of the genome-editing era. Aim of the TF-based genetic engineering strategy was to widely affect genetic regulation of diatoms, likely accelerating the discovery of new regulators and new phenotypes. This approach appeared particularly suitable for *P. tricornutum* considering that forward genetic screens (random insertion or chemical mutagens) are not feasible in this species presenting a diploid genome and lack of sexual cycle.
Abstract

Accounting for more than 40% of marine primary production, diatoms represent one of the
most prominent phytoplankton groups in the Oceans. Despite their ecological relevance, the
knowledge on the regulation of diatom photosynthesis and on the genetic control of its plasticity is
still very limited. To identify the unknown molecular regulators underlying these processes, we
applied a transcription factor genetic engineering approach to the diatom model species
*Phacodactylum tricornutum* and generated a collection of bHLH transcription factor knock-down
transgenic lines via RNA interference. Phenotyping by non-invasive imaging techniques based on
measurements of chlorophyll fluorescence changes lead to the identification of the bHLH-PAS
protein bHLH2 as a candidate regulator of diatom chloroplast activity. Lines bearing a *bHLH2RNAi*
construct show a light intensity-dependent reduction of PSII maximal quantum yield and maximal
thylakoid Electron Transfer Rate. Remarkably, this light-intensity conditional phenotype also
translates in a reduced growth capacity of *P. tricornutum* following transition from low to high light
conditions. Gene expression analysis of *bHLH2* showed a diurnal regulation of its expression, with a
strong, plastid-signals mediated downregulation of *bHLH2* transcripts following dark to light
transition. These results indicate the existence of a strong regulatory cross-talk between plastids and
nucleus for the optimization of diatom growth capacity under changing lights. bHLH2 represents the
first example of bHLH-PAS transcription factor characterized as a light-intensity dependent
modulator of photosynthesis and growth and it represents an ideal entry point for further dissection
of the gene regulatory network controlling these critical responses in diatoms.
Introduction

Although diatoms represent one of
the most abundant taxa of marine microalgae
and a major actor of carbon biogeochemical
cycles at the global level, the regulation of gene
expression involved in their photosynthetic
physiology is still largely uncharacterized. The
proper functioning of photosynthetic
metabolism and its dynamic adaptation to
environmental stimuli require fine-tuning of
gene expression. The complex evolutionary
processes that led to the acquisition of the
plastid by diatoms make the study of this
regulatory process particularly complex. As
secondary endosymbionts of the red plastid
lineage, diatom chloroplasts are distinct from
the green lineage ones and therefore, direct
extrapolation on the regulation of
photosynthetic processes from higher plants
and chlorophytes is not always applicable or
relevant (see Chapter I, paragraphs 3.a and
3.b). In recent years, particular genomic
features and metabolic pathways have been
identified in these algae by genomic analysis.
Initial regulators of light stress mechanisms
have been also identified by targeted
modulation of the expression of specific
proteins such as the LHCXs (Bailleul et al.,
2010; Buck et al., 2019; Giovagnetti et al.,
2022) However, information on the regulation
of diatom photosynthesis and the genetic
control of its plasticity remain still very limited.
As already presented in Chapter I,
approximately 15% of the entire proteome in
diatoms is represented by genes targeted to the
plastid or otherwise derived from the
endosymbionts that gave rise to the plastid
(Moustafa et al., 2009). A large part of these
genes have been transferred into the nuclear
genome of the host, so that only a reduced
fraction of genes remain in the plastid genome
(130 in the case of *P. tricornutum*, Oudot-Le
Secq et al., 2007). The alteration of cellular
processes by means of targeted deregulation of
specific enzymes has already been proven as a
potentially effective system to study diatom
physiology and metabolism (Daboussi et al.,

2014; Levitan et al., 2015). A valid alternative
that may allow a wide-ranging description of
the regulatory mechanisms of plastid
physiology is offered by transcriptional
engineering, *i.e.* the deregulation of
transcription factors (TFs, Bajhaiya et al.,
2017). TFs are master regulators of gene
expression and represent perfect candidates
for obtaining a profound alteration of
processes of interest through a single, pin-
point mutation. Hence TF engineering
represents a powerful strategy to identify novel
regulators of diatom biology (Bajhaiya et al.,
2017; Huang and Daboussi, 2017). In a TF
engineering approach, the modulations of TFs
expression by RNA-mediated interference is a
valid alternative to gene knock-out since a
complete loss of TF gene function could lead
to a highly noxious phenotype in the case of
factors controlling vital processes for the cells.
Differently, a partial deregulation of gene
expression is expected to lead to less
destructive phenotypes but still sufficient to
characterize the physiological alterations. This
kind of strategy is feasible in *P. tricornutum*
since RNAi is an already established
techniques (De Riso et al., 2009; Siaut et al.,
2007). Hence, deregulation of gene
expression via RNAi can allow the relatively
straightforward generation of a multitude of
mutants for different genes and favour a
characterization of their phenotypic
alterations. Following this principle, I
performed a characterization of the
photosynthetic physiology of a set of
transgenic lines presenting an antisense
construct aimed at silencing genes encoding
bHLH proteins. The aim was to evaluate their
possible involvement in the regulation of
plastid activity and, in parallel, further analyze
the proteins involved in the circadian clock of
*P. tricornutum*. To this end, I have
implemented a variety of biophysics
techniques that rely on chlorophyll
fluorescence as a reporter of photosynthetic
<figure><img src="image_113.png" /><figcaption>Figure 1. Principles of chlorophyll fluorescence analysis of PSII activity</figcaption></figure>

A) When chlorophyll absorbs light it gets excited to its singlet-electron excitation state (¹Chl*), which tends to quickly get back to its basal condition in a time span of few ps. To return to its ground energy state chlorophyll can: emit light (as fluorescence), dissipate heat via photoprotective processes or fuel the photosynthetic machinery (photochemistry process). ¹Chl* can also decay to a triplet excitation state (³Chl*), which in turn is able to generate singlet oxygen (¹O₂*). B) Sequence of a typical chlorophyll fluorescence trace. Cells are adapted to dark so to allow full opening of PSII centres and oxidation of their downstream electron acceptors. A measuring light (ML) is switched to measure basal fluorescence (F₀) with no induction of photochemistry. Application of a saturating light pulse (SP) allows measurement of maximum fluorescence of the dark-adapted chlorophyll (Fm). The variable fluorescence (Fv) is equal to Fm-F₀. A light able to drive photosynthesis (AL) is switched on causing a rapid increase in fluorescence. After a period of time another SP is applied, allowing to measure the maximum fluorescence in the light (Fm'). The level of stationary fluorescence when Fm' is measured is named F₀. Maximal fluorescence variation (Fm- Fm') represent the contribution of non-photochemical quenching (NPQ). Adapted from van Kooten 1990.

processes. The following section is devoted to
introducing the principles of these techniques.

Analysis of photosynthesis via
chlorophyll fluorescence

The techniques used for studying
photosynthetic and photoprotective capacity
are based on the analysis of chlorophyll
fluorescence. When excited by a photon, a
special pair of chlorophyll molecules switch to
a singlet-electron excitation state ('Chl*),
which is highly energetic and tends to quickly
get back to its basal condition (in a time span
of a few picoseconds). The relaxation process
can occur in three ways: by transferring the
excited electron to other components of the
photosynthetic machinery (photochemical
quenching), by fluorescence emission or by
heat energy dissipation via the Non

Photochemical Quenching processes (Müller et al., 2001). These three processes are mutually-exclusive, so that an increase in the efficiency in one of the three leads to a decrease in the others (Fig.1). Measuring the fluorescence emitted by chlorophyll can be then a useful strategy to quantify the yield of photosynthesis and heat dissipation (Maxwell and Johnson, 2000; van Kooten and Snel, 1990). At room temperature, the observable fluorescence basically derives only from the chlorophyll special pairs of the Photosystem II, PSII. Under room temperature conditions, in fact, electron-acceptance reactions by molecules downstream of the PSI are relatively constant and occur at high efficiency and speed. Specifically, the de-excitation of the special pair of PSI by electron transfer to
downstream acceptors occurs at a rate greater
than by fluorescence or non-photochemical
dissipation, which are negligible. The analysis
of chlorophyll fluorescence consequently
allows to obtain a picture of the functionality
of the PSII, but also the condition of other
electron transport within the cell and the
global efficiency of the electron transfer and
redox processes downstream of it.

For evaluating the efficiency of PSII to
absorb incoming photons and funnel their
energy into photochemical processes
(maximal PSII quantum yield, or ΦPSIImax),
a short adaptation of cells in the dark (in the
range of a few minutes) is used. This treatment
leads to the full reduction of the non-irradiated
special pairs of PSII, and the oxidation of their
downstream electron acceptors mobile
plastoquinones (PQs), hence setting them in a
state ready for charge separation. PSII
reaction centers which are fully reduced and
ready to cease their electrons to downstream
acceptors are commonly defined “open”.
Dark-adapted PSII are hence irradiated with a
detection light (or ML, measuring light) that is
sufficient to excite basal fluorescence of
chlorophyll (Fo) but too feeble to induce
changes in photochemical electron transfer.
Cells are then subjected to a brief and intense
saturating pulse (SP) of actinic light (i.e. light
capable of exciting photochemistry, AL) that
causes the full oxidation of PSII reaction
centers and the consequent reduction of
electron acceptor PQ. If the SP is short
enough, Non-Photochemical Quenching
processes cannot take place. Also, due to the
slow kinetics of electron transfer from PQs to
downstream acceptors, if the SP is short
enough no photochemistry can occur either
and all the absorbed energy is released by
fluorescence. The saturating pulse therefore
leads to complete “closure” of the PSII pool
and subsequent induction of maximal
fluorescence (Fm). Being Fm the highest
amount of fluorescence of dark adapted PSII,
the difference between Fm and Fo is equal to
the variable fluorescence (Fv). Fv/Fm

represents the proportion of photons that
activate the thylakoid electron transfer chain in
absence of NPQ and that are not released as
basal fluorescence. In other words, Fv/Fm is
equal to the maximal quantum yield of PSII
(ΦPSIImax). ΦPSIImax gives information
about the overall performance of the
photosynthetic apparatus and of the
downstream metabolism consuming ATP and
redox potential produced by it (Maxwell and
Johnson, 2000; van Kooten and Snel, 1990).

When cells are irradiated with
continuous actinic light, the electron acceptors
of PSII are quickly reduced, leading to an
increase of chlorophyll fluorescence (Kautsky
effect, (Kautsky, 1960; Maxwell and Johnson,
2000). Fluorescence then decreases over a
time scale of a few minutes. This progressive
decline in fluorescence is due to the initiation
of electron transfer along the thylakoid chain.
When light pressure is sufficiently high to
grant a saturation of electron transport chain,
this decline is also associated with an increase
in NPQ efficiency, whose magnitude depends
on the actinic light intensity. When the
stationary fluorescence reaches a steady state
(Fs), a SP is applied to calculate the emission
of maximal fluorescence in presence of light
(Fm'). Under this condition, (Fm'-Fs)/Fm'
represents the fraction of energy used for
photosynthesis under illumination, i.e. the
photochemical quantum yield of PSII in the
light, or ΦPSII. The fraction (Fm-Fm')/Fm' is
then equal to the energy dissipated via NPQ
reactions (Müller et al., 2001). It is important
to remember that part of the reduction in
maximal fluorescence observed in light is
attributable to either by energy-dependent
heat dissipation (qE), or inactivation of PSII
centers due to photodegradation of the D1
protein, a phenomenon that represents the
photoinhibitory (qI) component of NPQ.
While the qE component of NPQ reverses
with rapid kinetics once light stress is
interrupted, the replacement of damaged D1
subunits by photoinhibition, which is
necessary for the reversion of the qI
component, requires a time lag of several
minutes to several hours.

Since the parameter ΦPSII represents by definition the efficiency of conversion of light energy in photochemical process, the number of electrons flowing along the thylakoid chain for a given amount of time will be proportional to the product between ΦPSII and the light intensity (I, expressed as a number of photons reaching a given surface for a unit of time). This product is also named relative Electron Transfer Rate, as already shown in chapter II.c.

It is important to stress that the chlorophyll fluorescence parameters described above refer only to PSII. As this complex is only the first of the thylakoid chain, an alteration in the activity of any of the downstream electron transport components can impact its physiology and alter its quantum yield. A slowdown in the activity of one of the plastid's redox players (e.g. due to a mutation or reduction in expression) will lead to an accumulation of electrons upstream of it, an

over-reduction of the complexes preceding it and, consequently, a decrease in the quantum yield of PSII. Therefore, analysis of chlorophyll fluorescence at ambient temperature is a powerful tool for obtaining information about the overall activity of photosynthesis, but it does not allow for the precise localization of any bottlenecks.

In the case of an altered quantum yield of PSII, several strategies can be applied to better localize the perturbation. One of these consists in quantifying the oxidoreductive state of PSI. If the bottleneck in the transfer of electrons occurs upstream of PSI (e.g. at the level of cytochrome b6f), at the same light intensity the amount of electrons that will reach the PSI per unit time will be lower and therefore PSI itself will be over-oxidized compared to normal (Fig.2A). Conversely, if the defect is located downstream, the PSI will give up less quickly than normal the electrons it receives and will be consequently over-reduced. This is the case in the presence of perturbations of the Calvin Benson Bassham Cycle, the first consumer of

<figure><img src="image_114.png" /><figcaption>Figure 2. Use of P700 redox state to localize perturbations in the plastid electron transfer chain</figcaption></figure>

Graphical representations of two possible scenarios of altered plastid electron transfer chain. A) depicts a situation where the bottleneck in the transfer of electron is placed downstream of PSI, specifically at the level of the CBBC. Due to the lowered capacity of the CBBC to consume the redox potential generated by PSI (NADPH), the photosystem will be limited on its electron acceptor side resulting in an over-reduction. B) depicts a situation where the bottleneck in the transfer of electron is placed upstream of PSI, in this example at the level of the cytochrome b6f. The impaired income of electron to the donor side of PSI will result in an over oxidation of the photosystem. The relative level of reduction of PSI can be determined exploiting its absorbance at 705nm, dependent on the redox state, hence allowing a finer localisation of electron transfer defects.
the reductive potential generated by the
thylakoid (Fig.2B). To evaluate the redox state
of PSI, it is possible to exploit the spectral
properties of this complex, in particular its
variable absorbance at 700 nm, absorbance
that increases with increasing reduction level.

The possibility of large-scale
characterization of the photosynthetic capacity
of a high number of cell lines is a useful tool
for the identification of regulators of plastid
activity. Such a screening strategy has been
developed on a variety of unicellular algae
(Niyogi et al., 1997; Ozaki et al., 2007) and,
recently, also on the *Eustigmatophyceae*
(*Stramenopila*) *Nannochloropsis gaditana*
(Perin et al., 2015). Large-scale screenings of
this kind can be applied for a forward genetics
strategy, in order to identify mutants of
photosynthetic capacity generated by random
mutagenesis and subsequently determine the
gene associated with these alterations. Equally,
large-scale screenings can be also applied to a
reverse genetics strategy, for testing whether
one or more genes of interest, obliterated by
targeted mutagenesis, influence plastid activity.
In addition, such analysis allows a
measurement of PSII activity in a non-invasive
way. As seen, the chlorophyll fluorescence of
PSII provides an overview of the global activity

of the electron transfer chain and the
metabolic processes that depend on them
downstream. Consequently, its analysis is an
extremely useful tool for a preliminary
identification of lines mutated in genes
involved in any of the various photosynthesis-
dependent physiological processes.

As reported below, through the
application of a set of these biophysical
techniques I was able to pursue a
characterization of alterations in
photosynthetic physiology in cell lines
containing a bHLH-PAS bHLH2 protein
silencing construct. Although not yet
conclusive due to the difficulty of obtaining a
stable and significant reduction of bHLH2
expression in these lines, these analyses seem
to suggest that this protein may play a role in
the regulation of plastid activity in *P. tricornutum*. If the results are confirmed on
KO lines for bHLH2 (analyses are currently
in progress), this would be a first
characterization of a bHLH-PAS protein as a
modulator of photosynthesis. Given the
absence of proteins with a similar architecture
in *Archaeplastida*, this could open new
perspectives on the study of regulation of
plastid activity in secondary endosymbionts.
# Methods

## Generation of bHLH RNAi lines

The selection of genes encoding for putative bHLH family transcription factors was performed following the initial prediction reported in (Rayko et al., 2010) and re-annotation on the new version of *P. tricornutum* genome (“Phaeodactylum_tricornutum - Ensembl Genomes 52,”). The list of the seven identified genes is shown in Table 1 along with their associated Phatr3 accession numbers. Transgenic and control lines of *P. tricornutum* used for bHLH2 have a Pt1 8.6; CCMP2561 genomic background. Lines targeted for gene silencing were generated following the protocol reported in (De Riso et al., 2009). Diatom cells were transformed by biolistic

bombardment (Falciatore et al., 1999) with a linear construct containing an anti-sense fragment of the target genes transcriptionally fused downstream of a *Sh ble* cassette conferring resistance to phleomycin (pKS-Sh ble-FA vector, (De Riso et al., 2009; Siaut et al., 2007)). Anti-sense fragments targeting sequences (~200bp) specific to each bHLH (coding sequence or transcribed regulatory UTR regions) were generated via PCR using the primers listed in Table 1 and cloned in the pKS-Sh ble-FA vector via EcoRI and StuI digestion. The expression of the *Sh ble* - antisense sequence construct was regulated via the constitutive *H4* promoter and the *FcpA* terminator. After transformation, transgenic lines expressing the antisense fragments were selected and maintained on F/2 media (Guillard, 1975) added with 1% agarose, in the presence of the selective phleomycin antibiotic.

<table><caption>Table. 1 Primers used for the construction of the RNAi cassettes</caption><thead><tr><th>Target gene</th><th>Accession</th><th>Primer name</th><th>5' to 3' sequence</th><th>Pairing length</th></tr></thead><tbody><tr><td rowspan="2">bHLH1a (RITMO1)</td><td rowspan="2">Phatr3_J44962</td><td>bHLH1a_3</td><td>TCCCAGCCAAAGTCGAGGTAGCTCGCCTTTGCTCAATATTCAACCATC</td><td>23</td></tr><tr><td>bHLH1a_4</td><td>ACGGCGGCCCACGGGTCCGAGGCCTGAAGTGTACAGCCACGTTGGTA</td><td>22</td></tr><tr><td rowspan="2">bHLH1b</td><td rowspan="2">Phatr3_J44963</td><td>bHLH1b_3</td><td>TCCCAGCCAAAGTCGAGGTAGCTCGCACGTGAAGAGCTAATGAATGC</td><td>22</td></tr><tr><td>bHLH1b_4</td><td>ACGGCGGCCCACGGGTCCGAGGCCTTGGAGTTGCTTTTTTGATTCTT</td><td>22</td></tr><tr><td rowspan="2">bHLH2</td><td rowspan="2">Phatr3_J54435</td><td>bHLH2_3</td><td>TCCCAGCCAAAGTCGAGGTAGCTCGGTTCAGCCAGCTATCGAGCTAT</td><td>22</td></tr><tr><td>bHLH2_4</td><td>ACGGCGGCCACGGGTCCGAGGCCTAATATCAGTACGGTCCCCCATT</td><td>22</td></tr><tr><td rowspan="2">bHLH3</td><td rowspan="2">Phatr3_J42586</td><td>bHLH3_3</td><td>TCCCAGCCAAAGTCGAGGTAGCTCGATGAGACAAGCCACAACAATGATAG</td><td>25</td></tr><tr><td>bHLH3_4</td><td>ACGGCGGCCCACGGGTCCGAGGCCTGCTAGTGATTTAGTGCGACTGTCTC</td><td>25</td></tr><tr><td rowspan="2">bHLH4</td><td rowspan="2">Phatr3_J31578</td><td>bHLH4_3</td><td>TCCCAGCCAAAGTCGAGGTAGCTCGACCATTTTCGGGAGGTACAAT</td><td>22</td></tr><tr><td>bHLH4_4</td><td>ACGGCGGCCACGGGTCCGAGGCCTCCGAATAATATGGAACGGAATG</td><td>22</td></tr><tr><td rowspan="2">bHLH5</td><td rowspan="2">Phatr3_J43365</td><td>bHLH5_3</td><td>TCCCAGCCAAAGTCGAGGTAGCTCGATGGGTCGATTTCTCAATGAAC</td><td>22</td></tr><tr><td>bHLH5_4</td><td>ACGGCGGCCCACGGGTCCGAGGCCTGTTGGTGATACGCACTTTCTGA</td><td>22</td></tr><tr><td rowspan="2">bHLH6</td><td rowspan="2">Phatr3_J48817</td><td>bHLH6_3</td><td>TCCCAGCCAAAGTCGAGGTAGCTCGACTTTCCGTCGAAGAGATTCAG</td><td>22</td></tr><tr><td>bHLH6_4</td><td>ACGGCGGCCACGGGTCCGAGGCCTGTGACTGTGCGTGAGGTAGTGT</td><td>22</td></tr></tbody></table>

The table reports the primers used for amplifying the anti-sense sequences used for targeting *bHLH* genes following de Riso et al., 2009. Common names and Phatr3 accessions are reported for each targeted gene. Red and green letters highlight the common sequences of forward and reverse primers respectively used for cloning in the pKS-Sh ble-FA vector following de Riso et al., 2009. Black letters refer to the target specific sequence of the primers.

## On plate colony screening

Pre-screening of lines containing a construct for RNAi of each bHLH gene was performed by adapting to *P. tricornutum* the procedure developed on the alga *Nannochloropsis gaditana* by (Perin et al., 2015). Briefly, lines transformed with RNAi constructs and exhibiting phleomycin

resistance were grown in liquid culture and maintained under exponential growth phase. Following optical density analysis at 750nm, aliquots of the different cultures were diluted to equalize their optical density to OD=0.2 and equal volumes were spotted onto plates of F/2 (Guillard, 1975) supplemented with 1% agarose. Colonies were subsequently grown under continuous light at an irradiation intensity of 30 µmol photons m⁻² s⁻¹. The effect
of insertion of transformation cassettes was analyzed by taking into account two parameters. First, colony growth capacity was calculated by checking the change in basal chlorophyll fluorescence in cells adapted for 20 minutes in the dark (Fo) and normalized for colony surface area. The variation of Fo/surface was followed over five days from the time of spotting on plate. Second, the maximum quantum yield of Photosystem II (PSII) was calculated by testing the ratio of the difference between chlorophyll fluorescence following exposure to a light-saturating pulse (Fm) minus the basal fluorescence of dark-adapted cells (Fo) divided by Fm (Maxwell and Johnson, 2000). This parameter allows us to obtain a rapid estimate of the maximum photosynthetic capacity over a large number of colonies. *In vivo* fluorescence analysis on colonies was performed using a FluorCam FC 800 video-imaging apparatus (Photon Systems Instruments), which allows a normalization of fluorescence signal over the surface of the colony.

Liquid culture and irradiation conditions

Cells were grown in batch culture in F/2 Guillard media. Liquid cultures were grown and maintained at 18°C in a Standard Growth Chamber SGC120 (Weiss Gallenkamp™). Illumination was performed with white light Master TL90 36W/950 neon lamps (Philips™). Experiments were performed in constant light, if not specified otherwise. Cell concentrations were measured using a Z2 Coulter Particle Count and Size Analyser (Beckman Coulter). Minimal and maximal size thresholds were set to 3 µm and 11 µm. Growth rates were calculated in exponential phase of growth over four days and expressed as the average number of divisions per day over the experiment. The formula used is division per day = (ln(c₄) - ln(c₀)) / (4 * ln(2)), where c₀ and c₁ represent the culture concentrations at day 0

and day 4. For experiments involving treatment with 3-(3,4-dichlorophenyl)-1,1-dimethylurea (DCMU), cultures in exponential growth phase were dark adapted for 50 hours and transferred to 30 µmol photons m⁻² s⁻¹ for 4 hours in presence of a 1000X dilution of DCMU ethanol solution (final concentration of 2µM). DCMU was added 10 minutes before the transfer to light in order to ensure its full penetration in the cells. Control replicates were treated with an equal amount of pure ethanol to exclude solvent dependent effects on photosynthesis.

ΦPSII analysis via chlorophyll fluorescence in liquid cultures

All fluorescence analyses were performed on cultures grown in liquid F/2 media and sampled during exponential growth. To measure PSII maximal quantum yield, 10ml of biological replicate cultures (~10⁶ cells ml⁻¹) were centrifuged at 2500 rpm for 15 minutes to be concentrated ten times in their own supernatant. Concentrated cells were adapted to dim light for 20 minutes to induce complete relaxation of PSII reaction centers. Fluorescence analysis was conducted using a Paradigme Camera Speedzen system (Beambio). Measuring light and actinic saturating flash were provided by blue and green LEDs, respectively. Saturating flash corresponded to 250ms of 2,000 µmol photons m⁻² s⁻¹. Chlorophyll fluorescence was detected through a low-pass filter at wavelength >700 nm. Experiments demanding the analysis of ΦPSII upon light irradiation were performed with an in-house spectrofluorometer allowing irradiation of the samples in the analysis chamber and using green LEDs for both detection and saturating pulses. qE calculations presented in Fig.10C were realized considering cells at the end of the relaxation kinetics as completely relaxed (qE=0). qE was therefore calculated as equal to (Fm'-Fm)/Fm, where Fm' is the maximal
fluorescence induced by the last saturating pulse.

## relative Electron Transfer Rate

Three biological replicates were adapted to continuous light (30 µmol photons m⁻² s⁻¹ or 200 µmol photons m⁻² s⁻¹). Analysis was performed on cultures in the exponential growth phase (1 to 2x10⁶ cells ml⁻¹). Concentrated cultures were left under dim light in order to allow full relaxation of PSII before analysis. Samples were then exposed to increasing intensity steps of actinic light to obtain the saturation light curves. Each light step lasted at least five minutes until full adaptation of the photosynthetic apparatus to the irradiation level. Illumination was performed with green LED lights. At the end of each light step, chlorophyll fluorescence was measured with a blue light measuring pulse and a 250 ms green light saturating pulse was used to induce PSII maximal fluorescence in the light (Fm'). Fluorescence of light adapted PSII (Fs) was measured just prior to saturating light application. PSII quantum yield ΦPSII was calculated as (Fm'-Fs)/Fm' (Maxwell and Johnson, 2000). For each light step, the relative Electron Transfer Rate through the photosynthetic chain (rETR) was calculated as ΦPSII x Light intensity. The light saturation curve resulting from plotting rETR as a function of light intensity was fitted to a double exponential decay function following (Platt et al., 1980; Ralph and Gademann, 2005). Fitting was performed via Origin software version 9.1 to the function

$$ETR = ETR_{max} * (1 - e^{(-\alpha E/ETR_{max})}) * (e^{(-\beta E/ETR_{max})})$$

## Evaluation of PSI redox state

The redox state of the PSI donor was calculated as the difference between the absorption kinetic at 705 nm similarly to (Long et al., 2021). Analysis was performed on cells adapted to continuous 5 µmol photons m⁻² s⁻¹ for two weeks and after one day of exposure

to 200 µmol photons m⁻² s⁻¹. Properties of PSI were calculated from the measurements of the absorption changes in the dark and in the light-adapted condition. PSI redox state was estimated upon increasing light intensity steps to which samples were adapted for 20 minutes to assure stationary activity of photosynthetic chain. Absorption changes at 735 nm were also calculated to eliminate spectral contributions due to diffusion and other thylakoid complexes. As a control of the maximal possible oxidation state of PSI, cells were treated with a saturating dose of DCMU and subject to a saturating pulse of light to obtain the Pmax value. All data were normalized to Pmax - P0 (full oxidation - maximal reduction states). Analysis was performed on a Joliot-type spectrophotometer (JTS-10, Biologic, Grenoble, France).

## Lincomycin treatment

Photoinhibition analysis via lincomycin treatment was performed following Lavaud et al., 2016. Cultures were preadapted in continuous light at 5 µmol photons m⁻² s⁻¹ for two weeks. Three biological replicates were treated with a saturating dose of inhibitor (500 µg ml⁻¹) and left 10 minutes under low light (<<5 µmol photons m⁻² s⁻¹) to allow full penetration of the compound. Cultures were subsequently exposed to 500 µmol photons m⁻² s⁻¹ for sixty minutes to induce photoinhibition of D1 protein and then moved to 10 µmol photons m⁻² s⁻¹ to allow relaxation of the residual non photodamaged PSII centers. Cultures aliquots were sampled periodically to determine ΦPSII using the in-house spectrofluorometer.

## RNA extraction and RT-qPCR analysis

For RNA purification, 10⁸ cells from exponential growth cultures (~10⁶ cells ml⁻¹) were filtered onto Whatman filter 589/2. Filters were then washed with 25mL of PBS
1X, cut in two to have technical replicates, flash frozen in liquid nitrogen and stored at -80°C. RNA was extracted with TriPure Isolation Reagent (Roche) following the protocol provided by the company and further purified by an ammonium acetate precipitation. RNA purity and integrity were checked on samples by the 260nm/230nm and 260nm/280nm OD ratios and by gel electrophoresis. 500ng of RNA were retro-transcribed via an oligo-dT and random primers mix using the QuantiTect Reverse Transcription Kit (Quiagen). For RT-qPCR,

ribosomal protein small subunit 30S (RPS) and TATA box Binding Protein (TBP) were used as reference genes (Siaut et al., 2007). For each sample, geometric mean of Ct was used as reference to evaluate the expression of genes of interest via the ΔΔCt Livak method (Livak and Shcmittgen, 2001). Expression data was normalized against the average Wt expression value. All RT-qPCR analyses were performed on two biological replicates obtained in distinct experiments. TBP, RPS and bHLH2 primers used for RT-qPCR analysis are listed in Table 2.

<table><caption>Table 2. List of the oligonucleotides used in this work.</caption><thead><tr><th>Target gene</th><th>Accession</th><th>Primer name</th><th>Strand</th><th>Sequence</th><th>Type</th></tr></thead><tbody><tr><td rowspan="2">TBP</td><td rowspan="2">Phatr3_J10199</td><td>TBP_Fw</td><td>Fw</td><td>ACCGGAGTCAAGAGCACACAC</td><td>RT-qPCR</td></tr><tr><td>TBP_Rv</td><td>Rv</td><td>CGGAATGCGCGTATACCAGT</td><td>RT-qPCR</td></tr><tr><td rowspan="2">RPS</td><td rowspan="2">Phatr3_J10847</td><td>RPS_Fw</td><td>Fw</td><td>CGAAGTCAACCAGGAAACCAA</td><td>RT-qPCR</td></tr><tr><td>RPS_Rv</td><td>Rv</td><td>GTGCAAGAGACCGGACATACC</td><td>RT-qPCR</td></tr><tr><td rowspan="2">RITMO1</td><td rowspan="2">Phatr3_J44962</td><td>RITMO1_total_Fw</td><td>Fw</td><td>ATTCTTGGTCCCACCCGGTA</td><td>RT-qPCR</td></tr><tr><td>RITMO1_total_Rv</td><td>Rv</td><td>ACGCCACATTGAAAAACCGAG</td><td>RT-qPCR</td></tr><tr><td rowspan="3">bHLH2</td><td rowspan="3">Phatr3_J54435</td><td>bHLH2_Fw</td><td>Fw</td><td>GCAAGGGAAGGGCTCAAAG</td><td>RT-qPCR</td></tr><tr><td>bHLH2_Rv</td><td>Rv</td><td>GCTGCTGGAGAATGCGAATG</td><td>RT-qPCR</td></tr><tr><td>bHLH2_Seq</td><td>Fw</td><td>CAAGGCGCTCATCAGTTCTC</td><td>sequencing</td></tr></tbody></table>

Target gene, name, pairing strand, sequence and use of the oligonucleotides used in this work.

## Subcellular localization prediction

bHLH2 protein sequence was scanned with multiple online tools for subcellular localization prediction. HECTAR (Gschloessl et al., 2008) and ASAFind (Gruber et al., 2015) tools were used to search for the heterokont specific signal peptide and ASAFAP plastid localisation sequences (Kilian and Kroth, 2004). DeepLoc (Almagro Armenteros et al., 2017), NucPred (Brameier et al., 2007), WoLF PSORT (Horton et al., 2007), LOCALISER (Sperschneider et al., 2017), SeqNLS (Lin and Hu, 2013) and NLStradamus (Nguyen Ba et al., 2009) were used for additional subcellular localisation

prediction and to search for Nuclear Localization amino acid Sequence (NLS).

## Generation of bHLH2 Knock-Out (KO) strains

bHLH2 KO mutants were generated by biolistic delivery of the CRISPR/Cas9 ribonucleoproteins according to (Serif et al., 2018). Selection of transformed lines with the riboprotein complex was accomplished by co-targeting the APT gene (Phatr3_J6834) to induce resistance to 2-fluoroadenine (2-FA). Wt cells were co-transformed with a total of 4 sgRNA, two for each gene (Table 3). 2-FA resistant lines were screened via Sanger sequencing (primer reported in Table 2).

<table><caption>Table 3. gRNAs used for *bHLH2* KOs generation</caption><thead><tr><th>Target gene</th><th>Accession</th><th>gRNA name</th><th>Strand</th><th>Sequence</th><th>3' PAM</th></tr></thead><tbody><tr><td rowspan="2">APT</td><td rowspan="2">Phatr3_J6834</td><td>APTg1</td><td>Rv</td><td>AAGCGTGGAATGCCTTTGAA</td><td>GGG</td></tr><tr><td>APTg3</td><td>Fw</td><td>CCTGGGTCCACCAATTGCCC</td><td>TGG</td></tr><tr><td rowspan="2">bHLH2</td><td rowspan="2">Phatr3_J54435</td><td>Guide_bHLH2_59</td><td>Rv</td><td>TACGTTAAGCGTAAGTGTGT</td><td>CGG</td></tr><tr><td>Guide_bHLH2_135</td><td>Fw</td><td>GATAGCGGGAGCTCGAACGG</td><td>CGG</td></tr></tbody></table>

List of the sgRNA used for CRISPR-Cas9 mutagenesis. Target gene, sgRNA name, target strand, gRNA sequence and the associated protospacer adjacent motif (PAM) are reported.
# Results and Discussion

## Generation of RNAi transgenic lines
targeting bHLH genes

Perin et al., 2015, recently developed a protocol for screening photosynthesis mutants based on chlorophyll fluorescence in the *eustigmatophyceae Nannochloropsis gaditana*. This protocol represents the first application of such screening on algae of the *Stramenopila* group. The screening was performed on cultures grown on solid medium, with the advantage of allowing the analysis of several strains in parallel in a more compact set up than would be provided by an analysis of cultures in liquid. In order to generate colonies with a sufficiently large surface (and hence a sufficiently large signal) to allow fluorescence analysis, small amounts of liquid culture were spotted on agar plates. The possibility to characterize multiple cell lines at once also allows a direct comparison of chlorophyll fluorescence in the same experiment, hence uniforming results. To date, such a screening strategy has not yet been proposed for the study of diatoms. To test the applicability of this screening strategy to *P. tricornutum*, model used for the study of photosynthetic processes in diatoms, a characterization of a collection of transgenic lines containing a construct for RNA interference of the bHLH transcription factors of this alga was performed. The choice was motivated by the number of transcription factors belonging to this family (seven in total, shown in Table 1), thus allowing a preliminary evaluation of the screening method on a reduced number of target genes. A total of 159 phleomycin-resistant transgenic lines containing *bHLH* gene-specific RNAi constructs were generated, distributed among

the various bHLH factors as shown in Table 2. For each bHLH gene, one single RNAi antisense construct was used. The on-plate characterization of these lines was performed in collaboration with the laboratory of Plant Biology at the University of Padua, Italy where the screening strategy was originally developed on *N. gaditana* (Perin et al., 2015). Fig.3 shows an example of the fluorescence profiles of Wt lines and RNAi lines. The results of chlorophyll fluorescence analysis for the study of growth capacity and maximum PSII quantum yield of the mutant lines grown on agar plates are summarized in Table 4. The on-plate analysis was performed on plates without antibiotic. This exploratory plate screening revealed a relevant number of clones (between 10 and 20%) exhibiting alteration in growth or photosynthetic capacity compared with Wt lines for four of the bHLH genes. These included the three bHLH-PAS genes among which *bHLH2* presented the highest percentage of altered lines at the level of on plate growth (20%, equal to 5 lines out of the 25 present). Cultures on agar does not represent the natural condition for *P. tricornutum* growth and consequently the physiological alterations observed may be independent of transformation with RNAi constructs. Therefore, a similar analysis of

<figure><img src="image_115.png" /><figcaption>Figure 3. Screening on plate based on chlorophyll fluorescence</figcaption></figure>

Representative FISH fluorescence results for *Wt* (top) and RNAi strains (bottom), spotted on plate at the same cellular concentration and analysed after 5 days of growth. Fluorescence signals are detected with a FluorCam FC 800 video-imaging apparatus (Photon Systems Instruments, Brno, Czech Republic).
<table><caption>Table 4. Summary results of on plate screening.</caption><thead><tr><th></th><th></th><th>Number of clones</th><th>Clones affected in growth</th><th>Clones affected in Fv/Fm</th><th>% of clones affected</th></tr></thead><tbody><tr><td>bHLH1a</td><td>Phatr3_J44962</td><td>22</td><td>2</td><td>0</td><td>9</td></tr><tr><td>bHLH1b</td><td>Phatr3_J44963</td><td>36</td><td>3</td><td>2</td><td>14</td></tr><tr><td>bHLH2</td><td>Phatr3_J54435</td><td>25</td><td>5</td><td>0</td><td>20</td></tr><tr><td>bHLH3</td><td>Phatr3_J42586</td><td>37</td><td>1</td><td>0</td><td>3</td></tr><tr><td>bHLH4</td><td>Phatr3_J31578</td><td>7</td><td>0</td><td>0</td><td>0</td></tr><tr><td>bHLH5</td><td>Phatr3_J43365</td><td>1</td><td>0</td><td>0</td><td>0</td></tr><tr><td>bHLH6</td><td>Phatr3_J48817</td><td>31</td><td>2</td><td>2</td><td>13</td></tr></tbody></table>

The table reports the number of *P. tricornutum* phleomycin resistant clones obtained after transformation with RNAi constructs for each of the *bHLH* genes. A total of 159 resistant colonies were analysed. The number of colonies showing a significant reduction of growth (calculated as the variation of Fo/surface over five days) and of PSII maximal quantum yield Fv/Fm is reported, as well as the percentage of lines showing a phenotypic alteration for each gene.

maximum photosynthetic capacity on agar-grown cultures was performed on 208 independent transformation control lines containing only phleomycin resistance. As presented in Fig.4, these lines exhibit high scatter in Fv/Fm values, suggesting that the level of noise associated with chlorophyll fluorescence measurements on *P. tricornutum* cultures grown on solid medium makes identification of mutant lines difficult. Given that on-plate analysis were performed in absence of antibiotics, such a level of noise is not due to photosynthetic parameters but is rather a result of the high sensitivity of the on-plate analysis.

To verify whether similar alterations are also found under optimal growth conditions for *P. tricornutum* and to eliminate false positives, subsequent characterization of photosynthetic parameters was repeated on a limited set of independent RNAi lines grown in liquid medium. Given the interest in the laboratory in bHLH-PAS family proteins, these additional investigations were conducted on transgenic lines containing constructs for RITMO1, bHLH1b, and bHLH2 RNAi.

*bHLH2* RNAi lines are affected in photosynthetic capacity in a light intensity dependent way

Characterization of the photosynthetic capacity of lines containing antisense constructs for bHLH-PAS proteins was performed in cells grown under continuous light, in continuity with the growth conditions used for plate colonies. In order to increase the detectability of any alteration of the photosynthetic capacity in the transgenic lines, the irradiation intensity was in this case increased to 200 µmol photons m⁻² s⁻¹. This light intensity is not to be considered stressful for Wt cells of *P. tricornutum* with unaltered

<figure><img src="image_116.png" /><figcaption>Figure 4. On plate screening of phleomycin resistant control</figcaption></figure>

Fv/Fm of 208 independent phleomycin resistant control lines grown on plate. Box plot reports the first, second and third quartile. Whiskers represent the 10th and 90th percentiles. Black dots report the values of cultures above and below 10th and 90th percentiles. Mean and s.d. values for the 208 samples are equal to 0.634 and 0.130 respectively.
non-photochemical quenching (Lavaud and Goss, 2014; Lepetit et al., 2013; Taddei et al., 2016). However, this intensity is strong enough to saturate photosynthetic activity and would hence sensitize for eventual defects. A transgenic line transformed only with the phleomycin resistance cassette and hereafter named Resistance Control RC1 was used to exclude possible effects of the biolistic transformation and antibiotic resistance on photosynthetic physiology. Maximum quantum yield of PSII, Fv/Fm, was used as an indicator of the overall photosynthetic capacity of the lines under study. This analysis revealed that none of the lines containing the antisense construct targeting for *RITMO1* or for *bHLH1b* present a significant reduction in Fv/Fm compared with the Wt line or the control for phleomycin resistance (Fig.5A). Analysis of a subset of *RITMO1* RNAi lines reveals a reduction in the expression of this gene, suggesting that deregulation of its transcription levels does not bring alterations in maximal photosynthetic activity (Fig.5B). In contrast, three of the four lines presenting the antisense construct targeted against *bHLH2* (KD2.1, KD2.3, KD2.4) showed a significant reduction in the maximal quantum yield of PSII, ranging from 14% to 21% reduction compared with control lines (Fig.5A).

An alteration of photosynthetic efficiency as the one described can be expected to impact cellular growth and,

<figure><img src="image_117.png" /><figcaption>B</figcaption></figure>

<figure><img src="image_118.png" /><figcaption>C</figcaption></figure>

<figure><img src="image_119.png" /><figcaption>Figure 5. bHLH2 lines are affected in photosynthesis and growth</figcaption></figure>

A) Wt, RC1 cells and transgenic lines containing RNAi constructs targeting the three different bHLH-PAS genes of *P. tricornutum* were grown under continuous 200 µmol photons m⁻² s⁻¹ to test possible alterations of PSII maximal quantum yield (Fv/Fm). KD1a: RNAi lines for *RITMO1*; KD1b: RNAi lines for *bHLH1b*; KD2: RNAi lines for *bHLH2* (n≥3, mean ±s.e.m., P-values obtained via One-way analysis of variance against Wt with Bonferroni post-test correction are shown as ***P<0.001). B) Expression of *RITMO1* KD1a.5 and KD1.12 RNAi lines(n=2, mean ± s.e.m.). Cells were grown under 12L:12D at 30 µmol photons m⁻² s⁻¹ and collected at ZT7. Expression values are normalized to the mean expression of Wt strain. C) Effect of *bHLH2* RNAi construct on growth under 200 µmol photons m⁻² s⁻¹ Growth capacity of Wt, RC1 and *bHLH2* RNAi lines grown under continuous 200 µmol photons m⁻² s⁻¹. Growth capacity was calculated on exponential growth cultures and is expressed as average number of divisions per day over four days. (n=3, mean ±s.e.m., P-values obtained via One-way analysis of variance against Wt with Bonferroni post-test correctionare shown as ***P<0.001)
<caption>Figure 6. Effects of light intensity on Fv/Fm and growth in *bHLH2* RNAi lines</caption>

Wt, RC1, KD2.1 KD2.3 and KD2.4 lines were grown in liquid media under continuous 5 µmol photons m⁻² s⁻¹ (A), 30 µmol photons m⁻² s⁻¹ (B) or 200 µmol photons m⁻² s⁻¹ (C) to test PSII maximal quantum yield (Fv/Fm). n≥10, mean ±s.e.m., P-values obtained via One-way analysis of variance against Wt with Bonferroni post-test correction are shown as **P < 0.01, ***P<0.001). D) Growth capacity of Wt, RC1, KD2.1 KD2.3 and KD2.4 RNAi lines grown under continuous the light conditions of panels A-C. Growth capacity was calculated on exponential growth cultures and is expressed as average number of divisions per day over four days (n=3, mean ±s.e.m.).

consequently, the growth capacity of these
RNAi cultures. In order to verify this
hypothesis, the growth rate of Wt, RC1
control cells and bHLH2 RNAi lines adapted
to continuous light 200 µmol photons m⁻² s⁻¹
was determined. In a manner comparable to
that shown at the maximal quantum yield level
of PSII, the KD2.1 KD2.3 and KD2.4 lines
are characterized by a significant reduction in
growth capacity under these conditions, thus
confirming the expected correlation between
the reduction of photosynthetic efficiency and
growth capacity (Fig.5C). Given the results
above, we hence decided to characterize the
photosynthetic alterations of the KD2.1
KD2.3 and KD2.4 lines at higher light
intensity (400 µmol photons m⁻² s⁻¹) assuming
to potentially accentuate the observed
phenotypes by imposing a stronger pressure
on the thylakoid apparatus. Notably, when
transferred to these irradiation conditions, the
three transgenic lines showed a total growth
arrest, thus making any characterization of
photosynthetic capacity impossible.

Such a result suggests that the
observed phenotype can be light intensity
dependent, with an increase of perturbation of
photosynthetic capacity under increasing
photon fluency. To test this hypothesis and
given the effects observed at 400 µmol photons
m² s⁻¹, it was decided to test whether the
phenotype of the KD2.1 KD2.3 and KD2.4
lines is also visible at lower irradiations,
specifically at 5 and 30 µmol photons m⁻² s⁻¹. At
both intensities, growth capacity and maximal
quantum yield of PSII were tested. *bHLH2*
RNAi lines KD2.1 KD2.3 and KD2.4 showed
a minor reduction in maximal PSII quantum
yield when grown under low light in

<figure><img src="image_120.png" /></figure>

<figure><img src="image_121.png" /></figure>

<figure><img src="image_122.png" /></figure>

<figure><img src="image_123.png" /></figure>
comparison to the control lines, but these
differences became progressively more
pronounced when cells were adapted to higher
light irradiation (30 and 200 µmol photons s⁻¹
m² Fig.,6A-C) confirming the light intensity
dependent alteration in maximal yield of PSII.

In parallel, growth analysis highlighted
that although control lines show a rise in
growth speed as light irradiation (and
consequently the energy available to power
cell metabolism) increases, *bHLH2* RNAi
manifest a saturation of growth rate capacity
under low light irradiation (Fig.6D) and do not
show a similar increase of growth capacity at
higher light fluencies. Indeed, Wild-type and
RC1 cells went from one and a half division
per day at 5 µmol photons s⁻¹ m² to two and a
half divisions per day at 200 µmol photons s⁻¹
m² (Fig.6D), while mutants impaired to
increase their growth rate in response to light
intensity and stayed at one and a half division
per day even under higher light intensity.

Effects of antisense RNAi on
*bHLH2* transcription

In order to verify the effects of the
antisense RNAi construct on the transcription
of *bHLH2*, an analysis of the expression of
this gene was performed on the KD2.1 KD2.3
and KD2.4 lines grown at the three light
intensities of the physiological characterization
of photosynthesis in the previous paragraph.
As shown in (Fig.7A-C) the expression of
*bHLH2* was not significantly altered in any of
the three KD lines at 5 µmol photons m⁻² s¹ and
30 µmol photons m⁻² s¹. At 200 µmol photons
m⁻² s¹, lines KD2.1 and KD2.3 showed a slight
but not statistically significant reduction in
*bHLH2* transcript content, whereas only
KD2.4 manifested a major and significant

reduction (approximately 50% compared to
Wt). Overall, this data suggests that although
the RNAi construct may lead to a perturbation
of the *bHLH2* expression, especially at higher
light irradiance under study, no correlation
between the phenotypes observed at the level
of photosynthetic activity and this deregulation
could be globally identified. It is important to
note that it has been suggested the existence of
two silencing mechanisms in *P. tricornutum*,
one based on repression of transcription and

<figure><img src="image_124.png" /><figcaption>**Figure 7. Gene expression of bHLH2 under the different continuous light irradiations**

Expression of *bHLH2* in Wt, RC1, KD2.1 KD2.3 and KD2.4 RNAi lines grown under continuous light at 5 µmol photons m⁻² s⁻¹ (A n=3, mean ± s.e.m.), 30 µmol photons m⁻² s⁻¹ (B n=4, mean ± s.e.m.) or 200 µmol photons m⁻² s⁻¹ (C, n=3, mean ± s.e.m.). Expression values are normalized to the mean expression of Wt strain. P-values obtained via One-way analysis of variance against Wt with Bonferroni post-test correction are shown as ***P<0.001).</figcaption></figure>
a second based on translational inhibition (De Riso et al., 2009). A reduction in the content of the target protein can therefore occur where there is no visible reduction in the content of the mRNA encoding it (De Riso et al., 2009). On this basis, an in-depth photosynthetic characterization of the KD2.1 KD2.3 and KD2.4 lines was carried out to dissect in detail their alterations at the physiological level. An extensive molecular analysis of *bHLH2* expression patterns was also conducted in parallel and is reported in the paragraphs at the end of the chapter.

*bHLH2*RNAi lines show a deficit in electron transfer at the thylakoid chain level

In order to characterize in detail the physiology of the KD2.1 KD2.3 and KD2.4 lines, I performed an analysis of their relative electron transport rate (rETR) in cells adapted under 30 µmol photons s⁻¹ m² and 200 µmol photons s⁻¹ m². This parameter represents the rate of electron transport along the thylakoid redox chain as a function of increasing light intensity and it consequently gives an estimate of the efficiency of the photosynthetic apparatus to convert the incident photon flux into downstream chemical energy. As reported in Fig.8A, *bHLH2* RNAi lines adapted under continuous light at 30 µmol photons s⁻¹ m² showed a similar rETR profile to that of the control lines. Equally, they did not exhibit an alteration of rETRmax (maximum electron flux along the thylakoid chain under saturating light) and of the Ek value, which indicates the light intensity at which such saturation of photosynthetic activity theoretically occurs (Table 5). In the case of cells adapted to continuous light at 200 µmol photons s⁻¹ m², KD2.1 KD2.3 and KD2.4 lines showed an altered rETR profile compared to control lines, with a lower rETRmax as well as a saturation of electron transport at lower light values than controls (Fig.8C, Fig.8D and Table 5). In other words, the mutant lines are unable

to adequately exploit the totality of the incident light and consequently cannot sustain a photosynthetic metabolism comparable to that of the controls. In association with the analysis of growth reported in Fig.6D, this phenotype suggests that a bottleneck at the level of thylakoid electron transport results in a metabolic impairment that prevents the cells from sustaining a normal growth under increasing light conditions.

To further characterize this phenotype, the activity of Photosystem I was also tested on cultures grown under the same conditions. This measure was performed by means of a PSI differential absorbance spectroscopy analysis. A preliminary analysis showed that, when exposed to high light, the *bHLH2* KD2.3 RNAi cells present a more oxidized PSI than the control, indicating a deficit in the rate of electrons reaching this Photosystem, while no difference is observed under low light adapted cells (Fig.8E and Fig.8F). This finding suggests that the anomalies of the mutants are located between PSII and PSI. On the contrary, a higher level of reduction of PSI would have suggested a diminished activity of the metabolism downstream of PSI such, *e.g.* at the level of the Calvin-Benson-Bassham cycle. These analyses remain preliminary at this time and will need to be repeated on the remaining transgenic lines to confirm this observation.

Response of photosynthesis upon transition to high light is affected in *bHLH2*RNAi lines

The light intensity-dependent alterations in photosynthesis described so far have been obtained in cells adapted in continuous light. The lack of a phenotype at the level of cells grown under weak irradiation may have two reasons: alterations in plastid physiology only occur
A

<figure><img src="image_125.png" /></figure>

B

<figure><img src="image_126.png" /></figure>

C

<figure><img src="image_127.png" /></figure>

D

<figure><img src="image_128.png" /></figure>

E

<figure><img src="image_129.png" /></figure>

F

<figure><img src="image_130.png" /><figcaption>Figure 8. bHLH2 RNAi lines show a defect in electron transfer located upstream of PSI</figcaption></figure>

A-D) Wt, RC1, KD2.1, KD2.3 and KD2.4 RNAi lines were grown under continuous light at 30 µmol photons m⁻² s⁻¹ (A and B, n=4, mean ± s.e.m.) or 200 µmol photons m⁻² s⁻¹ (C and D, n=3, mean ± s.e.m.). Panels A and C show the mean rETR of the five lines under increasing light intensities after sampling from the growth conditions. Dashed lines represent the double exponential decay fitting to the raw data. Panels B and D report the maximal PSII quantum yield of the replicates of panels A and C at the moment of the analysis. P-values obtained via One-way analysis of variance against Wt with Bonferroni post-test correction are shown as **P<0.01, ***P<0.001). E-F) PSI is over-oxidised in bHLH2 RNAi line exposed to high light. Representative results of the analysis of PSI redox state performed by differential absorbance analysis at 705nm in control line RC1 and bHLH2 RNAi line KD2.3. Cells grown under 5 µmol photons m⁻² s⁻¹ (E) or exposed to 200 µmol photons m⁻² s⁻¹ for one day (F) were used for the analysis. Graphs report the percentage of oxidised PSI under increasing light intensity steps, normalized to the maximal oxidation observed upon treatment with a saturating dose of DCMU that inhibits electron flow downstream of PSII. Higher oxidation of PSI in KD2.3 exposed to 200 µmol photons m⁻² s⁻¹ locates the defect in electron flow upstream of PSI.
<table><caption>Table 5. Summary results of the RLC analysis</caption><thead><tr><th></th><th>Wt</th><th>RC1</th><th>KD2.1</th><th>KD2.3</th><th>KD2.4</th></tr></thead><tbody><tr><td>30 µmol photons</td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>rETRmax</td><td>89.8±3</td><td>81.9±2</td><td>90.3±3</td><td>81.2±3</td><td>80.6±4</td></tr><tr><td>Ek</td><td>106±4</td><td>101±4</td><td>114±5</td><td>106±6</td><td>94±7</td></tr><tr><td>200 µmol photons</td><td></td><td></td><td></td><td></td><td></td></tr><tr><td>rETRmax</td><td>386±5</td><td>398±7</td><td>277±6</td><td>247±10</td><td>188±11</td></tr><tr><td>Ek</td><td>653±18</td><td>690±23</td><td>496±23</td><td>501±48</td><td>405±60</td></tr></tbody></table>

Maximal rETR and light of saturation of the photosynthetic apparatus (Ek) calculated on the curves reported in Fig.9 A and C via the Platt equation. n=3, mean ± s.e.m.

lines are characterized by an increased sensitivity to transitions from weak to strong light, showing a more pronounced reduction of photosynthetic capacity already in the four hours following light transition (Fig.9A). These results are further matched by the growth curves obtained on cells grown at 5 µmol photons s⁻¹ m⁻² and transferred to 200 µmol photons s⁻¹ m⁻² presented in Fig.9B. As shown, *bHLH2* RNAi lines exhibit a major perturbation of growth rate already after the

under higher photon fluences, whereas at weak irradiations the physiological state is effectively unaltered; a second possibility is that the physiological state of the plastid is affected independently of the growth light intensity and that phenotypes at the level of photosynthetic performance is detected only when the light flux is high enough to be stressful for transgenic strains. In order to clarify this point and to test whether the ability of *bHLH2* RNAi cells to respond to high light is also altered when grown in low light, Wt, RC1, KD2.1 KD2.3 and KD2.4 cultures adapted to 5 µmol photons s⁻¹ m⁻² were transfected at 200 µmol photons s⁻¹ m⁻². Aliquots of the cultures were sampled at 5 and after 20, 60 and 240 minutes from the beginning of the light treatment to characterize their PSII maximal quantum yield. As shown in Fig.9, Wt and RC1 lines show a partial reduction of this parameter following the transition, in accordance with what previously observed in (Lavaud et al., 2016) under similar conditions. This reduction reflects the changes in photosynthetic capacity triggered by irradiation change and can be attributed to the establishment of NPQ phenomena functional to dissipate excess radiant energy, including a photo-damage phenomenon of the D1 protein, a photosensitive component of PSII (Campbell and Tyystjärvi, 2012). Compared to control lines, *bHLH2* RNAi

A

<figure><img src="image_131.png" /></figure>

B

<figure><img src="image_132.png" /><figcaption>Figure 9. Response to light intensity transition is affected in *bHLH2* RNAi lines</figcaption></figure>

A) Wt, RC1, KD2.1 KD2.3 and KD2.4 RNAi lines were grown under continuous light at 5 µmol photons m⁻² s⁻¹ and subsequently exposed to 200 µmol photons m⁻² s⁻¹ for four hours to assess response of PSII maximal quantum yield. Samples were collected before transition (T0) and after 20, 60 and 240 minutes after exposure to high light and their Fv/Fm was evaluated after full relaxation of PSII centers (N=4, mean ± s.e.m.). B) Growth profiles of Wt, RC1, KD2.1 KD2.3 lines grown under 5 µmol photons m⁻² s⁻¹ and subsequently exposed to 200 µmol photons m⁻² s⁻¹ on day 0 (N=3, mean ± s.e.m.).
time of transition to high light and never
recover a growth rate comparable to that of
controls over the course of the analysis.
Overall, these data indicate that plastid
physiology is potentially altered already in cells
adapted to weak light irradiances in the
*bHLH2* RNAi lines and that such
perturbations do not build up progressively
during the exposition to higher light. An
alternative explanation would be that the
perturbation of plastid physiology emerges
with a quick kinetics upon irradiation with
strong light (in the time of few minutes), while
cells are still not affected under low light. Our
experimental set up does not allow us to
discriminate between these two possible
scenarios.

bHLH2 RNAi lines are not affected
in response to photoinhibition

The alterations in photosynthetic
activity shown in the previous paragraphs can
be caused by increased sensitivity to
photoinhibition phenomena, *i.e.* the
reduction of photosynthetic efficiency due to
light induced damage of D1. To verify this
possibility, the *bHLH2* RNAi lines under
study grown under 5 µmol photons s⁻¹ m² were
subject to a treatment with 500 µg ml⁻¹ of
lincomycin, an inhibitor of plastid protein
translation (Lavaud et al., 2016), and moved to
500 µmol photons s⁻¹ m² for inducing photo
degradation of D1. Such a treatment allows to
highlight the loss of PSII quantum yield in a
condition where the damaged D1 cannot be

<figure><img src="image_133.png" /></figure>

<figure><img src="image_134.png" /></figure>

<figure><img src="image_135.png" /><figcaption>Figure 10. bHLH2 RNAi lines are not affected in photoinhibition.</figcaption></figure>

A) Wt, RC1, KD2.1 KD2.3 and KD2.4 RNAi lines were grown under continuous light at 5 µmol photons m⁻² s⁻¹ and subsequently exposed to 500 µmol photons m⁻² s⁻¹ for one hour in presence of 500µg ml⁻¹ of lincomycin to assess sensitivity to photoinhibition of D1 protein when plastid translation is inhibited. After high light treatment cells were transferred for 2 hours under 10 µmol photons m⁻² s⁻¹ to grant full relaxation of NPQ processes. PSII quantum yield under light irradiation ΦPSII is reported in the graph (n=3, mean ± s.e.m.). B) ΦPSII relaxation of Wt, RC1, KD2.1 KD2.3 and KD2.4 RNAi lines grown under continuous light at 200 µmol photons m⁻² s⁻¹ and subsequently exposed to 5 µmol photons m⁻² s⁻¹ to induce relaxation of PSII reaction centers under non saturating light irradiation (n=3, mean ± s.e.m.). C) qE component of NPQ in the same samples reported in panel A. NPQ values were calculated considering the cells at the end of the kinetics (t=26 minutes) as completely relaxed. qE values at time t were hence calculated as (Fm'₁-Fm'₂₆)/Fm'₂₆.
replaced by newly synthesized subunits, hence
making the photodamage of D1 itself the only
factor involved in the kinetics of quantum
efficiency reduction. As shown in Fig.10A, no
difference emerged between control lines and
mutants, hence excluding an increased
sensitivity to photodamage under deregulation
of *bHLH2* gene. An alternative explanation
for the impaired maximum photosynthetic
capacity of the mutants is that it is not the
sensitivity of PSII to photodegradation that is
altered, but rather the ability of the cells to
replace degraded proteins, *i.e.* an alteration in
protein resynthesis at the plastid level that
causes an unbalance between the degradation
of D1 subunits (that occurs at a normal rate)
and its substitution with neo synthesized
subunits. This hypothesis is supported by the
fact that cultures grown under continuous light
at 200 µmol photons s⁻¹ m⁻² show a slower
recovery of photosynthetic activity than
controls once moved to non-stressful low light
(5 µmol photons s⁻¹ m⁻² (Fig.10B) Analysis of
qE component of NPQ of the same cells upon
relaxation (Fig.10C), highlights that *bHLH2*
RNAi strains manifest a maximal NPQ at 200
µmol photons s⁻¹ m⁻² (time zero) comparable to
control strains. qE shows a slower relaxation in
*bHLH2* RNAi lines in the first minutes after
the transition to non-stressful light, but this
difference still does not account for the
impairment in ΦPSII shown by transgenic
lines after 25minutes from transition.

Photosynthetic activity represses
*bHLH2* expression

Available transcriptomic data of *P. tricornutum* cells exposed to different light conditions (Chauton et al., 2013; Matthijs et al., 2016; Nymark et al., 2013; Smith et al., 2016; Valle et al., 2014) show that *bHLH2* expression is strongly regulated by light cues. Its transcription is induced after a few hours in the dark and repressed following the dark to light switch. This pattern is independent of the photoperiod used for growing the cultures and

of the light colour and shows no anticipation
of dark to light transitions, differently from
what is observed for *RITMO1* and *bHLH1b*
(Fig.11) This data suggests that inhibition of
*bHLH2* transcription by light can be related to
the activity of the plastid. Plastid development
and gene expression are tightly regulated by
nuclear encoded factors, a process called
anterograde control (Eberhard et al., 2008).
However, the plastid itself is in turn capable of
transmitting information about its functional
state to the nucleus to regulate expression of
Photosynthesis Associated Nuclear Genes
(PhANGs:). This signalling system originating
in the plastids is referred to as retrograde
signalling (Eberhard et al., 2008; Kleine et al.,
2009). In diatoms, one of the main markers of
photosynthetic activity to be involved in
retrograde signalling is the redox state of
plastoquinones (PQs see Chapter I, section
I.3.a), similarly to what occurs in primary
endosymbiotic phototrophs (Kleine et al.,
2009; Lepetit et al., 2013). In *P. tricornutum*,
the redox state of PQs has been shown to
regulate the expression of a number of
PhANGs (Lepetit et al., 2013; Taddei et al.,
2016). To test the hypothesis that *bHLH2*
transcription is controlled by this process,
qRT-PCR analysis of *bHLH2* expression was
performed on Wt cells adapted in the dark for
50 hours and then transferred back to light in
the presence and in the absences of DCMU
(3-(3,4-dichlorophenyl)-1,1-dimethylurea).

DCMU acts as a competitor of the mobile
plastoquinone as electron acceptor on the
donor side of PSII and induces the arrest of
the linear electron flow along the thylakoid
(Kyle, 1985; Metz et al., 1986; Trebst, 2007).
As reported in Fig.12A, control replicates
show a drastic reduction of *bHLH2*
expression upon illumination, with *bHLH2*
transcript being ten times less expressed
relative to dark after one hour of light
exposure and remaining constant to this
expression level after 4 hours, but still
detectable. In cells treated with DCMU the
repression of *bHLH2*, although still present, is
strongly slowed down. This result is
<figure><img src="image_136.png" /></figure>

<figure><img src="image_137.png" /></figure>

<figure><img src="image_138.png" /></figure>

<figure><img src="image_139.png" /></figure>

<figure><img src="image_140.png" /></figure>

<figure><img src="image_141.png" /><figcaption>Figure 11. Expression profiles of *bHLH2* from publicly available transcriptomic data</figcaption></figure>

Expression profiles of *bHLH2* under different illumination conditions as retrieved from different publicly available transcriptomic datasets. A) 12L :12D grown cells from Smith et al. 2014. High iron concentration was chosen as a representative study case from the article. *bHLH2* expression profiles did not appear to be altered by different iron concentrations (data not shown). B) Cells grown under 16L:8D from Chauton et al. 2013. Cells treated with 48h dark and exposed to white (C, from Nymark et al. 2013) red, blue or green light (D-F, from Valle et al. 2014). Expression values were normalized to the maximal expression of *bHLH2* during the course of the time course. Grey boxes indicate dark exposure, white boxes indicate exposure to white light, while red, blue and green boxes refer to exposure to different light colours. Further information on light quality and intensity can be found in the relative papers.
comparable to what is shown by other light-inhibited PhANGs such as the one encoding for the LHCX4 antenna (Taddei et al., 2016). The fact that inhibition of photosynthetic activity significantly slows down the repression of *bHLH2* expression demonstrates that its transcriptional regulation in the light depends on the activity of the electron transport chain. These results corroborate the existence of a regulatory cross-talk between plastids and nucleus for the optimization of diatom growth capacity under changing lights, a process in which *bHLH2* appears to play a relevant role. To note, *bHLH2* protein was not predicted to be targeted to the plastid by HECTAR and ASAFind localization tools, while all other tools used (see Materials and methods) identified a NLS in *bHLH2* sequence. Given its possible function as a bHLH-PAS transcription factor, we can speculate that *bHLH2* can act as a relevant regulator of PhANGs expression in response to light inputs and, consequently, in photoacclimation. Still, further characterization of *bHLH2* target genes is needed to deepen this hypothesis.

Given the information on dark transcription of *bHLH2*, we tested the effect of the antisense construct against *bHLH2* in the absence of light. To do so, the Wt, RC1, KD2.1 KD2.3, and KD2.4 lines were grown in a 12L:12D photoperiod and harvested at the end of the dark phase, when Wt expression levels are expected to be highest (Chauton et al., 2013; Smith et al., 2016). In a second analysis, the five lines pre-adapted in continuous light were exposed to 24 hr of darkness and sampled for RNA extraction to

verify the effect of prolonged dark treatment on *bHLH2* RNAi lines. As shown in Fig. 12B and Fig. 12C, the results obtained do not allow to demonstrate any major alteration in the expression level of *bHLH2* under these conditions compared to Wt, thus making it inconvenient to continue the analysis of photosynthetic physiology under these conditions. Indeed, a correlation between any

<figure><img src="image_142.png" /><figcaption>Figure 12. *bHLH2* expression in the dark</figcaption></figure>

A) Wt cells were left in the dark for 50 hours to induce maximal expression of *bHLH2* gene before being exposed to light (30 µmol photons m⁻² s⁻¹) in presence or absence of 2µM of DCMU. Cells were collected just before exposure to light and after 1 and 4 hours from exposure to extract RNA and determine *bHLH2* expression. Expression values are normalized to the mean expression of Wt strain. n=3, mean ± s.e.m.
B) Expression of *bHLH2* in Wt, RC1, KD2.1 KD2.3 and KD2.4 RNAi lines grown under 12L:12D and collected at ZT0 before transition to light.
C) Expression of *bHLH2* in Wt, RC1, KD2.1 KD2.3 and KD2.4 RNAi lines grown under continuous 5 µmol photons m⁻² s⁻¹ and transitioned for 24 hours under dark before being collected for RNA extraction. Expression values are normalized to the mean expression of Wt strain. n=2, mean ± s.e.m.
phenotypes in plastid physiology with the level of *bHLH2* messenger RNA is currently not possible. In the absence of antibodies to check for possible alterations of *bHLH2* at the protein level, we decided to generate Knock-Out lines to confirm the correlation of the phenotypes seen so far with a role of *bHLH2* in chloroplast regulation. Preliminary results are reported in the following paragraph.

## Preliminary characterization of bHLH2 KO mutants

KO mutants for *bHLH2* gene were generated in the laboratory by Marianne Jaubert. These mutants were generated to allow bHLH2 characterization in more detail in a controllable and stable genetic background. The mutagenesis of *bHLH2* was performed by proteolistic transformation of a CRISPR/Cas9 system in a manner similar to that performed for the mutagenesis of RITMO1 reported in Chapter II.c. By Sanger sequencing it was possible to identify two independent lines presenting a biallelic mutation at the *bHLH2* locus, henceforth

called RNP1 and RNP2 (for RiboNucleoProtein bombardment KO1 and 2). Sequencing was performed using a non-allele-specific primer to ensure early identification of biallelic mutants. The resulting chromatograms reported in Fig.13 demonstrate that RNP1 mutant presents a 67bp deletion in both alleles at the nt-65 position, implying a double cut of the Guide_bHLH2_59 and Guide_bHLH2_135 sgRNA target sites (Fig.14). The RNP2 mutant exhibits a chromatogram compatible with a 67bp deletion in one allele (as in RNP1) and a 2bp deletion in the second allele at the nt-65 position, at the level of the Guide_bHLH2_59 target site (Fig.13 and Fig.14). In all cases, the mutations induce a frameshift that causes a disruption of the downstream amino acid sequence. A more detailed characterization of the *bHLH2* gene by allele-specific sequencing is currently underway to precisely characterize the effect of the mutation on the two *bHLH2* alleles in the RNP2 mutant and determine the resulting sequences. An additional colony resistant to 2-FA but not targeted for *bHLH2* locus mutagenesis was chosen as control (CTR line).

<figure><img src="image_143.png" /><figcaption>Figure 13. Mutants on *bHLH2* gene induced by CRISPR/Cas9</figcaption></figure>

Chromatographic traces obtained by Sanger sequencing of the *bHLH2* locus in Wt, RNP1 and RNP2 lines. Dashed line highlights the nt-65 position where mutations occurred in both alleles in both RNP1 and RNP2 lines. RNP1 mutant show an identical 62 bp deletion starting from nt-65 position, leading to a single chromatographic trace downstream of the mutation site. RNP2 mutant show a 62 bp deletion in one of the two alleles starting at nt-65 and a 2bp (AC) deletion in the second allele at the same position.
<figure><img src="image_144.png" /><figcaption>Figure 14. Sequence alignment of the KO mutants for *bHLH2* gene.</figcaption></figure>

Alignment of the reference coding sequence in Pt1 Wt strain with that of the *bHLH2* KOs RNP1 and RNP2. The alignments were generated via EBI Clustal Omega webtool and visualized via Mview. Bases are colored by identity to the Wt reference sequence. sgRNA Guide_bHLH2_59 and Guide_bHLH2_135 target sites are highlighted in dark and light purple respectively. Reference coding sequence of *bHLH2* was retrieved from EnsemblProtist web page
Currently, physiological characterization of KO mutants of bHLH2 has been initiated in order to determine the precise role of this gene in the regulation of plastid activity and to verify a possible consistency of the phenotypes of these lines with that shown by RNAi lines. A first analysis was performed to verify if these lines present a response to the transition from weak light conditions, 5 µmol photons s⁻¹ m⁻², to 200 µmol photons s⁻¹ m⁻² similar to that shown by the KD2.1 KD2.3 and KD2.4 lines. As preliminary, the results shown in Fig.15 demonstrate that the KO bHLH2 mutants exhibit a response to such light transitions similar to that shown by the RNAi lines and, likewise, a more prominent reduction in the quantum yield of PSII than the control lines. A full range of analyses comparable to those performed for RNAi mutants is currently planned to elucidate the function of *bHLH2*. Similarly, special attention will be paid to verify the responses of KO lines in the transitions from dark to light, which is currently not probed in RNAi transgenic lines and which, from what has been reported in the previous paragraphs, represents one of the conditions under which the role of this protein should emerge more strongly.

<figure><img src="image_145.png" /><figcaption>Figure 15. Response to light intensity transition in bHLH2 KO lines</figcaption></figure>

Wt, CTR, RNP1 and RNP2 lines were grown under continuous light at 5 µmol photons m⁻² s⁻¹ and subsequently exposed to 200 µmol photons m⁻² s⁻¹ for four hours to assess response of PSII maximal quantum yield. Samples were collected before transition (T0) and after 20, 60, 180 and 240 minutes after exposure to high light and their Fv/Fm was evaluated after full relaxation of PSII centers (N=2, mean ± s.e.m.). The results for KD2.4 RNAi line showed in Fig.10 A are overlayed to allow the comparison between RNAi and KO lines