# Red light induces starch accumulation in *Chlorella vulgaris* without affecting photosynthesis efficiency, unlike abiotic stress

A. Six, A. Dimitriades-Lemaire, C. Lancelon-Pin, J.-L. Putaux, D. Dauvillée, D. Petroutsos, P. Alvarez Diaz, J.-F. Sassi, Y. Li-Beisson, G. Fleury

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## To cite this version:

A. Six, A. Dimitriades-Lemaire, C. Lancelon-Pin, J.-L. Putaux, D. Dauvillée, et al., Red light induces starch accumulation in *Chlorella vulgaris* without affecting photosynthesis efficiency, unlike abiotic stress. Algal Research - Biomass, Biofuels and Bioproducts, 2024, 80, pp.103515. 10.1016/j.algal.2024.103515. hal-04629808

HAL Id: hal-04629808

https://hal.science/hal-04629808v1

Submitted on 10 Oct 2024

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# Red light induces starch accumulation in *Chlorella vulgaris* without affecting photosynthesis efficiency, unlike abiotic stress

A. Six<sup>a</sup>, A. Dimitriades-Lemaire<sup>a</sup>, C. Lancelon-Pin<sup>b</sup>, J.-L. Putaux<sup>b</sup>, D. Dauvillée<sup>c</sup>, D. Petroutsos<sup>d</sup>, P. Alvarez Diaz<sup>a</sup>, J.-F. Sassi<sup>a</sup>, Y. Li-Beisson<sup>e</sup>, G. Fleury<sup>a,∗</sup>

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## ARTICLE INFO

**Keywords:**

*Chlorella vulgaris*

Starch

Light quality

Nitrogen deprivation

Supra-optimal temperature

Bioplastics

## ABSTRACT

Microalgae show great promise as sources of starch, one of the most widely consumed macromolecules. In this study, we evaluated the impact of three starch-inducing factors, namely nitrogen deprivation, supra-optimal temperature, and red light, on the physiology and starch accumulation capacity of *Chlorella vulgaris*. This starch accumulation was monitored by measuring the total carbohydrate content and transmission electron microscopy (TEM) imaging. Nitrogen deprivation and a supra-optimal temperature of 39 °C resulted in carbohydrate contents of 69.7 and 64.3 % of dry weight (DW) respectively. This constituted a 5.3- and 3.3-fold increase in carbohydrate productivity compared to the control, after 4 days of cultivation. During this period, carbohydrates represented over 80 % of the produced material (DW basis). However, nitrogen deprivation and supra-optimal temperature were accompanied by extensive stress, leading to lower cell division rates and damage to the photosynthetic apparatus. Red light illumination resulted in a more moderate production of carbohydrates. After 4 days of cultivation, the carbohydrate content reached 46.8 %, representing a 3.0-fold increase in productivity compared to control. The composition of the starch formed under red light was surprisingly poor in amylose, similar to transitory-type starch rather than storage starch. Most notably, the starch accumulation under red light was sustained over 7 days without affecting the rate of cell division and quantum yield efficiency. To the best of our knowledge, red light is the only factor reported so far to induce a significant starch accumulation without hindering cell division and photosynthesis efficiency, even after long-term exposure (7 days). Furthermore, all three conditions induced a cell wall thickening, albeit without affecting the recovery of accumulated starch by high-pressure homogenization. These results highlight the potential of red light as a starch inducer in *Chlorella vulgaris* and open up perspectives for the production of starch-based bioplastics from microalgae.

## 1. Introduction

Starch is a polymer of glucose, composed of amylose and amylopectin, that acts as a form of carbon and energy storage for plants and green microalgae. Its highly structured granular structure represents a compact glucose reservoir, able to deliver substantial energy to organisms in food, feed and fermentation applications. As a polymer, starch displays notable properties of sizing and binding agent that are largely exploited in papermaking and textile industries [1]. This combination of energy reserve and mechanical properties makes starch highly regarded

as a primary feedstock for the bioplastics industry. Indeed, the main types of biosourced-biodegradable plastics can be prepared from starch, either by fueling fermentation processes after saccharification (polylactic acid, polyhydroxyalkanoates) or by direct plasticization of the polymer (thermoplastic starch).

The increasing demand for amylaceous feedstock drives the exploration of new ways to produce starch. In this regard, microalgae have a strong complementarity with current starch crops [2]. While non-competing for arable land, microalgae can recover nutrients from wastewaters and recycle CO₂ from industrial processes. Yet, the full potential of microalgae as a reliable source of biomass is still to be

<footnote><sup>a</sup> CEA, MicroAlgae Processes Platform, Cadarache, F-13108 St Paul Lez Durance, France</footnote>

<footnote><sup>b</sup> Univ. Grenoble Alpes, CNRS, CERMAV, F-38000 Grenoble, France</footnote>

<footnote><sup>c</sup> Univ. Lille, CNRS, UMR 8576 - UGSF - Unité de Glycobiologie Structurale et Fonctionnelle, F-59000 Lille, France</footnote>

<footnote><sup>d</sup> Department of Organismal Biology, Uppsala University, 75236, Uppsala, Sweden</footnote>

<footnote><sup>e</sup> Aix Marseille Univ, CEA, CNRS, Institute of Bioscience and Biotechnology of Aix Marseille, BIAM, Cadarache, France</footnote>

<footnote>* Corresponding author.</footnote>

<footnote>E-mail address: gatien.fleury@cea.fr (G. Fleury).</footnote>
<table><thead><tr><th colspan="2">Abbreviations</th><th></th><th></th></tr></thead><tbody><tr><td></td><td></td><td>QY<sub>max</sub></td><td>Maximal quantum yield</td></tr><tr><td></td><td></td><td>LC</td><td>Light curve</td></tr><tr><td>N+</td><td>Control growth condition with complete media, optimal temperature and white light illumination</td><td>F<sub>S</sub></td><td>Relative fluorescence yield in steady-state illumination</td></tr><tr><td>N-</td><td>Nitrogen-deprived media condition</td><td>F<sub>M</sub></td><td>Relative maximum fluorescence yield in dark-adapted conditions</td></tr><tr><td>HT</td><td>Harsh (i.e. supra-optimal) temperature condition</td><td>F'<sub>M</sub></td><td>Relative maximum fluorescence yield during illumination</td></tr><tr><td>RL</td><td>Red-light illumination condition</td><td>ΦPSII</td><td>Quantum yield of PS II photochemistry at steady-state</td></tr><tr><td>OD<sub>880</sub></td><td>Optical density measured at 880 nm</td><td>ΦNPQ</td><td>Quantum yield of ΔpH- and xanthophyll-regulated thermal energy dissipation at steady-state</td></tr><tr><td>DW</td><td>Dry weight</td><td>Φf,d</td><td>Φf – quantum yield of fluorescence, Φd – quantum yield of constitutive thermal energy dissipation at steady state</td></tr><tr><td>wt%</td><td>Percentage of dry weight</td><td>ΔpH</td><td>Trans-thylakoid pH gradient</td></tr><tr><td>λ<sub>max</sub></td><td>Wavelength at the maximal absorbance</td><td>HPH</td><td>High-pressure homogenizer</td></tr><tr><td>Chla/b</td><td>Ratio of chlorophyll a and chlorophyll b</td><td></td><td></td></tr><tr><td>Chla</td><td>Chlorophyll a</td><td></td><td></td></tr><tr><td>Chlb</td><td>Chlorophyll b</td><td></td><td></td></tr></tbody></table>

unlocked, and requires further development of the whole production chain. Abiotic stress is known to further increase energy storage in green microalgae, either in the form of lipids or starch depending on the stress duration and the strain metabolism [3,4]. Many abiotic factors trigger starch accumulation in green microalgae, namely addition of cell division inhibitors like cycloheximide or deuterium [5,6], nutrient deprivation [7], supra-optimal temperature [8] or light quality [9].

Macronutrient deprivation (N, P, S) is generally acknowledged to be the most efficient strategy for starch accumulation. Phosphorous is the less praised starvation, as it triggers high starch accumulation but is not optimal for both productivity and ease of operation [10]. Conversely, sulfur starvation is recommended for its stable and high starch accumulation with sustained photosynthesis efficiency [5,11]. In comparison, nitrogen starvation is considered more harmful for the cells despite a high accumulation of starch [10]. However, the comparison of nitrogen and sulfur starvation in two different strains of *Chlorella vulgaris* led to contradictory starch accumulation results [5,12]. Among these various methods, nitrogen starvation is the most commonly used method to induce carbon reserve accumulation. Overall, nitrogen starvation is regarded as a practical yet efficient strategy for starch accumulation that can reach 60 % in *Chlorella* species [13].

The impact of temperature on starch production is less studied, but is interesting for application in closed photobioreactor (PBR) systems under natural light, where the temperature increases naturally with solar light availability. Supra-optimal temperature triggers starch accumulation in *Chlamydomonas reinhardtii* and *Parachlorella kessleri* when algae are grown at 39 or 40 °C, respectively [6,14].

Light quality manipulation is proposed as a way to optimize microalgal cultivation. Not only changing light quality can improve growth dynamics in *Chlorella vulgaris* [15,16], but applying only red light also increases the carbohydrate content in *Chlamydomonas reinhardtii* and in *Chlorella* biofilms [17,18]. Interestingly, the addition of a small amount of blue light reverses the carbohydrate accumulation promoted under red light [19]. Likely, blue light directly triggers the catabolism of carbohydrates, as shown in *Chlorella* [20].

In this study, we evaluated the response of the microalga *Chlorella vulgaris* CCALA924 to three starch induction methods: nitrogen starvation, supra-optimal temperature and red-light illumination. Although these starch inducers have previously been described in the literature, a thorough comparison of these methods represented an unaddressed area. Our study aimed at filling this gap by providing a detailed analysis and comparison of these three inducers. Starch accumulation kinetics were thus investigated, and the starch structure, photosynthetic apparatus status and biomass composition were analyzed. We also assessed the starch recovery from the microalgae by examining cell rupture under high-pressure homogenization.

## 2. Materials & methods

### 2.1. Strain and growth conditions

*Chlorella vulgaris* CCALA924 was purchased from the Culture Collection of Autotrophic Organisms (CCALA, Czech Republic). This strain was originally identified as a promising starch producer by Brányiková et al. [5]. Cultures were grown autotrophically in an automated incubator ZWYC 290 A (LABWIT Scientific, Melbourne, Australia) under constant environmental conditions (photoperiod: 16-h light and 8-h darkness, agitation: 150 rpm, 2 vol% CO₂). The built-in white LEDs were used for all the experiments, except for the RL experiment, where they were substituted by red LEDs. A comprehensive description of these two light sources is given in the next section. Microalgae were cultivated in 500-mL Erlenmeyer flasks (Schott, Germany) filled with 200 mL of culture. The nutritive medium was a modified version of the Beijerinck medium, containing 26.15 mM NaNO₃ (instead of the usual 7.5 mM of NH₄Cl), 0.34 mM CaCl₂·2H₂O, 0.41 mM MgSO₄·7H₂O, 0.52 mM K₂HPO₄, 0.39 mM KH₂PO₄, 0.18 mM BO₃H₃, 0.077 mM ZnSO₄·7H₂O, 0.026 mM MnCl₂·4H₂O, 0.018 mM FeSO₄·7H₂O, 0.007 mM CoCl₂·6H₂O, 0.006 mM CuSO₄·5H₂O, 0.001 mM Mo₇O₂₄(NH₄)₆·4H₂O, 19.98 mM C₇H₁₅NO₄S. The pH was adjusted to 6.8 with 1 M NaOH (around 6 mL L⁻¹). For nitrogen-deprived media, NaNO₃ was omitted. The cells were routinely maintained in these conditions by applying a weekly dilution, until the acclimation phase described afterwards started.

### 2.2. Experimental design of starch induction

Before starting the starch accumulation experiment, exponential phase cultures were acclimated to the control conditions. In addition to photoperiod and CO₂ concentration, the control conditions included growth in complete medium, a temperature of 25 °C and white-light illumination. This acclimation phase lasted for 4 days, during which the cultures grew approximately from 0.3 to 1.6 OD₈₈₀ (optical density measured at 880 nm).

For every condition tested, the inoculum culture was centrifuged at 4500 g (Allegra X15R, rotor model: SX4750A, Beckman Coulters, USA) for 5 min. The supernatant was discarded and the biomass pellet was resuspended in fresh medium. In the case of nitrogen starvation, the renewed medium was nitrogen-free. For supra-optimal temperature, the incubator temperature was adjusted to 39 °C. This temperature was arbitrary chosen among those (39 and 40 °C) reported to induce starch accumulation in green microalgae [6,14]. For red-light illumination, 4 red LEDs (LH CPxP, Osram, Germany) were set up in the incubator.

Emission spectra measurements (BTS256-E, Gigahertz-Optik GmbH, Germany) confirmed that red LEDs emitted within the red color range only (600–700 nm), whereas built-in white LEDs emitted in the
photosynthetically active radiations and near-infrared domain (400–800 nm), as illustrated in Fig. 1. The photon flux density was measured at the stirred plate level of the incubator (15 measurements in different locations) with a ULM-500 light meter and its MQS-B flat sensor (Walz, Germany). Despite differences in light homogeneity, the mean photon flux density was similar for the white LEDs of the incubator and the red LEDs, with respectively 184 µmol$_{photon}$ m$^{-2}$ s$^{-1}$ and 197 µmol$_{photon}$ m$^{-2}$ s$^{-1}$ (n = 15).

## 2.3. Microalgal biomass characterization

Cultures were grown in triplicate flasks. We ensured that each flask of the triplicate behaved similarly to the others by measuring the OD$_{880}$ nm and maximal quantum yield (QY$_{max}$) before each sampling. The corresponding data, showing an excellent homogeneity among the biological triplicate, is provided as Supplementary Material (Figs. S7 and S8). At each sampling, an equivalent volume of culture was sampled from each flask of the biological triplicate and mixed. This mixture from the three different flasks was subsequently analyzed. All analyses were conducted in technical triplicates unless specified otherwise.

### 2.3.1. Growth monitoring

The microalgal growth was monitored by determining the number and size of the cells, OD$_{880}$ nm, and dry weight in suspension. OD$_{880}$ nm was evaluated by measuring the optical absorption in 1-cm light path cuvettes (ref. 11,602,609, ThermoFisher, USA) of 10-fold diluted samples with a spectrophotometer (UV-Vis Epoch2, BioTek Instruments, USA). The corresponding data of each biological triplicate is provided as Supplementary Material (Fig. S7). The blank, measured with a cuvette filled with fresh medium, was subtracted from all measurements. A technical triplicate was made for each measurement. The cell density and cell volume were determined using a particle sizer and counter (Multisizer 4 Counter, Beckman Coulter, USA). Depending on the optical density of the sample, between 50 and 100 µL of 10-fold pre-diluted culture was suspended in 10 mL of filtered saline water (Isoton® II Diluent, Beckman Coulter, USA). The cellular exponential growth rate µ was calculated as $µ = \ln(C_2/C_1)/(t_2-t_1)$ with C$_x$ the cell concentration at time t$_x$ (in days). The dry weight (DW) in suspension was determined from 4 to 5 mL of culture. Briefly, culture aliquots were centrifuged at 4500 g for 10 min at 4 °C. The pellets were washed with pure water and centrifuged again to eliminate any remaining mineral solutes. The washed pellets were transferred to pre-weighed and pre-dried aluminum cups. The samples were then stored in a ventilated oven at 105 °C for 24

<figure><img src="image_5.png" /><figcaption>**Fig. 1.** Emission spectra of white LEDs used in N+, N-, HT experiments and red LEDs used for RL starch induction. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>

h. The weight loss upon drying was used to calculate DW and corresponding concentration. The supernatant of the centrifuged culture was recovered to determine nitrate, phosphate, and sulfate concentrations by ion chromatography (940 Professional IC Vario, Metrohm, Switzerland) as described in the Supplementary Material section.

### 2.3.2. Biomass analysis

During sampling, known volumes of culture were centrifuged, frozen at –20 °C, and dried (lyophilizer COSMOS 20 K, Cryotec™, France). The dry weight of samples was calculated from the dry measurement performed for growth monitoring. The samples were then hermetically sealed and stored in the dark at –20 °C until analysis.

#### 2.3.2.1. Carbohydrate and starch-equivalent content.
The starch accumulation was evaluated by the quantification of total carbohydrates. The total carbohydrate content was measured using the protocol developed by Dubois et al. [21]. Briefly, 1–3 mg samples of lyophilized biomass were digested in 1.25 M H$_2$SO$_4$ (0.5 mL mg$^{-1}$ DW) at 100.5 °C for 3 h. The digestate was diluted in ultra-pure water to a final volume of 0.5 mL and mixed with 500 µL of 5 % (w/v) phenol solution and 2.5 mL of 95 % H$_2$SO$_4$. The total carbohydrate content in the suspension was determined by comparing the light absorption at 483 nm with a glucose calibration curve.

#### 2.3.2.2. Protein content.
The protein content in the microalgae was measured using the nitrogen content as a proxy, as described in Chambonniere et al. [22]. To evaluate total nitrogen (TN) and total carbon (TC) contents in the biomass, about 10 mg of dry biomass were sampled in the liquid culture, centrifuged, and washed. The resulting pellet was freeze-dried, and the dry pellet was manually ground with a mortar, to homogenize the particle size. The dry biomass was then suspended in a 50-mL bottle and kept under magnetic stirring to prevent precipitation. The analyses were carried out on the Shimadzu TOC-LCSH system (Shimadzu™, Japan) equipped with an 8-channel OCT-L autosampler and a TNM-L Total Nitrogen module. After combustion at 720 °C in a quartz tube, the O$_2$ carrier gas (Alphagaz Air 1, Air Liquide, France) led to infrared detection (NDIR) thermostated at 65 °C, used for total carbon (TC), and to a chemiluminescence detection module for total nitrogen (TN). The flow was set at 150 mL min$^{-1}$. The protein fraction in the algal biomass was computed assuming a nitrogen-to-protein ratio of 5.04, as reported for *Chlorella vulgaris* [23].

#### 2.3.2.3. Lipid content.
The lipid content and profile were assessed by total fatty acids analysis using GC-FID (GC-2010 Pro AOC-20i/AOC-20s, Shimadzu™, Japan). Lyophilized biomass samples (2–10 mg) underwent transmethylation to produce fatty acid methyl esters (FAMEs). Briefly, the samples were suspended in 3 mL of a transesterification agent (1.25 M hydrogen chloride in methanol, ref: 17935, Supelco, USA) and combined with 0.2 mL of a 3 mg L$^{-1}$ TAG C15:0 solution (Tripentadecanoin, ref. T4257, Sigma-Aldrich™, USA) in anhydrous hexane 95 % (ref. 296,090, Sigma-Aldrich™, USA) as an internal standard. This mixture was incubated at 85 °C for 1 h. Subsequently, 3 mL of HPLC-grade hexane was added, followed by vigorous mixing with 1 mL ultra-pure water. After centrifugation (524 g, 5 min), the upper hexane phase was collected for GC analysis. The total lipid content was calculated by comparing the sum of peak areas to that of the internal standard. Compound identification utilized the retention index method and a set of standards of 37 FAMEs (ref. CRM47885, Supelco, USA).

#### 2.3.2.4. Pigment content.
The pigment content was measured by Wellburn's colorimetric method [24] after pigment extraction in dimethyl sulfoxide (DMSO). Briefly, pure DMSO was added in samples of 0.5–1.5 mg of freeze-dried biomass to reach 2 mL mg$^{-1}$ DW. Samples were heated in the dark at 60 °C for 40 min. After centrifugation, the optical density of supernatant was measured at 450, 649 and 665 nm in 1-cm
light path cuvette (ref. 11,602,609, ThermoFisher, USA) using a UV-Vis Epoch2 (BioTek Instruments, USA). Concentrations in chlorophyll a, chlorophyll b and total carotenoids were calculated according to Well-burn's Eqs. [24] and detailed in the Supplementary Material section. The total chlorophyll content was calculated as the sum of chlorophyll a and chlorophyll b contents.

### 2.3.2.5. Calculation of productivities and carbohydrate conversion into starch values.
The productivity in biomass, carbohydrates, lipid, protein and pigment were calculated as $\frac{C_2-C_1}{t_2-t_1}$, with $C_i$ the concentration at time $t_i$ (in hours). In the following, reported productivities are calculated from $t_1 = 0$ and $t_2 = 88$ h. Other periods are summarized in Table S3. The measurements and calculation of carbohydrate productivities were used to evaluate the starch productivities. We assumed that the measured increase in carbohydrate content during our experiments was only related to the starch increase. Furthermore, we converted carbohydrate values into starch by applying a multiplication factor of 0.9, to take into account the added H₂O molecules during starch hydrolysis into glucose, as proposed for instance by Brányiková et al. [5].

## 2.4. Photosystem II status

The photosynthetic apparatus status was qualified at the photosystem II (PSII) level using quantum yield protocol (QY), non-photochemical quenching protocol (NPQ3) and light curve assay (LC3) of the AquaPen-C AP 100-C fluorometer (Photon Systems Instruments, Czech Republic). Protocols details are described in the manufacturer manual and user guide. The fluorometer was set with the following parameters: measuring color: 630 nm, measuring flash pulse: 1000 µmol$_{photon}$ m⁻² s⁻¹, saturating pulse: 1500 µmol$_{photon}$ m⁻² s⁻¹, actinic light: 100 µmol$_{photon}$ m⁻² s⁻¹.

Soon after culture sampling, fresh microalgae cultures were diluted to an OD₈₈₀ nm of 0.1–0.3 in 1-cm cuvette and placed in a dark enclosure at room temperature for at least 5 min. After dark adaptation and the opening of PSII reaction centers, the response of chlorophyll a fluorescence to the different patterns of light stimulus was measured. The three measurement protocols were conducted once per condition and time. As the maximal quantum yield (QY$_{max}$) was measured for each of the three protocols, the reported results of QY$_{max}$ were calculated from triplicates.

Partitioning of absorbed light energy was calculated from the parameters $F_S$, $F_M$, $F'_M$ – relative fluorescence yield in steady-state illumination, relative maximum fluorescence yield in dark-adapted conditions or during illumination, respectively [25]. First, the fraction of photons absorbed by the PSII antenna allocated to PSII and photosynthetic electron transport was calculated as $\Phi$PSII = $1 - F_S/F_M'$. The fraction of regulated energy dissipation due to trans-thylakoid pH gradient- (ΔpH) and/or xanthophyll-dependent non-photochemical quenching (NPQ) was calculated as $\Phi$NPQ = $F_S/F_M' - F_S/F_M$. Finally, the fraction lost by either constitutive energetic dissipation via heat or fluorescence, i.e. the non-regulated and non-photochemical energy dissipation (f, d), was calculated as $\Phi$f,d = $F_S/F_M'$.

## 2.5. Electron microscopy and cell wall thickness measurement

At 0, 40, and 88 h of treatment, the cells were fixed in suspension with a solution of 2.5 % glutaraldehyde and 0.1 M PBS (pH 7.2). Aliquots were post-fixed with reduced osmium tetroxide / potassium ferrocyanide (1 % OsO₄ / 1.5 % K₃Fe(CN)₆), then treated with 1 % tannic acid in 0.1 M PBS, dehydrated in water / ethanol solutions with increasing ethanol concentration, infiltrated with an ethanol / EMbed 812 resin mixture, and embedded in pure EMbed 812 resin (Electron Microscopy Sciences, USA). 70-nm-thick sections were cut with a diamond knife in a Leica UC6 ultramicrotome. The sections were collected on carbon-coated copper grids, stained with uranyl acetate and lead citrate, and observed with a JEOL JEM-2100-Plus microscope operating at an

accelerating voltage of 200 kV. Images were recorded with a Gatan Rio 16 camera. The cell wall thickness was measured using the ImageJ software (National institutes of Health, USA) in 3 separate regions on a single cell. This thickness corresponds to the inner layer of the cell wall only. The distinction between inner and outer layers of the cell wall is described in Fig. S1. At least 60 individual cells per condition and per time were analyzed, and the value and standard deviation reported for each cell wall thickness measurement are therefore issued from at least 180 measurements. Since the height of the section inside one cell could vary and might result in wrong measurement of the wall thickness, we assumed that the largest cells in the section would more or less cut in their center. In that case, the cell wall was perpendicular to the plane of the section and the measured wall thickness was assumed to be meaningful. However, considering the variability of the cell diameter, we checked that, within the population of cells sections that were measured, there was not a clear correlation between the apparent cell size and cell wall thickness (Fig. S2).

## 2.6. Starch structure

Starch granules were extracted from the microalgae after complete cell rupture with a high-pressure homogenizer (HPH, CF2 Cell Disruptor, Constant Systems Ltd., England) at 250 MPa for 5 cycles. The biomass broth was centrifuged for 5 min at 4500 g to pellet the starch granules. The supernatant was discarded and the pellet was resuspended in 1 mL of Percoll (GE Healthcare Bio-Science AB, Sweden). After strong agitation followed by centrifugation, the supernatant of biomass debris and Percoll were removed. The starch pellet was rinsed in pure water and freeze-dried.

Starch granules (2 mg) were disrupted by incubation at 100 °C for 10 min after resuspension in 200 µL DMSO. Amylopectin and amylose were recovered by precipitation with 800 µL ethanol and centrifugation at 10,000 g for 10 min. The polysaccharide pellet was resuspended in 300 µL 10 mM NaOH and applied to a gel permeation chromatography Sepharose CL-2B column (0.5 cm inner diameter, 65 cm length) eluted in 10 mM NaOH to separate amylose and amylopectin which were collected in 300 µL fractions. 80 µL of each fraction were mixed with 20 µL of iodine solution (0.02 % I₂, 0.2 % KI), and the maximal absorbance and the wavelength at the maximum absorbance of the iodine polysaccharide complex ($\lambda_{max}$) was measured as described in [26].

## 2.7. Assessment of cell rupture

Samples of biomass were stored at –20 °C without freeze-drying to preserve the integrity of cell structures. Thawed biomass samples were diluted to approximately 1 mg DW mL⁻¹ in pure water and cells disrupted by a high-pressure homogenizer (HPH, CF2 Cell Disruptor, Constant Systems Ltd., England) at 250 MPa. The biomass broth was retrieved and passed in HPH for 4 additional cycles. Cell concentration was measured in technical triplicate with a particle counter (Multisizer 4 Counter, Beckman Coulter, USA) after each disruption cycle. Particles outside the typical cell size range of *Chlorella vulgaris*, from 2.6 to 10 µm, were excluded. The cell rupture was calculated as the ratio of the difference in present and initial cell concentration to the initial cell concentration.

## 2.8. Statistical analysis

Statistical significance was assessed using the ANOVA test with Prism 9 (Graphpad Software, LLC). When the null hypothesis was rejected ($p < 0.05$), data were further analyzed using Tukey's multiple comparison test. The following results are presented as the average of the technical replicate ($n = 3$), while the error bars account for the standard deviation.
**3. Results and discussion**

*Chlorella vulgaris* was grown under 4 different culture conditions to evaluate the effect of the abiotic factors on starch accumulation. The conditions were: replete medium (N+) considered as the control condition, nitrogen-depleted medium (N-), supra-optimal temperature (called HT for harsh temperature), and red light (RL).

The microalgae were grown under night periods of 8 h, and with high initial DW concentration. Cultures were sampled both before and after the night period over 4 days, and once after 7 days (160 h). Media in the N+ and RL conditions became sulfur-depleted by the end of the day 4 (88 h), as attested by ionic chromatography (Supplementary Material Table S1). Therefore, N+ should not be considered as proper replete-medium control on day 7.

**3.1. Starch accumulation**

Each inducer efficiently triggered starch accumulation, as visible in TEM images (Fig. 2). While the control N+ showed no accumulation of starch granules, the number of starch granules notably increased after 2 days (40 h) of nitrogen deprivation (N-), supra-optimal temperature (HT), or red-light illumination (RL). The number of starch granules within the cells further increased after 4 days (88 h), eventually filling up most of the cellular volume in N- condition. Two types of starch granules were observed: pyrenoidal and stromal starch granules, which had a similar aspect for all starch induction methods.

Fig. 3 illustrates the starch accumulation during growth. All cultures started at 0 h with the same biomass concentration of 0.9 g L⁻¹. In the case of control N+, the biomass DW concentration reached 6 g L⁻¹ after 7 days (160 h). Surprisingly, the DW concentration evolved differently depending on the applied induction method. During the first three days,

<figure><img src="image_6.png" /><figcaption>**Fig. 2.** TEM images of ultrathin sections of resin-embedded *Chlorella vulgaris* cells after 0, 44 and 88 h of starch induction. Starch granules (in white) are visible in the chloroplast, and rapidly increased in number and size from 44 h for N-, HT and RL. Scale bars correspond to 1 µm.</figcaption></figure>
<figure><img src="image_7.png" /><figcaption>**Fig. 3.** Impact of abiotic factors on biomass and carbohydrate accumulation in *Chlorella vulgaris* CCALA924. (A) Biomass dry weight concentration, (B) cell concentration, and (C) carbohydrate content. The carbohydrate content per cell (D) was determined from cell and carbohydrate concentrations. Sulfur was naturally depleted at 88 h in the control culture N+ (dotted vertical line), which explains the high amount of starch observed after 96 h of cultivation in C and D. Shaded regions represent the 8-h night periods.</figcaption></figure>

the DW concentration was the highest in N-, followed by RL, then HT and the control N+ (significant difference, *p* < 0.001) (Fig. 3A). However, after 3 days, the DW concentration kept on increasing regularly only for N+ and RL, whereas the increase in DW concentration slowed down from day 3 for HT, and from day 4 for N-. The DW concentration even decreased after day 4 for HT. Overall, the dry weight biomass productivities over the first four days (88 h) were significantly higher in N- and RL (0.72 g L⁻¹ d⁻¹ and 0.67 g L⁻¹ d⁻¹, respectively) than in N+ (0.58 g L⁻¹ d⁻¹) (*p* < 0.0001).

The starch induction method also had an impact on cellular division (Fig. 3B). In control N+, the cell concentration regularly increased after a short lag phase during day 1. In N-, the cell concentration doubled during the first two days and then completely stopped increasing. More remarkably, HT completely inhibited cell division from the beginning. RL behaved similarly to control N+ (Turkey's multiple comparison test, *p* = 0.92), with significantly close exponential cellular growth rates of 0.59 ± 0.01 d⁻¹ and 0.61 ± 0.03 d⁻¹ after 40 h for RL and N+, respectively (Supplementary Material Table S3).

Although all the cultures started with comparable dry matter at *t* = 0 h, there were variations in the initial carbohydrate content, with HT and RL having higher levels (Fig. 3C). For control culture N+, the carbohydrate content remained stable during 4 days (96 h), between 14.7 ± 3.7 and 25.8 ± 2.0 wt%. After day 4 (96 h), the medium became depleted in sulfur and carbohydrates therefore increased up to 42.8 ± 4.1 wt% on day 7. Nitrogen-depleted cultures (N-) had the fastest and highest accumulation of carbohydrates (significant content difference at

day 4, *p* < 0.001), with a considerable increase in carbohydrate content at day 1, from 14.7 ± 3.7 to 46.5 ± 0.8 wt% in only 16 h. This accumulation continued up to day 4 (88 h) when it reached 69.7 ± 3.5 wt%. From day 4 to 7, the carbohydrate content remained constant. HT cultures followed the same trend of fast and high carbohydrate accumulation. However, from day 4 to 7, the carbohydrate content decreased from 61.7 ± 0.3 to 49.3 ± 1.1 wt%. RL cultures also induced a fast starch accumulation during day 1, with a carbohydrate content increase from 24.2 ± 0.6 to 43.7 ± 0.9 wt% in 16 h. After 16 h, the carbohydrate content remained stable around 45 wt%, even between days 4 and 7 where the control N+ showed an increase in carbohydrate content due to the natural sulfur depletion of the medium. Finally, carbohydrate productivities over 4 days (88 h) in N-, HT, and RL were 0.64, 0.40, and 0.36 g L⁻¹ d⁻¹, respectively (Table 1). These values represent a 5.3-fold, 3.3-fold, and 3-fold increase, respectively, compared to control N+

<caption>**Table 1**</caption>

Biomass, carbohydrate, and starch-equivalent productivities during 4 days (88 h). The detailed content and productivities are reported in Supplementary Material Table S3.

<table><thead><tr><th rowspan="2"></th><th colspan="4">Productivity during 4 days (88 h) (mg L<sup>-1</sup> d<sup>-1</sup>)</th></tr><tr><th>N+</th><th>N-</th><th>HT</th><th>RL</th></tr></thead><tbody><tr><td>Dry weight biomass</td><td>584.3 ± 2.0</td><td>723.0 ± 12.1</td><td>493.4 ± 9.9</td><td>670.2 ± 4.2</td></tr><tr><td>Carbohydrates</td><td>122.8 ± 4.5</td><td>640.8 ± 19.6</td><td>406.2 ± 25.8</td><td>364.5 ± 2.8</td></tr><tr><td>Starch-equivalent</td><td>101.5 ± 4.1</td><td>576.7 ± 17.6</td><td>365.6 ± 23.2</td><td>328.1 ± 2.5</td></tr></tbody></table>
(0.12 g L⁻¹ d⁻¹).

In other works, the *Chlorella vulgaris* carbohydrate productivity reached 0.92 and 0.69 g L⁻¹ d⁻¹ for the CO₂-adapted strain AE10 [12,13] and 0.82 g L⁻¹ day⁻¹ for CCALA924 [5] under various nutrient deprivation and continuous light. The productivities reported in our work are slightly lower (Table 1), but can be explained by the lower light intensity and the 8-h night-period applied. In addition to weight loss and starch consumption due to uncompensated respiration, this 8-h night-period decreased photosynthesis by a third within a 24-h cycle. For instance, in N- condition, the initial carbohydrate productivity reached 970 ± 47 mg L⁻¹ d⁻¹ within the first 16 h, but decreased to 611 ± 23 mg L⁻¹ d⁻¹ over the entire first 24 h, representing a 37 % decrease. For N-, HT, and RL conditions, the decrease in productivity due to the extended 8-h night period ranged between 35 and 50 % over the course of 4 days.

Interestingly, the carbohydrate content per cell was lower for N- than for supra-optimal temperature HT, with maximum values of respectively 23.2 ± 0.7 vs. 32.8 ± 1.8 pg cell⁻¹ at 88 h (Fig. 3D). Meanwhile, the carbohydrate content in RL cells increased only on day 1, from around 10 to 15 pg cell⁻¹, and remained stable until 96 h. Control N+ increased its cellular carbohydrate content during day 1, possibly due to the increased light availability or medium renewal at the start of culture. Then, the carbohydrate content per cell decreased and varied between 8.6 ± 0.8 pg cell⁻¹ at 16 h and 2.5 ± 0.5 pg cell⁻¹ at 96 h. After day 4, and the onset of sulfate depletion, the cellular carbohydrate content increased at day 7 (160 h) for both N+ and RL, reaching respectively 14.7 ± 1.4 and 21.3 ± 4.3 pg cell⁻¹. This increase in cellular carbohydrate content in RL was not concomitant with an overall increase of carbohydrate synthesis, but rather with a general increase of cell dry weight. Indeed, the carbohydrate productivity remained stable between 96 and 160 h, while the exponential growth rate decreased (Supplementary Material Table S3).

These results demonstrated that the three selected inducers efficiently triggered starch production in *Chlorella vulgaris* CCALA924, consistent with several previously published works on other green microalgae [6,9,10]. Nitrogen deprivation resulted in the highest carbohydrate accumulation, with a carbohydrate content remaining high and stable from day 4 to 7. In contrast, the supra-optimal temperature induced a rapid accumulation with the highest cellular content, but the carbohydrate content significantly decreased from day 4. The red-light illumination resulted in a lower but constant carbohydrate content associated with a high biomass production, and tripled its carbohydrate productivity comparing to the control N+ over the first four days.

According to the literature, all known starch induction methods inhibited cell division, including nitrogen starvation and supra-optimal

temperature [6,8]. Here, cell division immediately stopped after being exposed under supra-optimal temperature and after 2 days of nitrogen starvation. However, we observed that cell division continued for the entire 7-day period under red light, in parallel to starch accumulation.

Intriguingly, the natural sulfur depletion occurring on day 4 had different effects on the control and RL cultures. Indeed, from day 4 to 7, the carbohydrate content tripled in N+ but remained stable in RL. This lack of a typical increase in carbohydrate content upon sulfur depletion could suggest that red light may modify the response to nutrient deprivation, which would require further investigations.

## 3.2. Starch structure

The starch granules produced in the four conditions were extracted from the cells harvested after 7 days (160 h) of cultivation. The starch was analyzed by gel permeation chromatography (Fig. 4). Iodine staining of eluted fractions revealed distinct amylopectin and amylose fractions, comparable to those previously found in *Chlorella sorokiniana* and *Chlamydomonas reinhardtii* [27,28], with a $\lambda_{max}$ of the iodine-amylopectin complex ranging from 539 to 560 nm. Qualitatively, starches produced in N+ and RL conditions contain similar very low amounts of amylose, while N- and HT conditions resulted in an apparent increase of this polysaccharide subfraction.

In *Chlamydomonas reinhardtii*, different types of starches are produced depending on growth conditions. During active division, transitory starch mainly accumulates around the pyrenoid which is responsible for CO₂ fixation. Conversely, nitrogen starvation results in a massive accumulation of storage starch in the stroma [29]. Transitory and storage starches differ in amylopectin structure (chain length distribution) and amylose content (higher in storage-like starch) [30]. Hence, the increase in amylose content in N- and HT can be attributed to the accumulation of stromal storage starch granules, as supported by TEM observation (Fig. 2). The absence of a noticeable amylose content increase in RL-produced starch suggests the accumulation of only transitory starch. However, TEM images revealed the presence of stromal granules in RL similar to HT and N-, whereas pyrenoidal starch granules exhibited no sign of hypertrophy, in contrast to the findings in a particular *Chlamydomonas* mutant [29]. Therefore, the stromal starch accumulation coupled with a low amylose content under red-light illumination may indicate that the carbon flux is not massively oriented towards the accumulation of storage compounds. This view is supported by the fact that the cells did not fully redirect their energy towards starch accumulation in response of the absence of blue light, even when nutrient depletion occurred after 4 days (Fig. 3C).

<figure><img src="image_8.png" /><figcaption>**Fig. 4.** Separation of amylopectin and amylose on CL-2B chromatography. Starch was extracted at day 7 (160 h). Maximal absorbance of the polysaccharide/iodine complex (black circles) was measured between 500 and 700 nm for each fraction and the wavelength at maximal absorbance determined ($\lambda_{max}$ in nm, red line). The gel permeation chromatography excluded the high molecular weight amylopectin (1st peak; between 8 and 15 mL of elution); while lower molecular weight amylose was eluted later (2nd peak, after 18 mL of elution). The $\lambda_{max}$ of the iodine-amylopectin complex for each condition are indicated on the corresponding graphs. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)</figcaption></figure>
<figure><img src="image_9.png" /><figcaption>**Fig. 5.** Changes in protein and lipid contents upon starch induction. The dotted vertical line indicates natural sulfur depletion in N+ at 88 h. Shaded area represent the 8-h night periods.</figcaption></figure>

### 3.3. Biomass composition

Apart from carbohydrates, the three main compounds of microalgal biomass are proteins, lipids and pigments. At 0 h, the protein content was approximately 50 wt% for N+, N- and RL, while it could not properly be measured for HT due to analytical problems (Fig. 5A). To perform the comparative calculations, we assumed that the initial protein content for HT was similar to those in other conditions (approximately 50 wt%). After 4 days (88 h), the protein content was $37.7 \pm 1.2$ and $35.0 \pm 1.0$ wt% in N+ and RL, respectively, and only $10.4 \pm 0.4$ and $14.3 \pm 1.0$ wt% for N- and HT. Additionally, protein productivity, calculated taking into account the created biomass together with its protein content after 4 days (88 h) was of $183.7$ and $200.7$ mg L$^{-1}$d$^{-1}$ in N+ and RL respectively, compared to $-28.8$ mg L$^{-1}$d$^{-1}$ for N- and $-12.4$ mg L$^{-1}$d$^{-1}$ for HT (Table 2). Therefore, the protein productivity was in a similar range for red-light illumination and the N+ reference, while it was negatively impacted by nitrogen deprivation and supra-optimal temperature. Nevertheless, the N-content of the N+ reference significantly decreased ($>10$%) during the first 4 days, which was not expected; in parallel, we observed only a slight increase in carbohydrates in the control during the same period.

Regarding nitrogen deprivation, Li et al. observed that the protein content decreased in *Chlorella vulgaris* by at least 2-fold after 7 days [31]. Under supra-optimal temperature, contrary to our results, Zachleder et al. reported a significant protein increase of 8-fold in *Parachlorella kessleri* after 48 h at 40 °C [6]. In the case of red-light illumination, Yuan et al. measured a stable protein content after 6 days in *Chlorella* biofilms [18]. Therefore, in our case, the marked decrease in protein synthesis observed in HT was intriguing. This result indicates a specific sensitivity of *Chlorella vulgaris* CCALA 924 to sustained exposure at supra-optimal temperatures, especially when combined with the alternating patterns of night and day.

Regarding lipid accumulation, *Chlorella vulgaris* CCALA924 did not exhibit any response to the three starch induction methods. At 0 h, the lipid content in biomass was approximately 10 wt% and subsequently remained relatively stable for the control N+ (Fig. 5B). From day 2 (40 h), the lipid content decreased for all induction methods, with the most important decrease occurring for RL, dropping below 5 wt%. Such a quick and high variation could also be linked with analytical problems during sampling preparation (the technical replicates were similar). Moreover, the RL culture produced $10.2$ mg L$^{-1}$d$^{-1}$ lipids over 4 days

<table><caption>**Table 2**
Protein, lipid and chlorophyll productivity during 4 days (88 h).</caption><thead><tr><th rowspan="2"></th><th colspan="4">Productivity during 4 days (88 h) (mg L<sup>-1</sup>d<sup>-1</sup>)</th></tr><tr><th>N+</th><th>N-</th><th>HT</th><th>RL</th></tr></thead><tbody><tr><td>Protein</td><td>183.7 ± 10.3</td><td>-28.8 ± 3.7</td><td>-12.4 ± 8.8</td><td>200.7 ± 9.3</td></tr><tr><td>Lipid</td><td>53.2 ± 4.9</td><td>43.3 ± 6.9</td><td>30.4 ± 3.2</td><td>10.2 ± 4.2</td></tr><tr><td>Total chlorophyll</td><td>28.1 ± 0.4</td><td>-0.3 ± 0.5</td><td>0.0 ± 0.7</td><td>17.9 ± 0.8</td></tr></tbody></table>

(88 h), whereas N+ and N- cultures produced $53.2$ and $43.3$ mg L$^{-1}$d$^{-1}$, respectively (significant difference, *p* < 0.001). Besides, the fatty acid profiles at day 7 (160 h) showed relatively low differences (Supplementary Material Fig. S3).

The decrease in lipid content under red-light illumination was unexpected, as it was shown to remain stable in *Chlorella* biofilms [18]. Moreover, this lower lipid synthesis cannot be only explained by the redirection of available energy towards starch synthesis. Indeed, the lipid productivities of N- and HT were at least 3-fold higher than RL, although concomitantly producing higher amounts of starch. This finding may suggest that blue light is essential to up-regulate lipid synthesis as shown in other microalgae species [32], although the underlying molecular mechanism remains to be determined.

Another important constituent of green microalgal biomass is the pool of photosynthetic pigments, namely chlorophyll a and b and carotenoids. We focused on the quantification of total chlorophyll, calculated as the sum of chlorophyll a and b. The total chlorophyll concentration in cultures remained stable for the beginning for N- and HT, while it increased continuously for N+ and RL cultures (Fig. 6A). Total chlorophyll productivities over 4 days were null or even negative for N- and HT, but were of $28.1 \pm 0.4$ and $17.9 \pm 0.8$ mg L$^{-1}$d$^{-1}$ for N+ and RL cultures, respectively (Table 2). Therefore, the chlorophyll synthesis was significantly reduced in the case of nitrogen starvation and supra-optimal temperature, while it was only slightly reduced by red-light illumination.

The observed decrease in chlorophyll synthesis during nitrogen deprivation and red-light illumination aligns with the literature [18,31]. Regarding supra-optimal temperature conditions, a similar inhibition of chlorophyll synthesis by harsh temperatures was described for higher plants [33].

The analyses of biomass composition have shown that starch was not the sole biomass constituent impacted by the three induction methods. Indeed, the synthesis of proteins and chlorophylls was significantly affected during nitrogen starvation and supra-optimal temperature, with near-to-zero or even negative productivities. Furthermore, the lipid synthesis was also strongly affected, as illustrated by the 5-fold productivity reduction under red-light illumination.

### 3.4. Photosynthetic capacity

The photosynthetic capacity provides direct insights into the ability of microalgae to convert light energy and utilize it to synthesize starch. The maximal quantum yield ($QY_{max}$) represents the fraction of absorbed light energy entering photosystem II (PSII) to produce photochemical energy, with the assumption of full PSII opening [34]. A decrease in $QY_{max}$ is typically associated with damage to the photosynthetic apparatus [35]. In Fig. 7A, $QY_{max}$ is around 0.72 at 0 h for all starch induction methods, indicating a healthy functioning photosynthetic apparatus. However, N- and HT cultures show a severe drop in $QY_{max}$ from $0.69 \pm 0.01$ and $0.70 \pm 0.00$ at 48 h, to $0.15 \pm 0.04$ and $0.06 \pm 0.02$ at 160 h, respectively. In the meantime, N+ and RL cultures maintain a stable $QY_{max}$ throughout the experiment. Interestingly, the evolution of the relative electron transport rate showed a similar pattern with $QY_{max}$ decreasing in N- and HT, while remaining stable in N+ and RL (Fig. S4). The decrease of $QY_{max}$ in N- and HT is expected from the literature, as the decrease in protein synthesis affects the renewal of the essential D1 protein and the chlorophyll-binding protein CP47 within the PSII core [36].

In addition to $QY_{max}$, which provides a broad assessment of the state of photosynthetic apparatus, we further explored the response of the key regulatory mechanisms of non-photochemical quenching (NPQ). NPQ refers to protective regulatory mechanisms leading to the dissipation of excessive absorbed light energy into harmless heat. Therefore, the extent of NPQ activation informs about the level of stress experienced by the photosynthetic apparatus, as well as its capacity to quench excessive energy. Energy partitioning at steady-state during the fluorescence
<figure><img src="image_10.png" /><figcaption>Fig. 6. Evolution of total chlorophyll concentration in the culture and per cell for the different starch induction methods.</figcaption></figure>

<figure><img src="image_11.png" /><figcaption>Fig. 7. Response of the culture photosynthetic capacity to starch induction methods: (A) maximum photochemical efficiency; (B) chlorophyll a/b ratio used as a proxy of the antenna size.</figcaption></figure>

<figure><img src="image_12.png" /><figcaption>Fig. 8. Evolution of absorbed light energy repartition through PSII photochemistry (ΦPSII) (black), fluorescence and constitutive thermal dissipation (Φf,d) (light grey) and regulated thermal dissipation (ΦNPQ) (dark grey) during steady-state photosynthesis.</figcaption></figure>

measurements revealed that NPQ was activated for all induction methods from 0 h (Fig. 8). In N+ and RL, the amount of energy quenched by NPQ (ΦNPQ) remained stable, around 10 %. In N-, ΦNPQ rapidly and gradually increased, which was seemingly associated with an equivalent decrease in PSII energy uptake (ΦPSII). In HT, ΦNPQ remained low and even decreased from day 3 (72 h) to almost zero at 96 h. This absence of NPQ implementation suggests a significant deterioration in cell health. Overall, protective mechanisms of NPQ appeared to be mildly-activated in N+ and RL, highly-activated in N-, and under-activated in HT.

A complementary approach for examining the behavior of the photosynthetic apparatus involves assessing the cell ability to harvest

light via their photocollective pigments. First, the cellular chlorophyll content ranged between 1 and 1.5 pg cell⁻¹ at 0 h for all induction methods (Fig. 6B). For N+ and RL cultures, the amount of chlorophyll per cell remained stable around 1.0 pg cell⁻¹ until day 7 (160 h), in contrast to HT where it rapidly decreased from day 4 (96 h). For N-, the cellular chlorophyll content dropped immediately as a consequence of the initial cell division (Fig. 3) and remained stable around 0.4 pg cell⁻¹.

Furthermore, the antenna size was approached through the proxy of chlorophyll *a* and *b* ratio (Chla/b) as proposed by Friedland et al. [37]. The Chla/b ratio was about 4 at 0 h for every induction method (Fig. 7B). Subsequently, the Chla/b ratio progressively increased for N+ and RL
cultures (i.e. associated with a decrease in antenna size), while it pro-
gressively decreased for N- and HT cultures (i.e. increasing antenna
size). In theory, the antenna size of microalgae should increase as the
culture becomes denser and light availability decreases. However, the
opposite phenomenon intriguingly occurred for the control N+ and
under red-light illumination. In the case of N- and HT, the decrease of
Chla/b was also unexpected, as Chla/b was shown to increase or stay
steady in N-deprived *Chlamydomonas reinhardtii* [38,39]. As a tentative
explanation, it can be postulated that the reaction centers of photosys-
tems (Chla-containing) degraded faster than the light-harvesting com-
plex (Chla- and Chlb-containing).

On the one hand, N- and HT exhibited a similar decrease in their photosynthetic capacities, as evidenced by the simultaneous decrease of QYmax. The NPQ was strongly activated in N- and under-activated in HT, while N- and HT cultures had distinct cellular chlorophyll contents due to differences in cell division. Overall, these results illustrate that the photosynthetic apparatus reacted negatively, albeit differently, to both nitrogen starvation and supra-optimal temperature.

On the other hand, the RL condition completely preserved the photosynthetic and light-harvesting capacities. Parameters such as QYmax, energy distribution at steady state, antenna size and cellular chlorophyll were very similar to the control N+. In conclusion, red-light illumination did not induce any visible damage to the photosynthetic apparatus of microalgae over 7 days.

3.5. *Cell wall thickness and cell rupture*

*Chlorella vulgaris* is known to have a tough and thick cell wall, largely constituted of chitin, a polymer of glucosamine [40,41]. This robustness makes the extraction of intracellular compounds like starch challenging. *Chlorella vulgaris* cells can be efficiently broken by high-energy input devices such as high-pressure homogenizer (HPH). Moreover, the mechanical strength of microbial cells and the thickness of their cell walls have been linked to the ability to withstand mechanical rupture, including HPH [42,43]. Therefore, we investigated the effect of the three starch induction methods on the cell wall thickness and cell rupture.

TEM images of cell walls are shown in Fig. S1. Although the outer
layer of the cell wall was also measured (Fig. S5), only the inner layer

was used for cell wall thickness measurement as most of the thickness
variation was attributed to this layer. At 0 h, violin plots of cell wall
thickness were similar for all conditions, with a mean thickness of
approximately 60 nm (Fig. 9). After 2 days (40 h), the cell walls slightly
thickened in HT and RL cultures (92 ± 2 nm and 79 ± 16 nm, respec-
tively). After 4 days (88 h), the differences in cell wall thickness
increased. The N+ cell wall thickness remained stable (59 ± 15 nm).
Conversely, N-, HT, and RL cells exhibited wall thickening (96 ± 27 nm,
127 ± 30 nm, and 84 ± 20 nm, respectively). Therefore, every starch
induction method, with HT in particular, caused a significant increase in
cell wall thickness compared to control after 4 days.

Interestingly, the cell wall thickening seemed to correlate with the cell division rate (Table S4). As mentioned in Section 3.1, over the first 4 days, the cell concentration regularly increased for N+ and RL, while it increased during one day and then stagnated for N-, and never increased for HT. In parallel, the cell wall thickness of N+ remained low over the 4 days, while it increased from day 2 for N- and from day 0 for HT. Therefore, cell walls tended to thicken when the cellular growth rate was low for N- and HT, as shown in Fig. 3B and illustrated in Supplementary Material Table S4. Intriguingly, RL cell walls thickened despite having a cellular growth rate similar to that of N+.

In the literature, cell wall thickening is often associated with sta-
tionary phase culture, in which cell division is reduced [44,45].
Furthermore, previous studies have established a relationship between
cell division and the reduction of cell wall thickness [46]. Therefore, the
cessation of cell division would result in an average increase in cell wall
thickness in the culture. In our work, the microalga cells exhibited
thicker cell walls when exposed to cell division inhibitors such as supra-
optimal temperature and nitrogen starvation. However, it is noteworthy
that red light appeared to induce cell wall thickening independently of
the cellular growth rate.

To investigate whether cell wall thickness influenced the cell resistance to mechanical disruption, the biomass was disrupted by five subsequent cycles of HPH at 250 MPa (Fig. 9). At 0 h, the cell rupture by HPH showed two distinct efficiencies. For N+, N-, and HT biomass, the cells were rapidly disrupted, with approximately 80 % of cell rupture after the first HPH cycle and approximately 98 % after 3 cycles. For RL cells, the breakage occurred at a slower rate, with approximately 62 ± 3

<figure><img src="image_13.png" /><figcaption>**Fig. 9.** Variations of cell wall thickening over time and corresponding effects on cell rupture. (A) Thickness of cell wall, measured on three regions for at least 60 cells per condition. (B) Cell rupture efficiency according to number of HPH cycles. The *p*-values for the comparisons indicated in the graph are calculated with Prism 9 (Graphpad Software, LLC) and based on ANOVA Dunnett's multiple comparisons test (**, *P* < 0.0021; ***, *P* < 0.0005; ****, *P* < 0.0001, *n* > 160).</figcaption></figure>
% after one cycle and 95 ± 0 % after three cycles. This distinction underscores a slight biological difference between RL inoculum and the inocula used for other conditions, even if the inocula were prepared in conditions as similar as possible. This difference was not emphasized by the other characterization techniques. After 2 days (40 h), cell rupture remained unchanged, except for a reduction in the rate of HT cell rupture that appeared similar to RL cells. After 4 days (88 h), cell rupture after one cycle of HPH became 64 ± 2 % for HT, 70 ± 1 % for N-, 74 ± 1 % for N+ and 78 ± 1 % for RL cells. On the 3rd cycle, the cell rupture efficiencies were similar for all conditions, ranging between 95 and 97 %. In conclusion, cell disruption efficiencies differed slightly between conditions, but no clear pattern emerged over the three-time points analyzed. Thus far, our analysis has not revealed a consistent correlation between the mechanical resistance of cells and the three starch-inducing conditions, nor have we observed consistent differences in mechanical resistance as a result of varying cell wall thicknesses. In conclusion, the three starch induction methods did not cause any extra difficulty for mechanical cell disruption by HPH, and thus ensured the same ease of recovery for the accumulated starch.

Yap et al. observed that differences in cell wall thickness between replete and N-starved *Chlorella vulgaris* did not impact cell rupture efficiency with HPH [43]. In the present work, we extend the validity of this result to two novel starch induction methods, namely supra-optimal temperature and red-light illumination, which increase the cell wall thickness up to 2-fold higher than control.

## 3.6. Application perspectives

### 3.6.1. Advantages and drawbacks of the three tested strategies

In this study, we evaluated the response of the microalga *Chlorella vulgaris* CCALA924 to three starch inducers: nitrogen starvation, supra-optimal temperature, and red-light illumination. The outcome provided valuable insights into the applicability of these starch-inducing methods. A supra-optimal temperature of 39 °C induced important starch accumulation, up to 64.3 wt% carbohydrate content, and starch-equivalent productivity of 0.37 g L⁻¹ d⁻¹ (i.e. 0.41 g L⁻¹ d⁻¹ carbohydrates) in 4 days. Notably, the starch accumulation was initially higher than under nitrogen deprivation during the first two days but considerably decreased afterward. The extensive stress caused by the high temperature was mainly visible through the complete stop in cell division, together with the sharp decrease in maximal quantum yield (QYmax) and highly reduced protein and chlorophyll productivities. Reducing the temperature could potentially mitigate this stress and thereby improve the stability of starch synthesis. Therefore, the supra-optimal temperature is a potentially highly efficient starch induction method for short-term accumulation but requires further optimization of the target temperature.

Nitrogen deprivation triggered the highest and fastest carbohydrate accumulation, with a starch-equivalent productivity of 0.58 g L⁻¹ d⁻¹ over the first 4 days (88 h) (i.e. 0.64 g L⁻¹ d⁻¹ carbohydrates), representing a 5-fold increase compared to the control. The maximal carbohydrate content reached 69.7 wt%. During this period, >80 % of the biomass produced was only carbohydrates, while the production of proteins, lipids and chlorophylls concomitantly decreased. Yet, after 48 h, the photosynthetic capacity of the culture rapidly decreased, eventually reaching a maximal quantum yield (QYmax) below 0.2 after 7 days. In conclusion, the nitrogen starvation method should be considered for short-term starch accumulation strategies.

Red-light illumination induced a 3-fold improvement in starch-equivalent productivity, which notably persisted for 7 days without decrease. During the first 4 days, the starch-equivalent productivity reached 0.33 g L⁻¹ d⁻¹ (i.e. 0.36 mg L⁻¹ d⁻¹ carbohydrates), and the maximal carbohydrate content reached 46.8 wt%. Meanwhile, the protein synthesis slightly increased compared to the control, while lipid productivity decreased by 5-fold. Interestingly, the amylose content in the produced starch was low, contrary to the expected storage-type

starch. Most notably, the dynamics of biomass growth under red light were similar to the control with continuous and regular cell division. In addition, the photosynthetic capacities were fully sustained over 7 days, with no decrease in quantum yield or NPQ mechanisms implementation. Therefore, red-light illumination induced a significant starch accumulation without causing stress to the culture. To our knowledge, this is the first report on a significant starch accumulation concurrent with sustained cell health. In this view, red-light illumination appears to be a novel and gentle way to induce starch accumulation in the microalgae *Chlorella vulgaris*, with interesting characteristics for long-term accumulation.

### 3.6.2. Technical challenges for culture scale-up

The present work opens intriguing perspectives for the large-scale production of starch from microalgae. On the one hand, a continuous culture system illuminated with red LEDs could achieve high starch and protein productivities simultaneously. On the other hand, the nitrogen deprivation induction can be easily implemented in batch cultivation by controlling the initial amount of nitrogen to start the deprivation at a targeted dry weight. Moreover, as suggested by Cheng et al., a two-reactor system could easily be used, with one reactor in replete conditions regularly supplying biomass to the second one operating in several-day batches under nitrogen deprivation [13].

Additionally, the present work raises a compelling scientific and technological interest in exploring the pathways responsible for the induction of starch accumulation in response to red light. A novel kinase signaling cascade has recently been described in *Chlamydomonas reinhardtii* where the blue-light perception mediated by the serine/threonine kinase phototropin is transduced via the Phototropin-Mediated Signaling Kinase 1 to activate gene expression of GAP1, acting as enhancer of starch accumulation [9]. Validating and gaining a profound understanding of these pathways in *Chlorella vulgaris* could potentially result in engineering strains capable of simultaneously sustaining their growth while tripling their starch productivity under solar spectrum.

## 4. Conclusion

In this work, three abiotic starch-inducing methods were compared in terms of carbohydrate productivities, biomass composition, starch structure, photosynthetic capacities, and cell rupture. *Chlorella vulgaris* CCALA924 was confirmed to be a high-starch-producing strain, while its lipid production appeared to be low. All three factors induced a cell wall thickening without any impact on the mechanical cell disruption performed with high-pressure homogenization at 250 MPa, ensuring a similar ease of starch recovery. Overall, nitrogen deprivation and supra-optimal temperature induced a fast but non-sustained starch accumulation and should be preferred for short-term accumulation strategies in batch culture. Despite achieving a lower starch content, red-light illumination induced an enhanced and steady starch production suitable for continuous culture systems. This work opens up novel prospects for the production of microalgal starch, especially for large-scale applications such as bioplastics.

### CRediT authorship contribution statement

A. Six: Writing – original draft, Validation, Methodology, Investigation, Formal analysis, Conceptualization. A. Dimitriades-Lemaire: Resources, Methodology, Investigation. C. Lancelon-Pin: Methodology, Investigation. J.-L. Putaux: Writing – review & editing, Investigation. D. Dauvillée: Writing – review & editing, Investigation. D. Petroutsos: Writing – review & editing, Conceptualization. P. Alvarez Diaz: Resources, Project administration. J.-F. Sassi: Writing – review & editing, Supervision, Project administration, Funding acquisition. Y. Li-Beisson: Writing – review & editing, Supervision. G. Fleury: Writing – review & editing, Supervision, Resources, Methodology, Funding acquisition, Conceptualization.
Declaration of competing interest

The authors declare that they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

Data availability

All data are provided in the Supplementary Material section.

Acknowledgments

The authors thank Florian Delrue for resources and Sophie Mailley for funding acquisition, as well as the NanoBio-ICMG Platform (UAR 2607, Grenoble) for granting access to the electron microscopy facility. This work was partly performed within the framework of SEALIVE (https://sealive.eu/) and Nenu2PHAr (https://nenu2phar.eu/) projects, funded by the European Union's Horizon 2020 Research and Innovation programme under grant agreement #862910 and the Bio Based Industries Joint Undertaking (BBI-JU) under grant agreement #887474, respectively. The BBI-JU receives support from the European Union's Horizon 2020 Research and Innovation programme and the Bio Based Industries Consortium. The PhD grant of A. Six was partly founded by the French Région Provence-Alpes-Côte-d'Azur (Région Sud). The graphical abstract was partly created by BioRender.com.

Appendix A. Supplementary data

Supplementary data to this article can be found online at https://doi.org/10.1016/j.algal.2024.103515.

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