# Laminaria digitata Extract Improved Leaf Meristem Protection Under Drought and Nitrogen Uptake After Rehydration Through Hormesis-Based Chemical Priming in Lolium perenne

Antoine Grandin-courbet, Annette Morvan-bertrand, Marine Dehail, Franck Hennequart, Marie-pascale Prud'Homme

► To cite this version:

Antoine Grandin-courbet, Annette Morvan-bertrand, Marine Dehail, Franck Hennequart, Marie-pascale Prud'Homme. Laminaria digitata Extract Improved Leaf Meristem Protection Under Drought and Nitrogen Uptake After Rehydration Through Hormesis-Based Chemical Priming in Lolium perenne. Plant, Cell and Environment, 2025, 48 (9), pp.6674-6690. 10.1111/pce.15646. hal-05209503

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ORIGINAL ARTICLE

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# Laminaria digitata Extract Improved Leaf Meristem Protection Under Drought and Nitrogen Uptake After Rehydration Through Hormesis-Based Chemical Priming in *Lolium perenne*

Antoine Grandin-Courbet<fnref n="1" />,² | Annette Morvan-Bertrand<fnref n="1" /> | Marine Dehail² | Franck Hennequart² |
Marie-Pascale Prud'homme<fnref n="1" /> ID

**Correspondence:** Marie-Pascale Prud'homme (marie-pascale.prudhomme@unicaen.fr)

**Received:** 13 September 2024 | **Revised:** 23 April 2025 | **Accepted:** 18 May 2025

**Funding:** The work was supported by the University of Caen Normandie and by the company Algaia. The Normandie Council and the European Union funded the work through the research projects AlgoEVA (Valorisation des algues en Normandie pour les secteurs de l'agriculture et de la santé animale, 2018-2021, in the framework of the ERDF-ESF) and BIOSTIM-ALMAP (Intérêt des biostimulants d'origine algale pour la gestion des cultures maraîchères et des prairies dans le contexte du dérèglement climatique et de la transition agroécologique, 2022-2025, in the framework of the EAFRD). Antoine Grandin-Courbet received a PhD grant (2022-2025) from the ANRT (Association Nationale de la Recherche et de la Technologie) (CIFRE 2021/1265).

**Keywords:** algal extract | bioassay | biostimulation indicators | membrane stability | N uptake | perennial ryegrass

## ABSTRACT

Drought is among the most damaging stress for plants, impacting crop yield and grassland sustainability. This study aimed to evaluate the biostimulant effect of an algal extract from *Laminaria digitata* on *Lolium perenne* cultivated in a growth chamber. Leaves were sprayed at different concentrations 7 days before stopping irrigation. This priming period was followed by fourteen days of drought and ten days of recovery. Algal extract supplied at 2 and 5 L.ha⁻<fnref n="1" /> stimulated nitrogen uptake during recovery, while higher doses were deleterious. During drought, algal extract 2 L.ha⁻<fnref n="1" /> increased water content in leaves and shoot 0-3 cm housing the leaf meristems. The improvement in water content arose from the smaller decline in leaf relative water content (RWC), suggesting better osmotic adjustment. Cell membrane stability was less impaired during drought and quickly returned to pre-drought levels during recovery, indicating better membrane protection. The higher fructan content may contribute to osmotic adjustment and membrane protection. The results show that algal extract improved leaf meristem protection under drought and N uptake after rehydration through hormesis-based chemical priming. The treatment limited sucrose accumulation during drought, so that sucrose content can be used as an indicator of biostimulation together with RWC and cell membrane stability.

## 1 | Introduction

The combination of land-use and climate changes due to human activities leads to modifications of production conditions in agriculture. Plants are subject to more intense and frequent abiotic stress such as drought. Drought is one of the most damaging stress for plants (Farooq et al. 2009; Seleiman

et al. 2021). This stress affects plant growth and development with an impact on crop yield and grassland sustainability (Bray et al. 2000; IPCC 2021). Grasslands represent an important land use in Europe and contribute not only to livestock feed but also provide many ecosystem services (Smit et al. 2008; Dumont et al. 2019). Drought has been shown to lead to a decline in grassland productivity (Felton et al. 2019; Carroll et al. 2021),

This is an open access article under the terms of the Creative Commons Attribution-NonCommercial License, which permits use, distribution and reproduction in any medium, provided the original work is properly cited and is not used for commercial purposes.

© 2025 The Author(s). *Plant, Cell & Environment* published by John Wiley & Sons Ltd.

<footnote n="1">¹Normandie Université, UNICAEN, INRAE, EVA Ecophysiologie Végétale, Agronomie & nutritions NCS, Caen, France | ²Algaia SAS, Saint-Lô, France</footnote>
illustrated in particular by a net loss of primary productivity, which depends on soil composition (Buttler et al. 2019), species richness and diversity (Craven et al. 2016) and timing of stress (Craine et al. 2012). Drought can also have an impact on the long-term functioning of the ecosystem (Haddad et al. 2002).

At the plant level, drought is known to cause stomatal closure, leaf temperature increase and leaf turgescence decline (Farooq et al. 2009). To limit the effect of drought, plants have developed different resistance mechanisms at the tissue, cellular and molecular levels (Volaire 2018). Among them, the accumulation of soluble compounds such as soluble carbohydrates allow plants to maintain osmotic potential in cells and therefore limit water dehydration (Farooq et al. 2009). Another key component of drought resistance is the protection of membrane integrity. Indeed, under water stress induced by cold or drought, the cytoplasmic content becomes increasingly viscous and this enhances the risk of molecule denaturation, membrane destabilization and membrane fusion (Hoekstra et al. 2001). To ensure the stability of the membranes, some molecules such as sucrose are able to interact with phosphate in head groups of phospholipids (Strauss and Hauser 1986).

Among soluble carbohydrates involved in drought resistance, fructans are particularly important in grasslands. Indeed, these soluble fructose polymers are synthesized from sucrose by 15% of angiosperms (Hendry 1993) including perennial ryegrass (*Lolium perenne*), one of the main species of temperate grasslands in which they represent the main form of carbohydrate storage (Chatterton et al. 1989). Fructans and the regulation of their degree of polymerization by fine enzyme activity tuning may contribute to cell integrity by both osmotic adjustment and membrane protection (Hincha et al. 2007; Valluru et al. 2008). Fructans are able to insert between the polar headgroups of phospholipids, thus maintaining the stability of membranes (Vereyken et al. 2003; Hincha et al. 2007; Livingston et al. 2009).

Perennial ryegrass dominates temperate grasslands and is widely cultivated for forage due to its high growth rate under conditions of high nutrients and water availability, and its high nutritive value for livestock, but it is not well adapted to hot summers and drought (Wilkins and Humphreys 2003). At vegetative stage, a plant of ryegrass is composed of several tillers. One tiller includes mature and elongating leaves. The leaf meristem, made of dividing and elongating cells, is located between 0 and 3 cm from the base of elongating leaves which are surrounded by the sheaths of mature leaves (Morvan-Bertrand et al. 2001). When plants are defoliated by mowing or grazing, only elongating leaves continue to grow through the activity of their meristems. Moreover, leaf meristems survive water stress better than most mature leaf tissues so that they represent key surviving organs for plant recovery after drought (Volaire et al. 2014; Barkaoui and Volaire 2023; Van Hautegem et al. 2025). Their dehydration resistance is at least partially mediated by high concentrations of water-soluble carbohydrates (WSC), especially fructans, allowing osmotic adjustment and membrane protection (Zwicke et al. 2015; Volaire et al. 2020). Breeding efforts have led to the development of perennial ryegrass cultivars with increased levels of WSC to improve protein digestion efficiency and milk production (Miller et al. 2001). Given the role of WSC, and fructans in particular, in drought

resistance, increasing WSC content could alleviate the negative impact of drought. However, as the expression of WSC traits is strongly controlled by genotype and environment, evaluation of the potential of high-WSC cultivars to improve drought resistance has been inconclusive (Robins and Alan Lovatt 2016). Summer productivity can be improved by irrigation, but this management strategy can entail high economic and environmental costs (Rogers et al. 2019). A new approach to reduce stress-induced yield limitations is the use of biostimulants.

According to the EU fertilizer regulation 2019/1009, a plant biostimulant is “a product stimulating plant nutrition processes independently of the product’s nutrient content with the aim of improving one or more of the following characteristics of the plant or the plant rhizosphere: nutrient use efficiency, tolerance to abiotic stress, quality traits, availability of confined nutrients in soil or rhizosphere” (Ricci et al. 2019). A scientific consensus agreed around five main types of products: humic substances, plant extracts including those of seaweed, amino acids and protein derivatives, inorganic compounds and microorganisms (du Jardin 2015; La Torre et al. 2016; Yakhin et al. 2017). The definition of the EU comes with the idea of harmonizing regulations and making sure that the biostimulants available in the market have proven their effectiveness. Numerous compounds, including algal extracts, are able to increase plant growth and development (Franzoni et al. 2022). The use of algal extract, in synergy with organic and inorganic fertilisers, can improve nutrient uptake and confer abiotic stress resilience, thus reducing costly synthetic inputs and indirectly environmental impacts by limiting leachates (Kergosien et al. 2023). In addition, as macroalgae grow in the sea and therefore do not compete with crops, and do not require water nor fertilizers, macroalgal extracts are developing rapidly on the biostimulant market (Kergosien et al. 2023). The effectiveness of various algal extracts in alleviating drought stress has been repeatedly demonstrated (Ali et al. 2021; Deolu-Ajayi et al. 2022), such as red algae *Kappaphycus alvarezii* extract on wheat (Patel et al. 2018), microalgae *Amphora ovalis* extract on onion (Almaroai and Eissa 2020), brown algae *Ascophyllum nodosum* extract on pepper (Dalal et al. 2019) and tomato (Goñi et al. 2018). *Laminaria digitata* extracts and in particular their laminarans are known for their protective effect against biotic stress (Klarzynski et al. 2000; Aziz et al. 2003; Agarwal et al. 2021), while their protective effect against abiotic stress remains largely unknown. Improved drought stress resistance has been obtained with mixtures containing *Laminaria* extract, in tomato (*L. digitata* with *A. nodosum* and yeast, Campobenedetto et al. 2021; *L. digitata* with *A. nodosum* and humic and fulvic acids, Liava et al. 2023), olive tree (*L. digitata* with *A. nodosum*, Graziani et al. 2022) and mint (*L. digitata* with *A. nodosum*, and vegetable proteins and amino acids or humic and fulvic acids, Chaski et al. 2023). However, to the best of our knowledge, only one study has reported the positive effect of an extract exclusively based on *L. digitata*. It was against heat and salinity in *Arabidopsis thaliana* (Wu et al. 2016). The drought protective effect of pure *L. digitata* extract has never been reported.

The objective of this study was to assess the protective effect of *L. digitata* extract to improve the drought resistance and resilience of *L. perenne*. For this purpose, plants were grown in a phytotronic chamber to tightly control environmental
conditions. A protocol made of three successive periods was designed, (i) seven days corresponding to the priming period following the treatment with the algal extract supposed to prepare the plants for the stress to come, (ii) fourteen days without irrigation leading to a gradual decrease in soil water content, (iii) ten days of recovery under irrigation. To assess the overall effect of the algal extract on drought resistance and resilience, we measured the biomass and water content of the different plant compartments at the end of each period. To assess more precisely the contribution of leaf meristem protection to drought resistance, we monitored leaf relative water content (RWC) as a proxy for osmotic adjustment (Matin et al. 1989), relative electrolyte retention capacity (RERC) as a proxy for membrane stability (Volaire 2002), and the content of water-soluble carbohydrate (WSC) including fructans, given their involvement in cell protection in grasses. In addition, to evaluate the contribution of mineral nutrition preservation to drought resilience, we monitored nitrogen uptake during recovery. The experiment was repeated as an independent assay to evaluate the feasibility of the protocol as a drought resistance bioassay and to define reliable and reproducible indicators of biostimulation.

## 2 | Materials and Methods

### 2.1 | Plant Growth Conditions and Treatments

Perennial ryegrass (Lolium perenne var. Delika) seeds were sown on a 1.4 mm mesh placed in contact to deionized water at 20°C in the dark for 3 days in a phytotronic chamber (FITOCLIMA D1200, ARALAB, Rio de Mouro, Portugal). The seedlings were then exposed to light with a 16 h photoperiod with photosynthetically active radiation of 400 µmol.m⁻².s⁻¹ measured with a light meter (LI-250, LI-COR, Lincoln, NE, USA), a temperature of 20°C during the day and 18°C at night and a relative humidity of 60%. Five days after sowing, the seedlings were transplanted into 2 L pots (11 × 11 × 20 cm). The pots are divided into four individual compartments (physically separated by a plastic cross system), each forming a replicate (5.5 × 5.5 × 20 cm for each replicate). Each pot contained a mixture of 1:1 (v:v) of sand (0–2 mm) and perlite (nr. 3; grain size 1–6 mm).

Twenty-five seedlings were transplanted for each replicate, i.e. 100 plants per pot. The pots were placed in a 60 × 40 cm tray (12 pots per tray), with drainage at the bottom of each pot, and kept in the same phytotronic chamber for the rest of the experiment with the same environmental conditions. The nutrient solution was delivered in four doses of 60 mL per 24 h at 3:00, 9:00, 15:00 and 21:00. The nutrient solution contained 1 mM K₂SO₄, 1 mM NH₄NO₃, 0.4 mM KH₂PO₄, 0.15 mM K₂HPO₄, 3 mM CaCl₂·H₂O, 0.5 mM MgSO₄·7H₂O, 0.2 mM EDTA (C₁₀H₁₆N₂O₈), NaFe·3H₂O, 10 µM H₃BO₃, 5 µM MnSO₄·H₂O, 3 µM ZnSO₄·7H₂O, 0.7 µM CuSO₄·5H₂O, 0.7 µM (NH₄)₆Mo₇O₂₄ and 0.1 µM CoCl₂.

The plants were grown for 33 days under these conditions. On day 33, the plants were cut at 5 cm from the ground. On day 40 (D0), the plants were cut at 10 cm from the ground and algal extract (or water for the control plants) was sprayed on the leaves. Seven days after the spraying (end of priming period, D7), the plants were cut at 5 cm from the ground and irrigation was stopped for fourteen days excepted for the irrigated control plants. After

fourteen days without irrigation (end of drought period, D21), the plants were cut at 5 cm from the ground and were re-irrigated with the same nutrient solution except that N was labelled using ¹⁵NH₄¹⁵NO₃ (with a ¹⁵N abundance of 4.06 atom % in the nutrient solution) for ten days (end of recovery period, D31). Camera monitoring and pot weighing were performed every two to 3 days throughout the experiment (Figure 1A).

The experiment was repeated once similarly as an independent assay. The first assay was named Bio-A and the corresponding results are presented in the main Figures 2–7 and in supplemental Figures S1–S3. The second assay was named Bio-B and the corresponding results are presented in Figure S4.

### 2.2 | Algal Extract Treatment

Leaves were sprayed with a commercial brown seaweed liquid extract from Algaia®, based on Laminaria digitata. The characteristics of the algal extract are reported in Borlongan et al. (2024). This extract consists of a mix of soluble compounds naturally present in brown seaweeds, in particular alginates, fucoidans, laminarans, mannitol, amino acids and mineral salts. The extraction takes place in controlled temperature and does not involve chemical addition.

Extract solutions were sprayed using 30 mL European standard medical bottle equipped with GCMI 28/410 bottle neck and sprayer. Liquid algal extract was diluted with pure water to supply the equivalent of 0.5, 2, 5, 10 and 15 L·ha⁻¹ (Bio-A experiment) or the equivalent of 0.5, 2 and 5 L·ha⁻¹ (Bio-B experiment). Specifically, each pot having a surface area of 121 cm², 0.605 µL (for 0.5 L·ha⁻¹), 2.4 µL (for 2 L·ha⁻¹), 6.05 µL (for 5 L·ha⁻¹), 12.1 µL (for 10 L·ha⁻¹) and 18.15 µL (for 15 L·ha⁻¹) of extract were diluted into a total volume of 400 µL with ultrapure water (≥ 18.2 MΩ·cm⁻¹) and supplied with four sprays of 100 µL for each dose. These dilutions are of the same order as those used in the field since the commercial recommendation is to dilute 2 L of extract in a tank of 200–400 L.

### 2.3 | Sampling

At the end of each period (D7, D21 and D31), the plants were sampled for analysis. For the determination of relative water content (RWC), the last fully developed leaf blade of the main tiller was sampled from ten different plants of each replicate. For the other analyses, the shoots were cut at 5 cm from the ground (Figure 1B) and the above part was called shoot > 5 cm. The below part of shoots (shoot 0–5 cm) was separated from the roots. After removing the senescent tissues, each shoot 0–5 cm sample was cut at 3 cm from the tiller base leading to two subsamples, the 0–3 cm basal part (shoot 0–3 cm) and the 3–5 cm upper part. The shoot 0–3 cm consisted of the base of elongating leaves, corresponding to leaf meristems (Morvan-Bertrand et al. 2001), surrounded by mature leaf sheaths. The rest of the aerial parts (the 3–5 cm upper part pooled with the shoot 0–5 cm senescent tissues) was called the shoot 3–5 cm (Figure 1B). The roots were harvested and cleaned before being weighed. Sand and perlite particles were removed from the roots by two baths of deionised water.
<figure><img src="image_4.png" /><figcaption>**FIGURE 1** | (A) Experimental design. (X) Number of photography sessions (camera monitoring) together with pot weighing (every 2, 3 days) during plant growth, priming, drought and recovery periods. Λ Measurement of parameters (biomass, water content, relative water content, relative electrolyte retention capacity, carbohydrate content, nitrogen content). (B) Description of a tiller of perennial ryegrass (Lolium perenne).</figcaption></figure>

## 2.4 | Total Nitrogen and N Uptake

Total N ($^{15}$N + $^{14}$N) content (mg.g$^{-1}$ DM) and $^{15}$N abundance ($^{15}$N/$^{15}$N + $^{14}$N)) were quantified in each plant tissue (shoot > 5 cm, shoot 3–5 cm, shoot 0–3 cm, root) and in the nutrient solution. The dried tissues were ground with a ball mill to a fine powder. An aliquot was weighed and placed in a tin cap. Nutrient solution (100 µL) was deposited on Chromosorb® in a tin cap. Samples were analysed in an isotope ratio mass spectrometer (IR-MS, Isoprime, GV Instruments, Manchester, United Kingdom) connected to a C/N analyser (EA3000, Euro Vector, Milan, Italy) by PLATIN' (Plateau d'Isotopie de Normandie, Caen, France).

The N taken up during the recovery period were calculated in mg.plant$^{-1}$ and in mg.g$^{-1}$ DM as follows for each tissue:

$$ 
\begin{aligned}
& N \text{ uptake (mg.plant}^{-1}\text{)} \\
& = \frac{(\text{tissue } 15\text{N abundance} - \text{pre} - \text{labelling } 15\text{N abundance})}{(\text{nutrient solution } 15\text{N abundance} - \text{pre} - \text{labelling } 15\text{N abundance})} \\
& \times \text{total } N_{\text{tissue}} (\text{mg.plant}^{-1})
\end{aligned}
$$

$$ 
\begin{aligned}
& N \text{ uptake (mg.g}^{-1}\text{ DM)} \\
& = \frac{(\text{tissue } 15\text{N abundance} - \text{pre} - \text{labelling } 15\text{N abundance})}{(\text{nutrient solution } 15\text{N abundance} - \text{pre} - \text{labelling } 15\text{N abundance})} \\
& \times \text{total } N_{\text{tissue}} (\text{mg.g}^{-1}\text{DM})
\end{aligned}
$$

The proportion of N taken up during the recovery period was calculated as follows:

$$ \text{Proportion of } N \text{ uptake} = N \text{ uptake}/\text{total } N. $$

## 2.5 | Water Content and Relative Water Content (RWC)

Determination of the water content was done by weighing the fresh mass of the tissue at harvest. The samples were then dried for 48 h at 60°C in an oven. A new weighing gave the dry mass of the tissue. For the determination of relative water content (RWC), the last fully developed leaf blade of ten tillers were weighed (fresh mass) and placed on absorbent paper soaked in
pure water for 24 h at 4°C. After this time, the blades were quickly blotted and weighed (turgid mass). The blades were then dried for 48 h at 60°C and weighed (dry mass). The RWC (g·g⁻¹) was calculated as the ratio of the mass of water in the freshly sampled leaf to the mass of water in the turgid leaves:

$$RWC = (fresh mass - dry mass)/(turgid mass - dry mass).$$

## 2.6 | Relative Electrolyte Retention Capacity (RERC)

As an indicator of membrane stability, the relative electrolyte retention capacity (RERC; adapted from Volaire (2002)) was determined on shoot 0–3 cm (150 mg for irrigated plants or 50 mg for plants sampled at the end of the drought). Tissues were cut into 5 mm long pieces and washed three times with pure water to remove surface electrolytes. The 5-mm long pieces were incubated in 60 mL of pure water. After 24 h of incubation at room temperature in the dark, the electrolyte conductivity of the solution (EC1) was measured with a conductivity meter (Pure Water Tester HI-98308; Hanna Instruments, Smithfield, RI, USA). Then, the solution containing the 5-mm long pieces was boiled at 100°C for 1 h to release all the electrolytes present inside the cells. When the temperature of the solution returned to room temperature, the electrolytic conductivity was measured again (EC2). The RERC was calculated as:

$$RERC = (EC2 - EC1)/EC2.$$

A stable membrane prevents electrolyte leakage and therefore leads to high electrolyte retention indicated by a low EC1 and a high RERC close to 1. On the contrary, an unstable membrane does not retain electrolytes, leading to a high EC1 and a low RERC. Thus, RERC is used as an indicator of cell membrane stability (Blum and Ebercon 1981; Volaire 2002).

## 2.7 | Water-Soluble Carbohydrate (WSC) Content

Water-soluble carbohydrate content was determined on the shoot 0–3 cm. Tissues were frozen in liquid nitrogen before freeze-drying. Freeze-dried tissues were ground with a ball mill and 50 mg of powder were weighed in a 2 mL-microtube. One mL of pure water containing 1 g·L⁻¹ mannitol (internal standard) was added to the powder. The microtube was mixed and incubated 15 min at 80°C. After the first water extraction, the sample was centrifuged 3 min at 15,000 × g. The supernatant was preserved and 1 mL of pure water was added to the pellet. The tube content was mixed and incubated 15 min at 60°C. After this second extraction, the sample was centrifuged 3 min at 20,000 × g. The supernatant was preserved and pooled with the first supernatant. The extraction was repeated a third time. The three supernatants were pooled and placed few hours at −80°C before freeze-drying. The freeze-dried sample was dissolved in 1 mL of pure water. For analysis of water-soluble carbohydrates, 100 µL of water-soluble extract were passed through mini-columns (Mobicols, MoBITec, Göttingen, Germany) packed, from bottom to top, with cotton ball, 250 µL of cation exchange resin in H⁺-form (Dowex 50 W X8-400, Sigma, Saint-Louis, MO,

USA) to remove cationic compounds, 80 µL of polyvinylpolypyrrolidon (PVPP, Sigma-Aldrich, St. Louis, MO, USA) to remove lipids, pigments and phenolic compounds and 150 µL of anion exchange resin in COO⁻-form (Dowex 1 × 8, Fluka, Buchs, Suisse) to remove anionic compounds. To collect the purified sample, the mini-column was successively centrifugated 1 min at 300 rpm, 1 min at 1000 rpm, 2 min at 10,000 rpm. Fructans, sucrose, glucose, and fructose were analyzed by high performance liquid chromatography (HPLC) on a cation exchange column (Sugar-PAK I, 300 × 6.5 mm, Millipore Waters, Milford, MA, USA) eluted at 0.5 mL·min⁻¹ and 85°C with 0.1 mM of Ca-EDTA in water, using mannitol as an internal standard and a refractive index detector (MilliporeWaters, Milford, MA, USA).

## 2.8 | Statistics

Statistical analyses were performed using Microsoft Office Professional Plus 2019 Excel and R software (version 4.0.3). Normality of data distribution (Shapiro-Wilk's test, 95%) and variance homogeneity (Bartlett's test, 95%) and were checked with R. When data were not normally distributed and/or the variances were not homogeneous between conditions, the Box-Cox transformation was performed using the 'powerTransform' function from the 'car' package in R (Fox and Weisberg 2024). At the end of priming (D7), the values obtained for plants treated with each dose of algal extract were compared to the ones obtained for control plants treated with water by the Dunnett's test with the 'DunnettTest' function from the 'DescTools' package in R (Signorell 2024) in using control plants as the reference. At the end of drought (D21) and recovery (D31), the values obtained for well-watered control plants were compared to the values obtained for drought-stressed control plants. When data were normally distributed, Student's tests were performed with Excel (t.test function), either with homoscedastic or heteroscedastic t-tests, when variances were homogeneous or not, respectively. When data were not normally distributed Wilcoxon's test was performed with R. At the end of drought (D21) and recovery (D31), the values obtained for drought-stressed control plants were compared to the values obtained for plants treated with each dose of algal extract by the Dunnett's test with the 'DunnettTest' function from the 'DescTools' package in R (Signorell 2024) in using the drought-stressed control plants as reference. To graphically visualize the hormetic curve on plotted data, nonlinear regression was applied to the data with the drc R package (Ritz et al. 2015) using the Brain-Cousens model with five parameters (BC.5 function) as proposed by Brain and Cousens (1989). A lack-of-fit F-test (modelFit in drc) was used to know if the proposed statistical model fits the design points well (Knezevic et al. 2007). The no-effect hypothesis was tested by a χ² test (noEffect in drc) for the comparison of the proposed statistical model with the simple regression model with slope 0 (a horizontal regression line corresponding to no dose effect) (Ritz et al. 2015).

# 3 | Results

## 3.1 | Plant Biomass

At the end of the priming period of 7 days (D7), the biomass of shoot > 5 cm was unaffected by the foliar spray with *Laminaria digitata* extract (Figure 2A). The same results were observed for
<figure><img src="image_5.png" /><figcaption>**FIGURE 2** | Legend on next page.</figcaption></figure>
the biomass of other tissues (Figure S1). After 14 days without irrigation (D21), all plants showed visual leaf degradation compared to the well-watered control and the algal extract had no impact on biomass of shoot > 5 cm (Figure 2B) and of other tissues (Figure S1) compared to the drought-stressed control. In contrast, effects of algal extract were observed at the end of the recovery period (D31) with a significant dose effect and maximum regrowth between 2 and 5 L.ha⁻¹ (Figure S5D, Table S1). Plants sprayed with 2 L.ha⁻¹ algal extract tended to regrow slightly better than the drought-stressed control plants while regrowth of plants sprayed with 15 L.ha⁻¹ algal extract was much lower than that of the drought-stressed control plants and the plants treated with all other doses (Figure 2C).

## 3.2 | N Uptake

The N taken up during the recovery period was determined in different compartments (shoot > 5 cm, shoot 0–5 cm, root) and in the whole plant using ¹⁵N labelling. Results were expressed in mg.plant⁻¹, mg.g⁻¹ DM and in proportion of total N (Figure 3).

Plants sprayed at 2, 5 and 10 L.ha⁻¹ tended to absorb more N than the drought-stressed control plants (Figure 3A) with a higher amount of N uptake in shoot 0–5 cm for plants treated at 10 L.ha⁻¹ (Figure 3G). This N was mainly found in shoot (Figure 3D, 3G), representing nearly 60% of total N in shoot 0–5 cm (Figure 3I) and 80% in shoot > 5 cm (Figure 3F). In contrast, plants sprayed at 15 L.ha⁻¹ absorbed less N than the drought-stressed control plants (Figure 3A, 3D, 3G, 3J). Consequently total N level (mg.g⁻¹ DM) at the end of recovery was higher in plants sprayed at 2 L.ha⁻¹ (*p* < 0.1) and 5 L.ha⁻¹ (*p* < 0.05) and lower in plants sprayed at 15 L.ha⁻¹ (*p* < 0.05) than in the drought-stressed control plants at the whole plant level (Figure 3B) as well as in total shoot (Figure S2B). This pattern was also found in shoot 0–5 cm (Figure 3H) and root (Figure 3K).

## 3.3 | Water Content

At the end of the priming period, the water content in different shoot compartments (Figures 4A, 4D, 4G, S3D) and roots (Figure S3M) was unaffected by the algal extract and was between 85% and 92%. At the end of the drought period, the water content in drought-stressed control plants was much lower (25% in the whole plant) than in well-watered plants (85%) (Figures 4B, 4E, 4H, S3E, S3H). The water content in tissues of plants sprayed at 2 and 5 L.ha⁻¹ decreased less than in drought-stressed control plants. The effect was particularly pronounced with the 2 L.ha⁻¹ treatment. Consequently, in these plants, the water content was much higher than in the drought-stressed control plants at the end of the drought period and was between 35% and 65% (Figures 4B, 4E, 4H, S3B, S3E,

S3H, S3K, S3N). The same trend was obtained in an independent assay (Figure S4A, S4B). At the end of the recovery period, the water content of all tissues was again greater than 80% (Figure 4C, 4F, 4I) in all conditions, except for plants treated with 15 L.ha⁻¹ in which the water content did not reach the pre-drought value, particularly in shoot tissues (Figures 4, S3).

## 3.4 | Assessment of Osmotic Adjustment by the Relative Water Content (RWC)

The last mature leaf blades were used to assess a possible osmotic adjustment through the relative water content (RWC) measurement. At the end of the priming period, RWC was unaffected by the algal extract and was between 0.89 and 0.94 g.g⁻¹ (Figure 5A). During the drought period, RWC strongly decreased in drought-stressed plants to 0.15 g.g⁻¹ (Figure 5B). This decrease was much less pronounced in plants treated with 2 L.ha⁻¹ than in drought-stressed control plants (0.40 g.g⁻¹; *p* < 0.001). By contrast, the decrease was more pronounced in plants treated with 10 L.ha⁻¹ compared to drought-stressed control plants (0.10 g.g⁻¹; *p* < 0.1). The same trend was obtained in an independent experiment (Figure S4C). During the recovery period, RWC increased in all treated plants and reached higher level in plants sprayed at 0.5 L.ha⁻¹ (*p* < 0.01) than in drought-stressed control plants (Figure 5C).

## 3.5 | Assessment of Membrane Stability by the Relative Electrolyte Retention Capacity (RERC)

The relative electrolyte retention capacity (RERC) was measured to assess membranes stability and the chosen tissue was shoot 0–3 cm, since it contains the leaf meristems. RERC did not change during the priming period and was around 0.7 (Figure 6A). At the end of the drought period, RERC of plants subjected to drought decreased drastically to 0.09 (Figure 6B). This decrease was less pronounced in plants treated with 2 L.ha⁻¹ which maintained this value at 0.23 (Figure 6B). The same trend was obtained in an independent assay (Figure S4D). During the recovery period, RERC increased in all plants subjected to drought (Figure 6C). Compared to the drought-stressed control plants, the increase tended to be higher in plants sprayed at 2 and 5 L.ha⁻¹ in which RERC reached nearly 0.60 at the end of the recovery period. By contrast, the increase was significantly lower in plants sprayed at 15 L.ha⁻¹ in which RERC reached only 0.30 (*p* < 0.1).

## 3.6 | Water Soluble Carbohydrate (WSC) Content

In control plants, at the end of the priming period, the main WSC in shoot 0–3 cm were fructans (113 mg.g⁻¹ DM), followed

**FIGURE 2** | Photographs of plants and rate of biomass production of shoot > 5 cm at the end of priming D7 (A), drought D21 (B) and recovery D31 (C). Dotted line represents the value of well-watered control plants (WW) at the end of priming and drought-stressed control plants (D) at the end of drought and recovery. Each bar represents mean ± standard error (n = 4). Differences between each dose with WW at the end of priming, and differences between each dose with D at the end of drought and recovery, were analyzed by Dunnett's test (†*p* < 0.1; \**(*p* < 0.05; \*\*\*(*p* < 0.01; \*\*\*\*(*p* < 0.001)). Differences between WW and D at the end of drought and recovery were analyzed by Student's or Wilcoxon's test (†*p* < 0.1; \*\*(*p* < 0.05; \*\*\*\*(*p* < 0.01; \*\*\*\*\*(*p* < 0.001) above the line drawn between WW and D). [Color figure can be viewed at wileyonlinelibrary.com]
<figure><img src="image_6.png" /><figcaption>**FIGURE 3** | N uptake during recovery in mg.plant⁻¹ (A, D, G, J), in mg.g⁻¹ DM (hatched bar in B, E, H, K) and in proportion of total N (C, F, I, L) in whole plant (A–C), shoot > 5 cm (D–F), shoot 0–5 cm (G–I), root (J–L). Solid bars in (B, E, H, K) indicate total N (mg.g⁻¹ DM) at the end of recovery period (D31). Dotted line represents the value of drought-stressed control plants (D). Each bar represents mean ± standard error (n = 4; except n = 3 for 10 L. ha⁻¹ treated plants). Differences between each dose with D at the end of recovery were analyzed by Dunnett's test (†p < 0.1; *p < 0.05; **p < 0.01; ***p < 0.001). Differences between WW and D at the end of recovery were analyzed by Student's or Wilcoxon's test (†p < 0.1; *p < 0.05; **p < 0.01; ***p < 0.001 above the line drawn between WW and D).</figcaption></figure>
<figure><img src="image_7.png" /><figcaption>**FIGURE 4** | Water content of ten last mature leaf blades at the end of priming (A), drought (B) and recovery (C), of shoot > 5 cm at the end of priming (D), drought (E) and recovery (F), and shoot 0–3 cm (containing the leaf meristems) at the end of priming (G), drought (H) and recovery (I). Dotted line represents the value of well-watered control plants (WW) at the end of priming and drought-stressed control plants (D) at the end of drought and recovery. Each bar represents mean ± standard error (n = 4). Differences between each dose with WW at the end of priming, and differences between each dose with D at the end of drought and recovery, were analyzed by Dunnett's test (†*p* < 0.1; *p* < 0.05; ***p* < 0.01; ***p* < 0.001). Differences between WW and D at the end of drought and recovery were analyzed by Student's or Wilcoxon's test (†*p* < 0.1; *p* < 0.05; **p* < 0.01; ***p* < 0.001 above the line drawn between WW and D).</figcaption></figure>

by fructose (22 mg.g⁻¹ DM), sucrose (14 mg.g⁻¹ DM) and glucose (14 mg.g⁻¹ DM) (Figure 7A, 7D, 7G, 7J). Treatment of plants with algal extract changed the WSC contents. The content of fructans tended to be lower than in control plants with significant difference at 0.5 and 5 L.ha⁻¹ (*p* < 0.05; 73 mg.g⁻¹ DM; Figure 7A). By contrast, sucrose, glucose and fructose contents tended to be higher than in control plants, except with 0.5 L.ha⁻¹, with significant differences for sucrose at 10 and 15 L.ha⁻¹ (20 mg.g⁻¹ DM) (Figures 7D, 7G, 7J).

In response to drought, two patterns of change were observed depending on the carbohydrate. On the one hand, the contents of fructans, glucose and fructose were lower in drought-stressed (91, 10, 17 mg.g⁻¹ DM, respectively) than in well-watered control plants (233, 14, 22 mg.g⁻¹ DM, respectively) (Figure 7B, 7H, 7K). On the other hand, sucrose content was much higher in drought-stressed (87 mg.g⁻¹ DM) than in well-watered control plants (19 mg.g⁻¹ DM) (Figure 7E). The main effects of algal treatment on WSC contents were obtained in plants treated with 2 L.ha⁻¹ where the levels of glucose and fructose were higher (20 and
<figure><img src="image_8.png" /><figcaption>**FIGURE 5** | Relative water content (RWC), as a proxy of osmotic adjustment, of ten last mature leaf blades at the end of priming (A), drought (B) and recovery (C). Dotted line represents the value of well-watered control plants (WW) at the end of priming and drought-stressed control plants (D) at the end of drought and recovery. Each bar represents mean ± standard error (n = 4). Differences between each dose with WW at the end of priming, and differences between each dose with D at the end of drought and recovery, were analyzed by Dunnett's test (†*p* < 0.1; *p* < 0.05; ***p* < 0.01; **p* < 0.05; ***p* < 0.001). Differences between WW and D at the end of drought and recovery were analyzed by Student's or Wilcoxon's test (†*p* < 0.1; *p* < 0.05; ***p* < 0.01; **p* < 0.05; ***p* < 0.001 above the line drawn between WW and D).</figcaption></figure>

<figure><img src="image_9.png" /><figcaption>**FIGURE 6** | Relative Electrolyte Retention Capacity (RERC), as a proxy of cell membrane stability (CMS), of shoot 0–3 cm (containing the leaf meristems) at the end of priming (A), drought (B) and recovery (C). Dotted line represents the value of well-watered control plants (WW) at the end of priming and drought-stressed control plants (D) at the end of drought and recovery. Each bar represents mean ± standard error (n = 4; except n = 1 for 0.5 L. ha⁻¹ treated plants at the end of recovery). Differences between each dose with WW at the end of priming, and differences between each dose with D at the end of drought and recovery, were analyzed by Dunnett's test (†*p* < 0.1; *p* < 0.05; ***p* < 0.01; **p* < 0.05; ***p* < 0.001). Differences between WW and D at the end of drought and recovery were analyzed by Student's or Wilcoxon's test (†*p* < 0.1; *p* < 0.05; ***p* < 0.01; **p* < 0.05; ***p* < 0.001 above the line drawn between WW and D).</figcaption></figure>

24 mg.g⁻¹ DM, respectively) (Figure 7H, 7K) and the level of sucrose was lower (55 mg.g⁻¹ DM; Figure 7E) than in drought-stressed control plants. The lower level of sucrose was thus associated with higher levels of hexoses together with a tendency towards higher fructan level (Figure 7B). The same trend was obtained but only for sucrose in an independent assay (Figure S4E, S4F, S4G, S4H).

At the end of the recovery period, the fructan content was still much lower in plants previously subjected to drought than in well-watered control plants (86 and 195 mg.g⁻¹ DM, respectively; Figure 7C). By contrast, the levels of sucrose, glucose and fructose were similar in drought-stressed and well-watered plants (Figure 7F, 7I, 7L). In plants treated with the algal extract at 0.5, 2, 5 and 15 L.ha⁻¹, fructan content was not different from

that in drought-stressed control plants, whereas it tended to be lower in plants treated with 10 L.ha⁻¹ (*p* = 0.14; Figure 7C). Sucrose contents was mainly affected by the 2 L.ha⁻¹ treatment, which led to higher level in treated plants than in drought-stressed control plants, at the end of the recovery period (Figure 7F).

## 3.7 | Dose-Response Curves

As several parameters showed a dose response revealing a hormetic relationship, i.e. a beneficial effect at low doses and a negative effect at higher doses, a nonlinear regression was applied to the corresponding data using the Brain-Cousens model with five parameters adapted for this type of non-
<figure><img src="image_10.png" /></figure>

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

<figure><img src="image_12.png" /><figcaption>FIGURE 7 | Legend on next page.</figcaption></figure>
monotonic bi-phasic relationship (Ritz et al. 2015) (Figure S5, Table S1). Significant dose effect and appropriate modelling were obtained for water content (Figure S5A) and osmotic adjustment (Figure S5B) in last mature leaf blades at the end of drought and water content in shoot 0–5 cm at the end of drought (Figure S5C), regrowth (Figure S5D) and N uptake (Figure S5E) during recovery.

4 | Discussion

4.1 | Laminaria digitata Extract Improved Leaf Meristem Protection Under Drought and N Uptake After Rehydration in L. perenne

*L. digitata* extract supplied before drought induced a dose-
response effect on plant growth after rehydration with maxi-
mum regrowth between 2 and 5 L.ha⁻¹, which can be explained
by leaf meristem protection during drought and enhanced N
uptake after rehydration.

In grasses, the leaf meristems are the surviving organs during
severe drought (Volaire et al. 2014) and their protection is a
crucial mechanism of drought resistance (Barkaoui and
Volaire 2023). The protection of leaf meristems was made
possible by limiting the loss of membrane stability and by
improving the leaf osmotic adjustment (Zwicke et al. 2015;
Volaire et al. 2020). In perennial ryegrass plants sprayed with
the algal extract and subjected to drought, the slowdown of
RWC decline can explain their higher water content in leaves
and shoot 0-3 cm than in the ones of the drought-stressed
control plants. This can be due to a better osmotic adjustment
resulting from net accumulation of osmolytes. Many com-
pounds act as osmolytes, amino acids and derivatives such as
proline (Li et al. 2020), polyols such as mannitol (Pharr
et al. 1995), sugars such as fructans, sucrose, fructose and
glucose (Garcia et al. 2011; Ozturk et al. 2021). Here, algal
extract supply resulted to higher levels of fructans, glucose
and fructose at the end of drought in shoot 0-3 cm which
include the leaf meristems. Interestingly, fructans not only
act as osmolytes but also as osmoprotectants by insertion
between polar headgroups of phospholipids, thus maintain-
ing the membrane stability (Hincha et al. 2007; Vereyken
et al. 2003). In addition, fructans can prevent membrane
peroxidation by scavenging the hydroxyl radical (OH.)
(Stoyanova et al. 2011). More generally, they play a crucial
role in stress resistance through their antioxidant properties
(Livingston et al. 2009; Nemati et al. 2018). Fructan accu-
mulation could be the result of downregulation of fructan
exohydrolase (FEH) genes as observed in wheat submitted to
drought (Zhang et al. 2009).

Improvement of N content in *Lolium perenne* by biostimulant have already been observed using microbial solution mainly based on lactic acid bacteria (Olszewska 2022). Following N uptake through ¹⁵N labelling during the recovery period gives accurate information on resilience underlying mechanisms (Karlowsky et al. 2018). However, the quantification of N uptake at the whole plant level is time consuming since it requires the sampling and analysis of all plant parts including the roots. Interestingly, the present study highlights the fact that following N uptake only in leaves below 5 cm was a good indicator of N content at the whole plant level since in both cases, value increased as a response to the same doses of the algal extract. The potential of seaweed extracts for plant-growth-promoting activities through N uptake improvement has been well-documented in the literature (Ali et al. 2021; Margal et al. 2023). This is for example the case for the brown algae *A. nodosum* extract in barley (Goñi et al. 2021) and grasses (Quille et al. 2022), and *Laminaria japonica* extract in sugarcane (Chen et al. 2023). To our knowledge, this is the first published report regarding the inducing effect of *L. digitata* on N uptake in plants.

4.2 | *Laminaria digitata* Extract Triggered a
Chemical Priming Effect

*L. digitata* extract supplied before drought was able to improve leaf meristem protection under drought and N uptake after rehydration. This indicates that the algal extract acted as a biostimulant through a priming effect.

The priming effect was first described in the case of systemic
acquired resistance to pathogens, observed after a previous
pathogenic infection (Ross 1961). The priming concept was then
also used in cases of induced resistance to abiotic stress such as
heat and drought following pre-exposure of the plant to the same
or another abiotic stress (Wang et al. 2014a, 2014b; Ben Abdallah
et al. 2017; Ganie et al. 2024). The priming concept is also used in
the case of pretreatment of plant by a chemical product, whether
naturally synthesized or artificial, to improve tolerance to future
biotic and abiotic stress: this is known as “chemical priming” or
“molecular priming” (Savvides et al. 2016; Kerchev et al. 2020).
In the present study, pretreatment with *L. digitata* extract 7 days
before stopping irrigation improved leaf meristem protection
under drought and N uptake after rehydration, demonstrated its
chemical priming action. This indicates that 7 days were suffi-
cient to induce plant resistance mechanisms leading to a “primed
state”, as was observed after a pretreatment of *Arabidopsis* with
an extract from *A. nodosum* seven, five or even 3 days before the
onset of drought (Santaniello et al. 2017; Fleming et al. 2019;
Rasul et al. 2021).

FIGURE 7 | Fructans content in shoot 0–3 cm at the end of priming (A), drought (B) and recovery (C), sucrose content in shoot 0–3 cm at the end of priming (D), drought (E) and recovery (F), glucose content in shoot 0–3 cm at the end of priming (G), drought (H) and recovery (I), and fructose content in shoot 0–3 cm at the end of priming (J), drought (K) and recovery (L). Dotted line represent the value of well-watered control plants (WW) at the end of priming and drought-stressed control plants (D) at the end of drought and recovery. Each bar represents mean ± standard error (n = 4). Differences between each dose with WW at the end of priming, and differences between each dose with D at the end of drought and recovery, were analyzed by Dunnett's test (†*p* < 0.1; *†p* < 0.05; ***†p* < 0.01; **†p* < 0.05; ***†p* < 0.01; ****†p* < 0.001). Differences between WW and D at the end of drought and recovery were analyzed by Student's or Wilcoxon's test (†*p* < 0.1; *†p* < 0.05; ***†p* < 0.01; **†p* < 0.05; ***†p* < 0.001 above the line drawn between WW and D).
In the present study, most of the parameters monitored (biomass production, N uptake, water content and membrane stability) were not altered during the priming period before stress. Nevertheless, slight changes were detected in the soluble carbohydrate composition of shoot bases at the end of the priming period. The lower doses of algal extract induced a decrease of fructan content, while the higher doses induced an increase in sucrose content in leaf base. These small changes may be the result of larger changes in cell signalling and gene expression triggered by chemical priming (Antoniou et al. 2016). It is well-known that seaweed extracts and some of their bioactive compounds act as elicitors upregulating numerous defense-related genes and plant hormones (Ali et al. 2022). In *Arabidopsis*, treatment with *A. nodosum* extract affected the expression of *LOX2*, a key gene for jasmonic acid synthesis, and *ERF1A* encoding a regulatory factor for the ethylene signalling pathway during the priming period preceding the drought period (Fleming et al. 2019). Moreover, *A. nodosum* extract also modulated expression of genes associated with ABA-responsive pathways (Santaniello et al. 2017; Ali et al. 2022) together with partial stomatal closure (Santaniello et al. 2017). Partial stomatal closure might lead to ROS production, which in turn could act as signals triggering antioxidant enzyme production (Karpinski et al. 1999). Indeed, partial stomatal closure following pretreatment with *A. nodosum* is associated with alteration in expression of genes associated with antioxidant system pathways (Santaniello et al. 2017). Stronger antioxidant defense system might prepare plants for drought resistance. As far as *L. digitata* and *L. perenne* are concerned, analyses linked to signalling compounds and gene expression at the priming state are required to identify the mechanisms of action and therefore, clarify the priming effect itself.

## 4.3 | Laminaria digitata Extract Triggered Hormetic Dose-Responses

*L. digitata* extract supplied before drought improved leaf meristem protection and N uptake after rehydration at low doses but had a toxic effect at higher doses, revealing a biphasic dose-response called hormesis, a phenomenon which has long been under-recognized in toxicology but is now generally accepted in different fields of biology and have been demonstrated for numerous physical, chemical and biological agents (Calabrese and Baldwin 2003). The hormetic effect of the algal extract was observed at the end of drought, in particular for tissue water content, which was improved with the 2 L.ha⁻¹ treatment and reduced with the 10 and/or 15 L.ha⁻¹ treatments, depending on the tissue considered. Hormesis was also clearly observed at the end of recovery, with low doses improving regrowth and N uptake, while the highest dose was deleterious. Similar hormetic response curves have previously been obtained for wheat and rice yield and grain quality following foliar applications of red algae extracts from *K. alvarezii* and *Gracilaria edulis* (Shah et al. 2013; Layek et al. 2018), suggesting that hormesis is a common process for the response of plants to algal biostimulants. Hormesis has also been demonstrated for various types of compounds so that it is a generalizable process for the response of plants to biostimulants (Godínez-Mendoza et al. 2023). At the physiological level, this phenomenon is considered to be a

response to a low level of stress which allows the organism to acclimate to its new environment (Calabrese et al. 2007; Vargas-Hernandez et al. 2017). More specifically, our study is a typical case of hormesis-based plant priming which promotes plant resistance to environmental adversities (Christou et al. 2022), indicating that biostimulants based on *L. digitata* can contribute to agricultural sustainability within the context of climate change. Indeed, as natural products are effective at low doses, the risk of persistence in the environment is very limited. In terms of costs of application in the field, the current application rates being low and targeted in the crop cycle, the financial impact per hectare remains reduced with an advantageous return on investment for the farmer, particularly in cases of great stress. However, because of their possible counter effects on plants at inappropriate higher doses, their use must be rigorously controlled.

## 4.4 | Development of a Plant Drought-Resistance Bioassay for the Detection of Biostimulants

To determine whether the protocol can be used as a bioassay to test the biostimulant effect of various products on grass resistance to drought, its reproducibility was checked. Two independent assays were carried out using the same experimental protocol and the same algal extract sprayed on leaves at 0.5, 2 or 5 L.ha⁻¹. The foliar treatment at 2 L.ha⁻¹ gave the best results in both assays. Although the drought-stressed control plants of the two independent assays appeared slightly differently altered, the algal extract triggered a similar improvement in leaf meristem protection based on shoot 0–3 cm water content and cell membrane stability, and a similar dose-response effect. In addition, the seaweed extract improved the last mature leaf blade RWC and limited the increase in sucrose content during drought, in both assays. The limitation of sucrose content increase might be due to its use for the synthesis of fructans by fructosyltransferases or for its hydrolysis by invertases, through the upregulation of the corresponding genes (Lasseur et al. 2011). In both cases, the production of glucose and/or fructose might be used for the production of other osmoprotectants such as proline that has been reported to accumulate in *L. perenne* in response to drought (Li et al. 2020). Since the algal treatment increased the content of fructans, glucose and fructose but only in plants most severely affected by drought, these parameters are not suitable general indicators of biostimulation. Instead, RWC of the last mature leaf blade, cell membrane stability and sucrose content of the shoot 0–3 cm represent reliable and reproducible indicators of biostimulation.

## 5 | Conclusion

The seaweed extract from *L. digitata* improved leaf meristem protection through fructan, glucose and fructose accumulation during drought and N uptake after rehydration in perennial ryegrass (*L. perenne* var. Delika) grown in controlled condition when plant leaves were sprayed at 2 L.ha⁻¹. The underlying mechanisms by which the seaweed extract and its constituents improve drought survival through hormesis-based chemical priming, will be investigated by metabolomic and transcriptomic approaches in future experiments. Identification of
the most active compounds and their mode of action may also
be useful to improve the effect on biomass yield.

Perennial ryegrass is an emblematic species of temperate grasslands, which will face more intense periods of drought in the near future due to climate change. Despite the weak effect of the biostimulant on biomass yield, the ability of *L. digitata* extract to improve N uptake may allow fertiliser rates to be limited, thereby reducing costs to the farmer and environmental impact. Field experiments will be carried out to evaluate the effectiveness of the algal extract in a natural environment and recommend its conditions of use. Indeed, due to dose effects, dose application must be carefully considered to ensure beneficial effects of the extract.

## Acknowledgements

The authors thank Magali Bodereau, Josiane Pichon, Anne-Françoise Ameline and Olivier Rose for helping with plant harvesting and sample grinding, Paul Moulin for his contribution to the preliminary tests during his scholarship. They acknowledge the PLATIN' (Plateau d'Isotopie de Normandie) core facility for element analysis used in this study. The work was supported by the University of Caen Normandie and by the company Algaia. The Normandie Council and the European Union funded the work through the research projects AlgoEVA (Valorisation des algues en Normandie pour les secteurs de l'agriculture et de la santé animale, 2018-2021, in the framework of the ERDF-ESF) and BIOSTIM-ALMAP (Intérêt des biostimulants d'origine algale pour la gestion des cultures maraîchères et des prairies dans le contexte du dérèglement climatique et de la transition agroécologique, 2022-2025, in the framework of the EAFRD). Antoine Grandin-Courbet received a PhD grant (2022-2025) from the ANRT (Association Nationale de la Recherche et de la Technologie) (CIFRE 2021/1265).

## Data Availability Statement

The data that support the findings of this study are available from the
corresponding author upon reasonable request.

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## Supporting Information

Additional supporting information can be found online in the Supporting Information section.