Research ArticleArticles

Growth, Physiology, and Root Development in Seedlings of Woody Species Treated with a Seaweed Extract

Sebastien Comin, Gloria Brocca, Noemi Valsecchi, Simone Fumagalli, Irene Vigevani, Denise Corsini, Francesco Ferrini, Giovanni Ravanelli and Alessio Fini
Arboriculture & Urban Forestry (AUF) September 2024, jauf.2024.013; DOI: https://doi.org/10.48044/jauf.2024.013

Abstract

The demand for saplings has risen in recent years as a consequence of massive planting campaigns targeted at increasing canopy cover. To test the hypothesis that seaweed extract can improve root biomass and length, an experiment was carried out at the ERSAF Regional Forest Nursery in Curno, Italy. The seeds of 5 woody species were planted in trays using a substrate amended with 0×, 1×, 2×, or 3× the label dose of a pure Ascophyllum nodosum extract. After germination, 6,400 seedlings were arranged according to a randomized complete block design with 10 blocks. After 1 growing season, plants were transplanted into 1.7-dm3 forest containers for 1 additional growing season. Root, stem, and leaf dry weights, total leaf area, total root length, and specific root length were measured over an 80-week period. Leaf gas exchange and greenness index were monitored for 78 weeks using an infrared gas analyser and a SPAD meter. Species differed for growth rate, biomass allocation to roots, and specific root length. The algal biostimulant increased stem and whole plant dry weights for 1 year only when applied at 3x the label dose. Significant effects on leaf gas exchange were found only at the highest dose and were mostly due to higher leaf greenness index than to lower diffusional limitations to photosynthesis. Results suggest that substrate amendment with Ascophyllum extracts may have short term positive effects on plant growth, likely due to a nutritional boost. However, they did not trigger structural changes in plant traits that can enhance transplant tolerance in the long run.

Keywords

Introduction

The growing awareness about the ecosystem services (ES) provided by urban vegetation (Bowler et al. 2010; Rahman et al. 2020) has led several municipalities and institutions to start extensive tree planting programs worldwide. Such initiatives have increased the demand for different types of nursery stocks, ranging from seedlings of native species for the development of urban and peri-urban woodlands to much larger plant material (e.g., from 14 to 16 cm to 20 to 25 cm in circumference) for park and street tree plantings.

One of the major threats to the success of such planting efforts is transplant stress, which can result in the mortality of up to one-third of newly planted trees within five years of transplant (Roman et al. 2014; Elmes et al. 2018; Breger et al. 2019). Environmental conditions of the planting site, quality of nursery stock, and availability of resources for site amelioration and postplanting care are crucial determinants of establishment rate (Koeser et al. 2014; Roman et al. 2014; Hilbert et al. 2019). A series of cultural techniques, collectively grouped as nursery preconditioning, have been proposed to produce sturdy nursery plants with a superior capacity to recover from transplant shock (Franco et al. 2006). Although it may result in lower growth rates in the nursery, preconditioning techniques can improve plant morphological (e.g., higher root:shoot ratio, higher fine root development), physiological (e.g., higher mesophyll:stomatal conductance ratio, constitutive osmotic adjustment), and biochemical traits (e.g., higher antioxidant capacity) that can induce a higher stress tolerance after transplanting (Franco et al. 2006).

The use of biostimulants as a preconditioning method has been recently proposed in fruit tree species (Conesa et al. 2020), while research on the suitability of biostimulants to harden forest and shade species is still scarce (MacDonald et al. 2012; Elansary et al. 2016). Plant biostimulants include any compounds or microorganisms that can improve nutrient uptake, promote tolerance to abiotic stresses, and enhance crop quality traits (du Jardin 2012). Seaweed extract is one of the 6 categories of biostimulants which is actually marketed (Critchley et al. 2021). Algae have been used as soil conditioners to increase fertility and crops productivity for millennia, but the first commercial seaweed extract was released for agricultural purposes about 70 years ago (Craigie 2011; Calvo et al. 2014). Brown algae are the most used in agriculture: nowadays, about 28.5-million tons of algae-based products are manufactured annually (FAO 2016), and these kinds of extracts are dominating the biostimulants global market (Critchley et al. 2021). Seaweeds can be applied directly to plants, as foliar sprays, or to growing substrates, as a liquid or powder formulation (du Jardin 2012). Seaweed extracts contain a complex mixture of compounds that can change according to the seaweed species, the harvest time, and the kind of extraction process used. Ascophyllum nodosum, the most widely used algae for producing seaweed extracts, is a brown macroalgae that belongs to the family Fucaceae of the order Fucales. It grows along intertidal, rocky shores throughout the North Atlantic region (Pereira et al. 2020). Ascophyllum nodosum biostimulants are available in liquid or dried form, raw or obtained after extraction (Shukla et al. 2019). The chemical composition of A. nodosum tissues has been carefully studied due to its increasing commercial interest. The major inorganic components are N, Na, Mg, K, Cl, and SO42−. Organic components include alginic acid, mannitol, amino acids, laminarin, fucoidan, polyphenols, and phytohormones (Pereira et al. 2020). Commercial extracts of A. nodosum contain about 45% of carbohydrates, 18% of ashes, 5% of proteins, 3% of lipids, 1.5% of polyphenols, and 13.5% of other compounds (Moreira et al. 2017).

Several studies have revealed that A. nodosum improved seed germination and crop growth by enabling greater nutrient availability and assimilation (Rayorath et al. 2008; Khan et al. 2009; Craigie 2011; Van Oosten et al. 2017). In particular, root growth was promoted by the application of A. nodosum extracts at a concentration of 0.1 g L−1 (Khan et al. 2009). Calvo et al. (2014) reported that the application of seaweed extracts could increase the number of lateral roots as well as the root volume and length in both woody and herbaceous plants. Similarly, Tkaczyk et al. (2022) found that A. nodosum extracts could promote root lengthening of Quercus robur seedlings. According to Jeannin et al. (1991), seaweed extract applied as a foliar spray increased the total fresh matter production of maize seedlings by 15% to 25% over the control. This was reflected in the increase of root and stem mass per plant of maize (Zea mays L.). Furthermore, Nelson and van Staden (1986) demonstrated that seaweed extracts could increase root:shoot ratio in wheat (Triticum aestivum). Ascophyllum nodosum extracts were also reported to increase leaf antioxidant capacity, net CO2 assimilation, and ameliorate plant water relations during drought stress in woody species (Elansary et al. 2016; Salvi et al. 2019; Frioni et al. 2021). Thus, the application of A. nodosum extracts may promote morphological and physiological adjustments in plants that can improve stress tolerance without depressing plant growth rate in the nursery, as occurring when other preconditioning techniques, such as deficit irrigation or root pruning, are used (Amoroso et al. 2010; Fini et al. 2011). This may be of great interest to the nursery industry, which is now forced to satisfy a massive demand for plants while trying to match environmental and economic sustainability and production quality (Nicese and Ferrini 2008).

The aim of this research was to test the hypotheses that the amendment of the nursery substrate with a pure A. nodosum extract: (1) can increase seed germination and plant growth, thereby accelerating plant production process, and (2) can induce persistent changes in morpho-physiological traits that promote resilience to transplanting stress. An additional aim of the experiment was to identify an optimal dose of application for seedlings of woody species.

Materials and Methods

Plant Material and Growing Conditions

The experiment was carried out at the Ente Regionale per i Servizi all’Agricoltura e alle Foreste (ERSAF) forest nursery in Curno (BG, Italy; 45°41′28″N, 9°36′45″E). The experimental site has a temperate climate with no dry season and a warm summer (Cfb according to Köppen-Geiger classification). The average temperature recorded from 1991 to 2021 was 11.5 °C, while the average rainfall was 1,420 mm per year. The experimental years were warmer (average temperature was 13.4 and 14.9 °C in 2021 and 2022, respectively) and moister (total rainfall was 1,614 and 1,842 mm in 2021 and 2022, respectively) than the 30-year average (Figure S1 in Appendix).

Seeds of Amelanchier ovalis (AOV), Carpinus betulus (CBE), Crataegus monogyna (CMO), Fagus sylvatica (FSY), and Ligustrum vulgare (LVU) were collected from certified seed forests according to the Italian legislation on plant production for forestry (Legislative Decree 10 November 2003, n. 386). The specific locations used for seed collection and their climatic characteristics are reported in Table 1. Seeds were cleaned with a sodium hypochlorite solution and sterilized using copper, according to standard practices. Seeds were then stratified into moist sand and vernalized for 12 weeks at 4 °C in a refrigerated room.

View this table:
Table 1.

Locations of the certified seed forests where seeds were collected and their climatic traits. The climate type (Koppen-Geiger classification), the average rainfall per year, and the average annual temperature data were collected from climate-data.org.

Seeds were sown into 200 32-cell forest seedling trays. Each cell had 0.4-dm3 volume. To ensure adequate plant material for the research, 3 seeds were sown in each cell (19,200 seeds were used in total for the experiment). After germination assessment (see Measurements section), seedlings were selectively thinned to retain only 1 plant per cell (6,400 seedlings). Trays were filled with 2,600 L of Hoochmoor® substrate (Terflor S.r.l., Carpiolo, BS, Italy), specifically designed for seedlings. The substrate was made of black peat (0.1 to 10 mm):blonde peat (10 to 20 mm):coconut fiber (0.1 to 15 mm)(2:1:1) amended with 10-kg m−3 montmorillonite and fertilized with 1-kg m−3 soluble N-P-K fertilizer (14-16-18) and 1-kg m−3 Osmocote® (ICL, Tel-Aviv, Israel)(8 to 9 months). Substrate was corrected with calcium carbonate to neutralize pH. Substrate pH, electric conductivity, bulk density, and porosity were 6.0, 0.3 dS m−1, 100 kg m−3, and 95%, respectively. Trays were placed in a shadehouse covered with a green woven polyethylene fabric which provided 50% shading, according to standard practices for forest nurseries. No additional fertilization was provided to seedlings other than that included in the substrate. Plants were irrigated once per day using a sprinkler system for 10 (Spring and Fall) to 15 (Summer) minutes according to standard nursery practices. Weeding was carried out manually. No pesticide application was carried out.

In January 2022, after a single growing season in the trays, 2 plants per tray (400 plants in total) were potted into 1.7-dm3 containers. Plants were selected as those with average height among those growing in the same tray. Containers were filled using a black peat (0.1 to 20 mm):blonde peat (20 to 40 mm):coconut fiber (0.1 to 15 mm):pumice (3 to 8 mm)(15:45:20:20) substrate, corrected with calcium carbonate to achieve a pH of 6.0. Plants were arranged in a shadehouse as previously described and subjected to similar management as in the first growing season.

Biostimulant Treatments

The biostimulant used in this research was a commercial pure extract of A. nodosum obtained by drying and grinding the algae at low temperature (Regulalg; Agrofertil, Allones, France). The biostimulant was made of 50% mineral components, 35% organic components, and 15% moisture. Organic components included: 54% carbohydrates, 24% alginic acid, 8% mannitol, 5% laminarin, and 9% fucoidan. The biostimulant included vitamins such as ascorbic acid (1,000 ppm), carotene (40 ppm), riboflavin (10 ppm), tocopherol (140 ppm), and nutrients such as nitrogen (0.8% to 3%), phosphorus (0.1%), potassium (1% to 3%), calcium (1% to 3%), sulphur (2% to 5%), iron (150 to 1000 ppm), boron (20 to 100 ppm), and zinc (40 to 200 ppm)(Agrofertil 2015).

The biostimulant was mixed with the substrate used for filling the forest trays at different doses: (1) no addition of biostimulant (D0), (2) biostimulant added at the dose recommended by the manufacturer of 1 kg m−3 of substrate (D1), (3) biostimulant added at a dose of 2 kg m−3 (D2), and (4) biostimulant added at a dose of 3 kg m−3 (D3). No further addition of biostimulant was made when plants were repotted into 1.7-L containers.

Measurements

Seed germination rate was measured 2 weeks after seedling emergence (which occurred around mid-March 2021) as the number of alive plants in each tray over the number of seeds seeded (96 per tray).

Biomass and leaf area measurements were conducted in early April 2021 (2 weeks after emergence), in between the end of May and the beginning of June 2021 (9 weeks), in mid-July 2021 (15 weeks), in mid-October 2021 (30 weeks), and in early October 2022 (80 weeks after emergence) on 1 plant per species, treatment, and block (200 plants in total in each sampling date), according to standard protocols (Fini et al. 2011). Within each tray, the plant with height closest to the tray average was harvested and cut at the root flare.

Roots were cleaned from the substrate with a flush of air. Total root length per plant (TRL) was calculated from the number of interceptions between the root system and a 1 × 1 cm grid, according to the modified line intercept method (Tennant 1975). A factor of 11/14 was used to convert intercept number to root length (Tennant 1975). Measurements were conducted on either entire root systems (2, 9, 15, and 30 weeks after emergence) or on a known fraction (about 30% w/w) of the root system (80 weeks after emergence).

Leaves were separated from the stem and were scanned with an A3 scanner (HP® OfficeJet Pro 7740; HP Inc., Palo Alto, CA, USA). An image-analysis software (ImageJ v. 1.53) was used to measure total leaf area per plant (TLA).

Dry weights of leaves, stems, and roots were measured after oven-drying the different plant organs separately at 70 °C until constant weight was reached (≈ 72 h). Total plant dry weight (DWplant) was calculated as the sum of the dry weights of leaves, stem, and roots. Specific root length (SRL) was calculated as the ratio between TRL and root dry weight (DWroots) (Amoroso et al. 2011). Root to shoot ratio was calculated as the ratio between DWroots and leaves + stem dry weight. The ratio between TLA and TRL was calculated as TLA/TRL. Relative growth rate (RGRplant) was calculated as (Fini et al. 2010): Embedded Image

where DWplantt1 and DWplantt0 denote plant dry weights at consecutive sampling periods and t1 – t0 denotes the number of days between samplings. For the estimation of RGRplant during the first 2 weeks after emergence, DWplantt0 was set equal to seed dry weight.

Leaf gas exchange was measured on 3 species (CBE, CMO, FSY), subsampled based on shade tolerance (CBE: partial shade-tolerant; CMO: sun-requiring; FSY: shade tolerant) and on differences in root traits recorded 9 weeks after emergence, when leaf gas exchange measurements started. Species subsampling was done because of the need to keep leaf gas exchange measurements within 3.5 hours to avoid excessive change in environmental conditions during the measurement. Measurement parameters were net CO2 assimilation (A, μmol m−2 s−1), transpiration (E, mmol m−2 s−1), stomatal conductance (gs, mmol m−2 s−1), and substomatal CO2 concentration (Ci, ppm). Measurements were performed using an infrared gas analyser (CIRAS-2; PP Systems, Amesbury, MA, USA) on 10 fully expanded leaves per species and biostimulant treatment (120 leaves per sampling date), and were conducted in June 2021 (9 weeks after seedling emergence), July 2021 (15 weeks), October 2021 (30 weeks), June 2022 (63 weeks), and September 2022 (78 weeks). In the cuvette, leaves were supplied with 420 ppm CO2 (Ca), provided using a CO2-cartridge, and saturating (1,300 μmol m−2 s−1) light intensity, provided using the integrated light source. In July 2021 and June 2022, simultaneous measurements of leaf gas exchange and chlorophyll fluorescence were carried out using the built-in chlorophyll fluorescence module (CFM; PP Systems, Amesbury, MA, USA). The actual quantum yield of PSII (ΦPSII) was measured after exposing light-adapted leaves to a saturating pulse of white light (6,000 μmol m−2 s−1, 0.8-s duration) (Genty et al. 1989). The linear electron transport rate (Jf) was calculated from ΦPSII as: Embedded Image

where a is leaf absorbance, set at 0.84, PAR is radiation, set at 1,300 μmol m−2 s−1, and 0.5 assumes equal distribution of irradiance between PSI and PSII (Tattini et al. 2015).

Mesophyll conductance to CO2 diffusion (gm, mmol m−2 s−1) was calculated using the variable J method (Harley et al. 1992) as previously described in detail (Fini et al. 2016). CO2 concentration in the chloroplasts (Cc) was finally estimated as: Embedded Image

Diffusive limitations to photosynthesis were estimated as the drawdown of CO2 from the ambient air to the substomatal chamber (stomatal limitation, Ca – Ci) and from the substomatal chamber to the chloroplasts (Ci – Cc)(Loreto et al. 2003).

Leaf greenness index was measured immediately after and on the same leaves used for leaf gas exchange measurements using a SPAD-meter (SPAD-502Plus; Konica Minolta, Inc., Tokyo, Japan)(Percival et al. 2008).

Experimental Design and Statistics

During the first growing season, when plants were grown into trays, the experimental design was a randomized complete block with 10 blocks and 1 tray (containing 32 seedlings) per species and biostimulant dose in each block (640 seedlings per block, 6,400 seedlings in total). During the second growing season, after repotting into containers, the experimental design was a randomized complete block with 10 blocks and 2 saplings per species and biostimulant treatment in each block (40 saplings in each block, 400 saplings in total).

All data were analysed using SPSS statistical package (Version 21.0; IBM, Armonk, NY, USA). To evaluate the effects of species, biostimulant dose, time after emergence, and their interactions on measured traits, a mixed-model procedure was applied. Homogeneous subsets were separated using the Sidak test. When significant species × time after emergence or biostimulant treatment × time after sampling interactions were found, the different species/biostimulant treatments were compared within each date of sampling.

Results

Effect of Biostimulant Application on Seed Germination

Seed germination was affected by species (P < 0.000) and by biostimulant treatment (P < 0.000). A significant (P < 0.000) species × biostimulant treatment interaction was found for the % of germinated seeds. Overall, FSY seeds had the highest germination rate among the species tested (62%) while AOV had the lowest germination rate (15%). The application of the biostimulant at 2× and 3× label doses significantly improved germination, compared to D0 and D1 treatment, only in FSY (+69%)(Figure 1).

Figure 1.
Figure 1.

Effect of biostimulant dose (D0, D1, D2, D3) on seed germination (%) in the 5 woody species tested: Amelanchier ovalis (AOV), Crataegus monogyna (CMO), Carpinus betulus (CBE), Fagus sylvatica (FSY), and Ligustrum vulgare (LVU). Different letters indicate significant differences among biostimulant doses within each species at P < 0.01, n.s. = P > 0.05.

Effect of Species and Biostimulant Application on Root Traits

Root dry weight (DWroots, g), specific root length (SRL, m g−1), and total root length per plant (TRL, m) were significantly affected by species and by time after seedling emergence, but not by biostimulant dose (Table S1 in Appendix).

Two weeks after seedling emergence (early April 2021), FSY had higher DWroots than other species (Figure 2A) and, despite a low SRL (Figure 2B), it had higher TRL than other species except CMO (Figure 2C). Higher SRL observed in CMO than in other species allowed CMO to reach similar TRL as FSY, although DWroots was 69% lower. Nine weeks after emergence (May to June 2021), FSY and CMO had higher DWroots than other species (Figure 2D). Nonetheless, low SRL displayed by FSY resulted in this species having similar TRL as AOV and LVU (Figure 2E-F). Fifteen weeks after emergence (mid-July 2021), FSY and CMO had higher DWroots than other species (Figure 2G). Conversely, LVU displayed the lowest DWroots among the species investigated. Nonetheless, because of higher SRL (Figure 2H), LVU displayed higher TRL than other species (Figure 2I). At the end of the first growing season (30 weeks after emergence, mid-October 2021), DWroots ranked as CMO > FSY > AOV > LVU = CBE (Figure 2J). SRL was higher in LVU than in CBE which, in turn, had higher SRL than other species (Figure 2K). TRL was higher in CMO than in LVU which, in turn, had higher TRL than AOV and FSY (Figure 2L). CBE had the lowest TRL among the investigated species after 30 weeks since emergence. After repotting and a single additional growing season (80 weeks after emergence, early October 2022), CMO had 2-fold higher DWroots than CBE and 3.5-fold higher DWroots than AOV, FSY, and LVU (Figure 2M). SRL was 1.7-fold higher in LVU than in AOV and CMO (Figure 2N). As a result, 80 weeks after emergence, TRL ranked as CMO > LVU = CBE > FSY > AOV (Figure 2O).

Figure 2.
Figure 2.

(A, D, G, J, M) Root dry weight (DWRoots [g]); (B, E, H, K, N) specific root length (SRL [m g−1]); and (C, F, I, L, O) total root length (TRL [cm]) of 5 woody species: Amelanchier ovalis (AOV); Carpinus betulus (CBE); Crataegus monogyna (CMO); Fagus sylvatica (FSY); and Ligustrum vulgare (LVU) at 2 weeks (A, B, C), 9 weeks (D, E, F), 15 weeks (G, H, I), 30 weeks (J, K, L), and 80 weeks (M, N, O) after seedling emergence. Different letters indicate significant differences among species within each sampling date at P < 0.01, n.s. = P > 0.05.

Neither TRL nor DWroots were affected by the application of the biostimulant at any dose (Figure 3A-B). In contrast, the application of the biostimulant at 2× and 3 × label doses decreased SRL, compared to control, on 2 and 80 weeks after seedling emergence (Figure 3C). On 15 weeks after emergence, D1 treatment had higher SRL compared to control.

Figure 3.
Figure 3.

Effect of biostimulant dose (D0 = no application; D1 = 1 kg m−3; D2 = 2 kg m−3; D3 = 3 kg m−3) on (A) average total root length (TRL [cm]); (B) root dry weight (DWroots [g]); and (C) specific root length (SRL [m g−1]) measured at 2 weeks, 9 weeks, 15 weeks, 30 weeks, and 80 weeks after seedling emergence. Different letters indicate significant differences among doses within each sampling date at P < 0.01, n.s. = P > 0.05.

Effects of Biostimulant Application on Plant Growth

Plant relative growth rate (RGRplant, mg g−1 day−1), total plant dry weight (DWplant, g), and total plant leaf area (TLA, cm2) differed among species (Table S2 in Appendix). RGRplant differed among species over the 80 weeks after seedling emergence: FSY displayed, on average, lower RGRplant than other species (data not shown). Consistently, on average over the entire experiment, DWplant was 41% lower in FSY than in other species, and TLA was 75% lower in FSY than in CMO and CBE (data not shown).

The biostimulant treatment affected DWplant, and a significant dose × time after seedling emergence interaction was found for DWplant, RGRplant, and TLA (Table S2 in Appendix). For the first 2 weeks after seedling emergence, plants amended with 2× (D2) and 3× (D3) label dose of seaweed extract increased RGRplant compared to plants amended with the extract applied at the label dose (D1) or unamended plants (D0)(Figure 4B). Later on, differences among treatments in RGRplant were no longer significant. DWplant and TLA were higher in D3 treatment compared to control plants on 2, 15, and 30 weeks after emergence, whereas D3 treatment showed similar DWplant and TLA as D0 plants on 9 and 80 weeks after emergence (Figure 4A-C). D2 and D1 treatments failed to consistently increase DWplant compared to D0.

Figure 4.
Figure 4.

Effect of biostimulant dose (D0 = no application; D1 = 1 kg m−3; D2 = 2 kg m−3; D3 = 3 kg m−3) on (A) average total plant dry weight (DWplant [g]); (B) relative growth rate of the whole plant (RGRplant [mg g−1 day−1]); and (C) total leaf area (TLA [cm2]) measured at 2 weeks, 9 weeks, 15 weeks, 30 weeks, and 80 weeks after seedling emergence. Different letters indicate significant differences among doses within each sampling date at P < 0.01, n.s. = P > 0.05.

Root to shoot ratio (R:S, g g−1) and the ratio between total leaf area and total root length (TLA:TRL, cm2 and cm−1 respectively) were significantly affected by species and by time after seedling emergence but not by biostimulant dose (Table S2 in Appendix). A species × time interaction was found for all investigated traits, and a dose × time interaction was found for SRL.

FSY had higher R:S than other species on all sampling dates except on 2 weeks after emergence, when AOV, LVU, and CBE had higher R:S than FSY (Figure 5A). R:S of FSY increased 6-fold over the first 30 weeks after seedling emergence. On the contrary, R:S of LVU increased only 4% over the same period, resulting in LVU showing lower R:S than other species 30 weeks after emergence. Eighty weeks after emergence, FSY had 43% higher R:S than CBE, 50% higher R:S than AOV and CMO, and 61% higher R:S than LVU. CMO had, in general, higher TLA:TRL than other species, except on 2 weeks after emergence, when FSY had higher TLA:TRL (Figure 5B). Except for 2 weeks after emergence, LVU had lower TLA:TRL than other species (Figure 5B).

Figure 5.
Figure 5.

Effects of species Amelanchier ovalis (AOV); Crataegus monogyna (CMO); Carpinus betulus (CBE); Fagus sylvatica (FSY); and Ligustrum vulgare (LVU) and biostimulant dose (D0 = no application; D1 = 1 kg m−3; D2 = 2 kg m−3; D3 = 3 kg m−3) on (A, C) root to shoot ratio (R:S [g g−1]) and on (B, D) the ratio between total leaf area (TLA) and total root length (TRL)(cm2 cm−1) measured at 2 weeks, 9 weeks, 15 weeks, 30 weeks, and 80 weeks after seedling emergence. Different letters indicate significant differences among species (A, B) and doses (C, D) within each sampling date at P < 0.01, n.s. = P > 0.05.

Biostimulant dose did not affect R:S or TLA:TRL throughout the experiment (Figure 5C-D).

Effects of Biostimulant Application on Leaf Gas Exchange

Leaf gas exchange data were significantly affected by species, dose of biostimulant, and time of sampling (Table S3 in Appendix). Significant species × time and dose × time interactions were found for transpiration (E) and net CO2 assimilation (A) per unit leaf area. A significant species × dose interaction was found for A. Significant species × dose × time interactions were found for both the drawdown of CO2 from the outer air to the substomatal chamber (Ca – Ci) and the drawdown of CO2 from the substomatal chamber to the chloroplasts (Ci – Cc) and for leaf greenness index (Table S3 in Appendix).

On average over the entire experiment, FSY displayed 26% and 51% higher E and A, respectively, compared to CBE (data not shown). Although CMO had 19% lower E than FSY, A did not differ between these 2 species. In general, the application of the biostimulant at 3× label dose increased both E (+9%) and A (+17%) compared to D0 plants, whereas only E (+11%) was increased by application at 2× label dose (data not shown). The effects of biostimulant application on E and A, however, varied depending on species and date of sampling.

D3 plants of CBE displayed 13% higher E compared to other treatments in July 2021 (15 weeks after emergence), whereas no significant differences among treatments were found in the other sampling dates (Figure 6A). CMO plants amended with 3× label dose showed 7% and 20% higher E compared to D0 treatment in July 2021 (15 weeks) and October 2021 (30 weeks), respectively. In October 2021, D2 CMO showed similar E as D3 treatment (Figure 6B). In FSY, D2 and D3 plants displayed 19% higher E than D0 and D1 treatments in July 2021 (15 weeks after emergence)(Figure 6C). Net CO2 assimilation of CBE plants amended a 3× label dose was 39% higher compared to D0 and D1 treatments in July 2021 (Figure 6D). In CMO, D3 treatment increased A, compared to D0 plants, in July (+35%) and October (+61%) 2021 (Figure 6E). In June 2022, D3 CMO plants had 42% higher A compared to D1 treatment, while control didn’t differ from either. D2 and D3 plants of FSY displayed higher A than D0 plants in June (+32%) and July (+44%) 2021 (Figure 6F).

Figure 6.
Figure 6.

Effect of biostimulant dose (D0 = no application; D1 = 1 kg m−3; D2 = 2 kg m−3; D3 = 3 kg m−3) on (A, B, C) transpiration per unit of leaf area (E [mmol m−2 s−1]) and (D, E, F) net CO2-assimilation (A [μmol m−2 s−1]) measured in Carpinus betulus (A, D), Crataegus monogyna (B, E), and Fagus sylvatica (C, F) during June 2021, July 2021, October 2021, June 2022 and September 2022. Different letters indicate significant differences among doses within each sampling date at P < 0.01, n.s. = P > 0.05.

CMO displayed 8% and 13% higher Ca – Ci compared to FSY and CBE, respectively. CBE had 8% and 16% higher Ci – Cc compared to CMO and FSY, respectively. CMO had 8% and 24% higher leaf greenness index compared to FSY and CBE, respectively. The application of the seaweed extract affected Ca – Ci and Ci – Cc only during the first year after its application, and consistent effects were only detected in FSY (Figure 7A-F). In this species, D2 and D3 treatments displayed higher Ca – Ci and lower Ci – Cc, compared to control (Figure 7C-F). Seaweed application at any dose failed to increase leaf greenness index in CBE, compared to control (Figure 7G). Conversely, D3 plants of CMO and FSY had 18% and 27% higher leaf greenness index compared to D0 treatment in 2021 (Figure 7H-I). D2 treatment was also successful in increasing leaf chlorophyll compared to D0 plants of FSY during 2021 (Figure 7I). For both CMO and FSY, differences among biostimulant treatments were no longer significant in 2022 (Figure7H-I).

Figure 7.
Figure 7.

Effect of biostimulant dose (D0 = no application; D1 = 1 kg m−3; D2 = 2 kg m−3; D3 = 3 kg m−3) on the drawdown of CO2 from (A, B, C) the outer air to the substomatal chamber (Ca-Ci [ppm]), from (D, E, F) the substomatal chamber to the sites of carboxylation (Ci-Cc [ppm]), and on (G, H, I) leaf greenness index (SPAD) measured in Carpinus betulus (A, D, G), Crataegus monogyna (B, E, H), and Fagus sylvatica (C, F, I) in 2021 and 2022. Different letters indicate significant differences among doses within each sampling date at P < 0.01, n.s. = P > 0.05.

Discussion

The first aim of this research was to test whether the amendment of nursery substrate with different doses of an A. nodosum extract could improve growth of plants during nursery cultivation. It may be crucial to hasten the production process of suitable ecotypes of a broad variety of plant species, to match the growing demand for plants triggered by the flourishing of extensive tree planting programs worldwide (Conway and Vander Vecht 2015). The development of urban and peri-urban woodlands is sensitive to the nursery availability of local ecotypes of native species (Jim 2017). Our results revealed that the application of A. nodosum extracts to the nursery substrate at rates of 2 or 3 kg m−3 improved seed germination, compared to untreated control, only in FSY. This effect may be due to a faster mobilization of endosperm and cotyledon reserves to support seedling growth, which can be achieved through a higher activity of α-amylase (Rayorath et al. 2008), and which may be relevant in beech seedlings which are characterized by large cotyledons. The mechanism through which A. nodosum extracts can enhance α-amylase haven’t been fully elucidated, but it is known that the enzyme activity is positively affected by gibberellins. Because gibberellins are absent in the extract, its positive effect might be due to nongibberellin components which can induce gibberellin-like effects in a gibberellin-independent manner (De Saeger et al. 2020). Such effect, however, cannot be generalized because the impact of biostimulant on germination was not significant in the other 4 species investigated.

After germination, seedlings displayed species-specific differences in early root development. The earlier development of lateral roots, which was observed in CMO within 2 weeks since seedlings emergence, when the roots system of other species was still made of the taproot only, is consistent with the early-invader ecological behaviour of CMO (Fichtner and Wissemann 2021). The development of thin and slender lateral roots with high SRL allowed CMO to reach similar TRL as FSY, although the investment in root biomass was significantly lower in the former than in the latter species. The elongation of lateral roots occurred in all species in between 2 and 9 weeks after emergence (mid-April to early June) and was followed by an increase in root taper and root density which occurred in all species but LVU between early June and mid-July (9 to 15 weeks), as shown by the decline in SRL from 9 to 15 weeks after emergence. LVU, instead, sustained root elongation during that period which resulted in LVU having higher SRL and TRL than other species at mid-July. This behaviour is consistent with LVU being the most Mediterranean species among those tested. A higher TRL may help LVU to explore the soil thoroughly to prevent the deleterious effects of the Mediterranean summer drought (Guidi et al. 2008). It can be noted that, at this stage, LVU had about 45% higher TRL than FSY, although root biomass was 3-fold higher in FSY than in LVU. To develop dense roots with low SRL and a high cellulose content may be advantageous to species growing on shallow, organic soils in cold, moist habitats such as FSY, where the support function of the roots system may be crucial for the tree to stand in shallow soils (Genet et al. 2005). Data collected at the end of the first growing season (30 weeks) indicate that root biomass is not fully explanatory of the complexity of the root system: although CMO displayed both higher root biomass and TRL than other species, LVU displayed only 15% lower TRL than CMO despite its roots biomass being 65% lower. AOV and CBE displayed, in general through the experiment, lower root development in terms of both root biomass and TRL than other species.

Minor effects were found on root traits because of the biostimulant application. Previous research highlighted that A. nodosum extracts applied at different concentrations failed to increase root biomass and resulted in decreases in TRL at increasing doses of application (MacDonald et al. 2012). These results are consistent with the lack of significant effects on root traits observed in this research, except for SRL, which decreased in plants amended with 2 and 3 kg m−3 of biostimulant, compared to control, on 2 and 80 weeks after seedling emergence. Low SRL has been associated with increased nutrient availability in soils (Ostonen et al. 2007). A longer lifespan and a higher hydraulic conductivity in resource rich environments have been associated with decreasing SRL, but also a lower metabolic activity and capacity to explore resource limited soils in search of water and nutrients (Poorter and Ryser 2015; Weemstra et al. 2020).

The effects of A. nodosum applications on growth of woody plants are still uncertain, because most research was conducted on herbaceous vegetation. Frioni et al. (2021) did not report any positive effect on grapevine vegetative growth following 6 applications of A. nodosum extracts per season, both when the extract was applied through foliar and through soil drench applications. On the contrary, Spinelli et al. (2009) reported increases in vegetative growth in apple trees treated with commercial extract through fertigation. Our results better conform with the latter research and showed that soil amendment with the seaweed extract applied at 2 and 3 kg m−3 transiently increased RGRplant in the very early stages of plant development. Nonetheless, such initial boost allowed plants amended with 3 kg m−3 to achieve a higher plant dry biomass compared to untreated plants throughout the first growing season. Such increase was mainly driven by higher stem dry weight in D3 plants (data not shown), as previously observed in Citrus (Conesa et al. 2020). The lack of significant differences in plant growth observed in the second growing season suggests the need for frequent applications (at least on a yearly basis), if increasing biomass production is the goal of the biostimulant treatment.

Transient increases in biomass production in plants amended with the highest dose of biostimulant was supported by a higher carbon gain, since D3 plants displayed both higher TLA and higher net CO2 assimilation per unit leaf area in the first year after germination. Larger differences in leaf area between plants treated with the highest dose of biostimulant and control were observed at the onset on leaf senescence (October 2021, 30 weeks after emergence). These differences may be due to: (1) direct supply of polyphenols from the extract. Flavonoids and anthocyanins in particular are well known for their antioxidant role and for their capacity to delay leaf physiological senescence and extend leaf lifespan (Renner and Zohner 2019; Agati et al. 2020). It is unclear, however, the mechanism through which polyphenols in the extract can be absorbed and transferred to leaves and whether this occurs; (2) activation of major genes involved in polyphenol biosynthesis such as chalcone isomerase and flavonoid 3-monooxygenase gene, which can be triggered by hormones contained in the biostimulant or by higher carbohydrate availability provided by higher assimilation rate (Calvo et al. 2014; Salvi et al. 2019); (3) direct supply of carbohydrates and amino acids (mainly betaine), which have also been reported to delay chlorophyll degradation and retard leaf senescence (Khan et al. 2009).

In the 3 species on which leaf gas exchange was measured, the application of biostimulant at high doses transiently increased net CO2 assimilation during the first year after emergence. Our data confirm, as hypothesized in previous research, that higher CO2-assimilation in treated plants may be due to transient increases in leaf chlorophyll concentration (Basile et al. 2020), thus being mainly driven by nutritional factors. These factors may include both improved nutrient retention capacity in amended substrate due to alginates and higher nutrient supply to plants due to inorganic ions contained in the extract (Khan et al. 2009). Although minerals in algae extracts are not at a concentration high enough to stimulate growth, when the extract is applied at 3 kg m−3, the supply of nitrogen may be relevant. Changes in diffusive limitations to photosynthesis were also expected because of biostimulant application (Santaniello et al. 2017), but they only transiently occurred in FSY. In this species, biostimulants may have acted as signalling molecules of a lenient stress, thus increasing stomatal limitations to photosynthesis in treated plants. This may be due to the downregulation of the expression of gene MYB60, which is related to stomatal movements, as previously reported in Arabidopsis (Santaniello et al. 2017). At the same time, mesophyll diffusion limitations were reduced by the biostimulant. This may be due to increased expression of PIP1;2 and bCA1 genes (Santaniello et al. 2017), which are related to the biosynthesis of COO-poorins and carbonic anhydrase, 2 crucial facilitators of CO2 diffusion from the substomatal chamber to the chloroplasts (Fini et al. 2016). The reduction of the ratio between mesophyll and stomatal limitations to photosynthesis has been proposed as a key functional trait to promote drought tolerance and water use efficiency (Flexas et al. 2013). However, the effect of biostimulant on this trait was short lived, and a single application of the extract during nursery cultivation may not yield a significant benefit to the plants throughout the establishment period. Further, biostimulant-induced effects on photosynthetic limitations cannot be generalized because they were only significant in one species.

Similarly, the increase in transpiration observed in plants treated with 2 and 3 kg m−3 of A. nodosum extract during the first, but not during the second growing season after emergence, was likely due to the supply of soluble carbohydrates, including mannitol, which have a key osmotic role in plants. Indeed, their active accumulation in plant tissue can lower water potential without hindering turgor, thereby promoting water uptake (Kozlowski and Pallardy 2002; Conesa et al. 2020). On the contrary, we found no evidence that biostimulant application can shift resource allocation to roots in order to promote a more favourable plant water balance in the long run. Neither root to shoot ratio, which has positively been related to drought and transplant stress tolerance (Franco et al. 2006), nor the ratio between TLA and TRL, which may be an even better indication of the amount of absorbing root surface which can supply the transpiring leaf area, were affected by biostimulant application.

Conclusion

In conclusion, our research revealed that the amendment of nursery substrate with a pure A. nodosum extract can accelerate the plant production process by improving seed germination and plant growth if the biostimulant is applied at 3 kg m−3, which is 3 times higher than the label dose, and if it is applied on a regular schedule (annually). Conversely, we found no evidence that the application of biostimulant during the cultivation period in the nursery can induce long term morphological and physiological adjustments which can enhance the tolerance to transplant. Future research may investigate whether A. nodosum application at transplanting may alleviate transplant shock, but the short persistence of the beneficial effects resulting from biostimulant application may lead to the need for multiple applications to be performed during the establishment period, which may not be economically feasible in urban woodlands.

Conflicts of Interest

The authors reported no conflicts of interest.

Acknowledgements

This research received no specific funding and was developed through an agreement between ERSAF (Ente Regionale per i Servizi all’Agricoltura e alle Foreste) and the Department of Agricultural and Environmental Sciences—Landscape, Production, Agroenergy, University of Milan.

Appendix.

View this table:
Table S1.

P-values obtained using mixed models for root traits. Significant (P < 0.05) factors and interactions are highlighted in bold. DWroots (root dry weight); R:S (root to shoot ratio); SRL (specific root length); TRL (total root length per plant).

View this table:
Table S2.

P-values obtained using mixed models for plant growth traits. Significant (P < 0.05) factors and interactions are highlighted in bold. RGRplant (relative growth rate of the whole plant); DWplant (plant dry weight); TLA (total plant leaf area); R:S (root to shoot ratio); TLA:TRL (total leaf area to total root length ratio).

View this table:
Table S3.

P-values obtained using mixed models for leaf gas exchange data. Significant (P < 0.05) factors and interactions are highlighted in bold. E (transpiration); A (net CO2 assimilation); Ca – Ci (CO2 drawdown from the outer air to the substomatal chamber); Ci – Cc (CO2 drawdown from the substomatal chamber to the chloroplasts); SPAD (leaf greenness index).

Figure S1.
Figure S1.

Average monthly temperature (°C, line) and total monthly rainfall (mm, columns) at the experimental site during the years when the experiment was carried out (2021 and 2022).

Literature Cited

  1. ↵
    1. Agati G,
    2. Brunetti C,
    3. Fini A,
    4. Gori A,
    5. Guidi L,
    6. Landi M,
    7. Sebastiani F,
    8. Tattini M.
    2020. Are flavonoids effective antioxidants in plants? Twenty years of our investigation. Antioxidants. 9(11):1908. https://doi.org/10.3390/antiox9111098
    OpenUrl
  2. ↵
    Agrofertil. 2015. Regulalg: 100% algues. Le Mans (France): Agrofertsil S.A.R.L. [Updated 2022 February 9; Accessed 2023 October]. https://www.agrofertil.fr/wp-content/uploads/2015/02/REGULALG.pdf
  3. ↵
    1. Amoroso G,
    2. Frangi P,
    3. Piatti R,
    4. Ferrini F,
    5. Fini A,
    6. Faoro M.
    2010. Effect of container design on plant growth and root deformation of littleleaf linden and field elm. HortScience. 45(12): 1824–1829. https://doi.org/10.21273/HORTSCI.45.12.1824
    OpenUrlAbstract/FREE Full Text
  4. ↵
    1. Amoroso G,
    2. Frangi P,
    3. Piatti R,
    4. Fini A,
    5. Ferrini F,
    6. Faoro M.
    2011. Evaluation of shrubs for side slope greening and protection in urban landscape. HortTechnology. 21(3):359–366. https://doi.org/10.21273/HORTTECH.21.3.359
    OpenUrl
  5. ↵
    1. Basile B,
    2. Rouphael Y,
    3. Colla G,
    4. Soppelsa S,
    5. Andreotti C.
    2020. Appraisal of emerging crop management opportunities in fruit trees, grapevines and berry crops facilitated by the application of biostimulants. Scientia Horticulturae 267:109330. https://doi.org/10.1016/j.scienta.2020.109330
    OpenUrl
  6. ↵
    1. Bowler DE,
    2. Buyung-Ali L,
    3. Knight TM,
    4. Pullin AS.
    2010. Urban greening to cool towns and cities: A systematic review of the empirical evidence. Landscape and Urban Planning. 97(3): 147–155. https://doi.org/10.1016/j.landurbplan.2010.05.006
    OpenUrlCrossRef
  7. ↵
    1. Breger BS,
    2. Eisenman TS,
    3. Kremer ME,
    4. Roman LA,
    5. Martin DG,
    6. Rogan J.
    2019. Urban tree survival and stewardship in a state-managed planting initiative: A case study in Holyoke, Massachusetts. Urban Forestry & Urban Greening. 43:126382. https://doi.org/10.1016/j.ufug.2019.126382
    OpenUrl
  8. ↵
    1. Calvo P,
    2. Nelson L,
    3. Kloepper JW.
    2014. Agricultural uses of plant biostimulants. Plant and Soil. 383:3–41. https://doi.org/10.1007/s11104-014-2131-8
    OpenUrlCrossRef
  9. ↵
    1. Conesa MR,
    2. Espinosa PJ,
    3. Pallarés D,
    4. Pérez-Pastor A.
    2020. Influence of plant biostimulant as technique to harden citrus nursery plants before transplanting to the field. Sustainability. 12(15):6190. https://doi.org/10.3390/su12156190
    OpenUrl
  10. ↵
    1. Conway TM,
    2. Vander Vecht J.
    2015. Growing a diverse urban forest: Species selection decisions by practitioners planting and supplying trees. Landscape and Urban Planning. 138:1–10. https://doi.org/10.1016/j.landurbplan.2015.01.007
    OpenUrl
  11. ↵
    1. Craigie JS.
    2011. Seaweed extract stimuli in plant science and agriculture. Journal of Applied Phycology. 23:371–393. https://doi.org/10.1007/s10811-010-9560-4
    OpenUrl
  12. ↵
    1. Critchley AT,
    2. Critchley JSC,
    3. Norrie J,
    4. Gupta S,
    5. Van Staden J.
    2021. Chapter 13. Perspectives on the global biostimulant market: Applications, volumes, and values, 2016 data and projections to 2022. In: Gupta S, Van Staden J, editors. Biostimulants for crops from seed germination to plant development: A practical approach. 1st Ed. Amsterdam (Netherlands): Elsevier. p. 289–296. https://doi.org/10.1016/B978-0-12-823048-0.00012-5
  13. ↵
    1. De Saeger J,
    2. Van Praet S,
    3. Vereecke D,
    4. Park J,
    5. Jacques S,
    6. Han T,
    7. Depuydt S.
    2020. Toward the molecular understanding of the action mechanism of Ascophyllum nodosum extracts on plants. Journal of Applied Phycology. 32:573–597. https://doi.org/10.1007/s10811-019-01903-9
    OpenUrl
  14. ↵
    1. du Jardin P.
    2012. The science of plant biostimulants—A bibliographic analysis: Final report [Contract 30-CE0455515/0096, Ad hoc study on bio-stimulants products]. Brussels (Belgium): European Commission. 37 p.
  15. ↵
    1. Elansary HO,
    2. Skalicka-Woźniak K,
    3. King IW.
    2016. Enhancing stress growth traits as well as phytochemical and antioxidant contents of Spiraea and Pittosporum under seaweed extract treatments. Plant Physiology and Biochemistry. 105:310–320. https://doi.org/10.1016/j.plaphy.2016.05.024
    OpenUrl
  16. ↵
    1. Elmes A,
    2. Rogan J,
    3. Roman LA,
    4. Williams CA,
    5. Ratick SJ,
    6. Nowak DJ,
    7. Martin DG.
    2018. Predictors of mortality for juvenile trees in a residential urban-to-rural cohort in Worcester, MA. Urban Forestry & Urban Greening. 30:138–151. https://doi.org/10.1016/j.ufug.2018.01.024
    OpenUrl
  17. ↵
    FAO. 2016. The state of world fisheries and aquaculture 2016: Contributing to food security and nutrition for all. Rome: Food and Agriculture Organization of the United Nations. 200 p. https://openknowledge.fao.org/server/api/core/bitstreams/20e618b3-93a1-488a-9697-798f6b6c6b35/content
  18. ↵
    1. Fichtner A,
    2. Wissemann V.
    2021. Biological flora of the British Isles: Crataegus monogyna. Journal of Ecology. 109(1):541–571. https://doi.org/10.1111/1365-2745.13554
    OpenUrl
  19. ↵
    1. Fini A,
    2. Ferrini F,
    3. Frangi P,
    4. Amoroso G,
    5. Giordano C.
    2010. Growth, leaf gas exchange and leaf anatomy of three ornamental shrubs grown under different light intensities. European Journal of Horticultural Science. 75(3):111–117. https://air.unimi.it/retrieve/handle/2434/660357/1266658/ejhs_28-09.pdf
    OpenUrl
  20. ↵
    1. Fini A,
    2. Frangi P,
    3. Amoroso G,
    4. Piatti R,
    5. Faoro M,
    6. Bellasio C,
    7. Ferrini F.
    2011. Effect of controlled inoculation with specific mycorrhizal fungi from the urban environment on growth and physiology of containerized shade tree species growing under different water regimes. Mycorrhiza. 21:703–719. https://doi.org/10.1007/s00572-011-0370-6
    OpenUrl
  21. ↵
    1. Fini A,
    2. Loreto F,
    3. Tattini M,
    4. Giordano C,
    5. Ferrini F,
    6. Brunetti C,
    7. Centritto M.
    2016. Mesophyll conductance plays a central role in leaf functioning of Oleaceae species exposed to contrasting sunlight irradiance. Physiologia Plantarum. 157(1): 54–68. https://doi.org/10.1111/ppl.12401
    OpenUrl
  22. ↵
    1. Flexas J,
    2. Scoffoni C,
    3. Gago J,
    4. Sack L.
    2013. Leaf mesophyll conductance and leaf hydraulic conductance: An introduction to their measurement and coordination. Journal of Experimental Botany. 64(13):3965–3981. https://doi.org/10.1093/jxb/ert319
    OpenUrlCrossRefPubMed
  23. ↵
    1. Franco JA,
    2. Martínez-Sánchez JJ,
    3. Fernández JA, Bañón S.
    2006. Selection and nursery production of ornamental plants for landscaping and xerogardening in semi-arid environments. The Journal of Horticultural Science and Biotechnology. 81(1):3–17. https://doi.org/10.1080/14620316.2006.11512022
    OpenUrl
  24. ↵
    1. Frioni T,
    2. VanderWeide J,
    3. Palliotti A,
    4. Tombesi S,
    5. Poni S,
    6. Sabbatini P.
    2021. Foliar vs. soil application of Ascophyllum nodosum extracts to improve grapevine water stress tolerance. Scientia Horticulturae. 277:109807. https://doi.org/10.1016/j.scienta.2020.109807
    OpenUrl
  25. ↵
    1. Genet M,
    2. Stokes A,
    3. Salin F,
    4. Mickovski SB,
    5. Fourcaud T,
    6. Dumail JF,
    7. van Beek R.
    2005. The influence of cellulose content on tensile strength in tree roots. Plant and Soil. 278:1–9. https://doi.org/10.1007/s11104-005-8768-6
    OpenUrlCrossRef
  26. ↵
    1. Genty B,
    2. Briantais JM,
    3. Baker NR.
    1989. The relationship between the quantum yield of photosynthetic electron transport and quenching of chlorophyll fluorescence. Biochimica et Biophysica Acta (BBA)—General Subjects. 990(1):87–92. https://doi.org/10.1016/S0304-4165(89)80016-9
    OpenUrlCrossRef
  27. ↵
    1. Guidi L,
    2. Degl’Innocenti E,
    3. Remorini D,
    4. Massai R,
    5. Tattini M.
    2008. Interactions of water stress and solar irradiance on the physiology and biochemistry of Ligustrum vulgare. Tree Physiology. 28(6):873–883. https://doi.org/10.1093/treephys/28.6.873
    OpenUrlCrossRefPubMed
  28. ↵
    1. Harley PC,
    2. Loreto F,
    3. Di Marco G,
    4. Sharkey TD.
    1992. Theoretical considerations when estimating the mesophyll conductance to CO2 flux by analysis of the response of photosynthesis to CO2. Plant Physiology. 98(4):1429–1436. https://doi.org/10.1104/pp.98.4.1429
    OpenUrlAbstract/FREE Full Text
  29. ↵
    1. Hilbert DR,
    2. Roman LA,
    3. Koeser AK,
    4. Vogt J, van Doorn NS.
    2019. Urban tree mortality: A literature review. Arboriculture & Urban Forestry. 45(5):167–200. https://doi.org/10.48044/jauf.2019.015
    OpenUrl
  30. ↵
    1. Jeannin I,
    2. Lescure JC,
    3. Morot-Gaudry JF.
    1991. The effects of aqueous seaweed sprays on the growth of maize. Botanica Marina. 34(6):469–474. https://doi.org/10.1515/botm.1991.34.6.469
    OpenUrl
  31. ↵
    1. Jim CY.
    2017. Conservation and creation of urban woodlands. In: Tan PY, Jim CY, editors. Greening cities: Forms and functions. 1st Ed. Springer Singapore. p. 307–330. https://doi.org/10.1007/978-981-10-4113-6_14
  32. ↵
    1. Khan W,
    2. Rayirath UP,
    3. Subramanian S,
    4. Jithesh MN,
    5. Rayorath P,
    6. Hodges DM,
    7. Critchley AT,
    8. Craigie JS,
    9. Norrie J,
    10. Prithiviraj B.
    2009. Seaweed extracts as biostimulants of plant growth and development. Journal of Plant Growth Regulation. 28:386–399. https://doi.org/10.1007/s00344-009-9103-x
    OpenUrlCrossRef
  33. ↵
    1. Koeser AK,
    2. Gilman EF,
    3. Paz M,
    4. Harchick C.
    2014. Factors influencing urban tree planting program growth and survival in Florida, United States. Urban Forestry & Urban Greening. 13(4):655–661. https://doi.org/10.1016/j.ufug.2014.06.005
    OpenUrl
  34. ↵
    1. Kozlowski TT,
    2. Pallardy SG.
    2002. Acclimation and adaptive responses of woody plants to environmental stresses. The Botanical Review. 68(2):270–334. https://doi.org/10.1663/0006-8101(2002)068[0270:AAAROW]2.0.CO;2
    OpenUrlCrossRef
  35. ↵
    Legislative Decree 10 November 2003, n. 386. Implementation of Directive 1999/105/EC on the marketing of forest reproductive material. Gazzetta Ufficiale della Repubblica Italiana. 23(29-01-2004): Ordinary Supplement no. 14. https://www.gazzettaufficiale.it/eli/id/2004/01/29/004G0024/sg
    OpenUrl
  36. ↵
    1. Loreto F,
    2. Centritto M,
    3. Chartzoilakis K.
    2003. Photosynthetic limitations in olive cultivars with different sensitivity to salt stress. Plant, Cell & Environment. 26(4):595–601. https://doi.org/10.1046/j.1365-3040.2003.00994.x
    OpenUrlCrossRef
  37. ↵
    1. MacDonald JE,
    2. Hacking J,
    3. Weng Y,
    4. Norrie J.
    2012. Root growth of containerized lodgepole pine seedlings in response to Ascophyllum nodosum extract application during nursery culture. Canadian Journal of Plant Science. 92(6):1207–1212. https://doi.org/10.4141/cjps2011-279
    OpenUrl
  38. ↵
    1. Moreira R,
    2. Sineiro J,
    3. Chenlo F,
    4. Arufe S, Díaz-Varela D.
    2017. Aqueous extracts of Ascophyllum nodosum obtained by ultrasound-assisted extraction: Effects of drying temperature of seaweed on the properties of extracts. Journal of Applied Phycology. 29:3191–3200. https://doi.org/10.1007/s10811-017-1159-6
    OpenUrl
  39. ↵
    1. Nelson WR,
    2. van Staden J.
    1986. Effect of seaweed concentrate on the growth of wheat. South African Journal of Science. 82:199–200.
    OpenUrl
  40. ↵
    1. Nicese FP,
    2. Ferrini F.
    2008. Competitiveness and environmental sustainability of ornamental nursery. Convention Proceedings “Development of Tuscany’s Nursery”. 2008 May 12; Pistoia, Italy. p. 79–99.
  41. ↵
    1. Ostonen I,
    2. Püttsepp Ü,
    3. Biel C,
    4. Alberton O,
    5. Bakker MR,
    6. Lõhmus K,
    7. Majdi H,
    8. Metcalfe D,
    9. Olsthoorn AFM,
    10. Pronk A,
    11. Vanguelova E,
    12. Weih M,
    13. Brunner I.
    2007. Specific root length as an indicator of environmental change. Plant Biosystems. 141(3): 426–442. https://doi.org/10.1080/11263500701626069
    OpenUrl
  42. ↵
    1. Percival GC,
    2. Keary IP,
    3. Noviss K.
    2008. The potential of a chlorophyll content SPAD meter to quantify nutrient stress in foliar tissue of sycamore (Acer pseudoplatanus), English oak (Quercus robur), and European beech (Fagus sylvatica). Arboriculture & Urban Forestry. 34(2):89–100. https://doi.org/10.48044/jauf.2008.012
    OpenUrl
  43. ↵
    1. Pereira L,
    2. Morrison L,
    3. Shukla PS,
    4. Critchley AT.
    2020. A concise review of the brown macroalga Ascophyllum nodosum (Linnaeus) Le Jolis. Journal of Applied Phycology. 32:3561–3584. https://doi.org/10.1007/s10811-020-02246-6
    OpenUrl
  44. ↵
    1. Poorter H,
    2. Ryser P.
    2015. The limits to leaf and root plasticity: What is so special about specific root length? New Phytologist. 206(4):1188–1190. https://doi.org/10.1111/nph.13438
    OpenUrl
  45. ↵
    1. Rahman MA,
    2. Stratopoulos LMF,
    3. Moser-Reischl A,
    4. Zölch T,
    5. Häberle KH,
    6. Rötzer T,
    7. Pretzsch H,
    8. Pauleit S.
    2020. Traits of trees for cooling urban heat islands: A meta-analysis. Building and Environment. 170:106606. https://doi.org/10.1016/j.buildenv.2019.106606
    OpenUrl
  46. ↵
    1. Rayorath P,
    2. Khan W,
    3. Palanisamy R,
    4. MacKinnon SL,
    5. Stefanova R,
    6. Hankins SD,
    7. Critchley AT,
    8. Prithiviraj B.
    2008. Extracts of the brown seaweed Ascophyllum nodosum induce gibberellic acid (GA3)-independent amylase activity in barley. Journal of Plant Growth Regulation. 27(4):370–379. https://doi.org/10.1007/s00344-008-9063-6
    OpenUrl
  47. ↵
    1. Renner S,
    2. Zohner CM.
    2019. The occurrence of red and yellow autumn leaves explained by regional differences in insolation and temperature. New Phytologist. 224(4):1464–1471. https://doi.org/10.1111/nph.15900
    OpenUrl
  48. ↵
    1. Roman LA,
    2. Battles JJ,
    3. McBride JR.
    2014. Determinants of establishment survival for residential trees in Sacramento County, CA. Landscape and Urban Planning. 129:22–31. https://doi.org/10.1016/j.landurbplan.2014.05.004
    OpenUrl
  49. ↵
    1. Salvi L,
    2. Brunetti C,
    3. Cataldo E,
    4. Niccolai A,
    5. Centritto M,
    6. Ferrini F,
    7. Mattii GB.
    2019. Effects of Ascophyllum nodosum extract on Vitis vinifera: Consequences on plant physiology, grape quality and secondary metabolism. Plant Physiology and Biochemistry. 139:21–32. https://doi.org/10.1016/j.plaphy.2019.03.002
    OpenUrl
  50. ↵
    1. Santaniello A,
    2. Scartazza A,
    3. Gresta F,
    4. Loreti E,
    5. Biasone A,
    6. Di Tommaso D,
    7. Piaggesi A,
    8. Perata D.
    2017. Ascophyllum nodosum seaweed extract alleviates drought stress in Arabidopsis by affecting photosynthetic performance and related gene expression. Frontiers in Plant Science. 8:1362. https://doi.org/10.3389/fpls.2017.01362
    OpenUrl
  51. ↵
    1. Shukla PS,
    2. Mantin EG,
    3. Adil M,
    4. Bajpai S,
    5. Critchley AT,
    6. Prithiviraj B.
    2019. Ascophyllum nodosum-based biostimulants: Sustainable applications in agriculture for the stimulation of plant growth, stress tolerance, and disease management. Frontiers in Plant Science. 10:655. https://doi.org/10.3389/fpls.2019.00655
    OpenUrl
  52. ↵
    1. Spinelli F,
    2. Fiori G,
    3. Noferini M,
    4. Sprocatti M,
    5. Costa G.
    2009. Perspectives on the use of a seaweed extract to moderate the negative effects of alternate bearing in apple trees. The Journal of Horticultural Science and Biotechnology. 84(6):131–137. https://doi.org/10.1080/14620316.2009.11512610
    OpenUrl
  53. ↵
    1. Tattini M,
    2. Loreto F,
    3. Fini A,
    4. Guidi L,
    5. Brunetti C,
    6. Velikova V,
    7. Gori A,
    8. Ferrini F.
    2015. Isoprenoids and phenylpropanoids are part of the antioxidant defense orchestrated daily by drought stressed Platanus × acerifolia plants during Mediterranean summers. New Phytologist. 207(3):613–626. https://doi.org/10.1111/nph.13380
    OpenUrl
  54. ↵
    1. Tennant D.
    1975. A test of a modified line intersect method of estimating root length. Journal of Ecology. 63(3):995–1001. https://doi.org/10.2307/2258617
    OpenUrlCrossRef
  55. ↵
    1. Tkaczyk M,
    2. Szmidla H,
    3. Sikora K.
    2022. The use of biostimulants containing Ascophyllum nodosum (L.) Le Jolis algal extract in the cultivation and protection of English oak Quercus robur L. seedlings in forest nurseries. Sylwan. 166(4):244–252. https://doi.org/10.26202/sylwan.2022032
    OpenUrl
  56. ↵
    1. Van Oosten MJ,
    2. Pepe O,
    3. De Pascale S,
    4. Silletti S,
    5. Maggio A.
    2017. The role of biostimulants and bioeffectors as alleviators of abiotic stress in crop plants. Chemical and Biological Technologies in Agriculture. 4:5. https://doi.org/10.1186/s40538-017-0089-5
    OpenUrl
  57. ↵
    1. Weemstra M,
    2. Kiorapostolou N,
    3. van Ruijven J,
    4. Mommer L,
    5. de Vries J,
    6. Sterck F.
    2020. The role of fine-root mass, specific root length and life span in tree performance: A whole-tree exploration. Functional Ecology. 34(3):575–585. https://doi.org/10.1111/1365-2435.13520
    OpenUrl
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Growth, Physiology, and Root Development in Seedlings of Woody Species Treated with a Seaweed Extract
Sebastien Comin, Gloria Brocca, Noemi Valsecchi, Simone Fumagalli, Irene Vigevani, Denise Corsini, Francesco Ferrini, Giovanni Ravanelli, Alessio Fini
Arboriculture & Urban Forestry (AUF) Sep 2024, jauf.2024.013; DOI: 10.48044/jauf.2024.013
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