Seaweed Polysaccharides as Potential Biostimulants in Turnip Greens Production

Abstract

Seaweed polysaccharides can act as substitutes for synthetic compounds present in commercial stimulants and fertilizers used in agriculture to improve crop yields and vigor. In this study, three different polysaccharides (alginate, agar, and carrageenan) were extracted from one brown seaweed, Saccorhiza polyschides, and two red seaweeds, Gracilaria gracilis and Chondrus crispus, respectively, and applied to potted turnip greens (Brassica napus L.), with the intention to analyze their impact on plant growth, development, and metabolism. Turnip greens treated with polysaccharides, especially carrageenan of C. crispus, showed the best results in improving the crop productivity in terms of plant length and weight, number of leaves, nutrient and pigment content, and soil fertility compared with turnip greens from the negative control or those treated with a commercial leaf fertilizer. λ-carrageenan extracted from the tetrasporophyte generation of C. crispus had the highest bioactivity and positive effect on turnip greens among all treatments. λ-carrageenan has been shown to improve plant growth; increase the plant’s biomass (plant leaves: CC(T) (40.80 ± 5.11 g) compared to the positive control (15.91 ± 15.15 g)) and root system; enhance photosynthetic activity; increase the uptake of soil nutrients; and protect plants against abiotic and biotic stresses, stimulating the production of secondary metabolites and managing its defense pathways. Seaweed-extracted polysaccharides have the potential to be used in sustainable agriculture.

1. Introduction

When compared to commercial fertilizers, seaweed-based extracts have drawn significant interest in agriculture by stimulating the quality and production of different plant crops [1,2]. Currently, there are a few seaweed-based fertilizers available on the market [3]. These biostimulants (seaweed extracts with biostimulant properties) contain bioactive components that regulate phytohormone imbalance, promote soil water retention, reduce nutrient deficiencies (both in the soil and the plant), and increase soil microbiota [2]. Seaweed extracts can also induce responses to pathogens in plants by activating their defensive mechanisms [4]. These extracts can enhance seed germination [5], crop production [6], plant vigor [6], soil nutrient absorption [7], fruit shelf life [6], and plant resistance to a variety of abiotic and biotic challenges [2,8].

Polysaccharides, polyphenols, phytohormones, minerals, and other inorganic and organic bioactive compounds found in seaweed extracts [4] may vary according to the seaweed class, species, and extraction technique. Polysaccharides account for the majority of seaweed biomass and are believed to have an impact on the plant development and protection against diseases, similarly as hormones do [9,10].

With the growing need in the European Union to limit the use of synthetic compounds in agriculture, seaweed polysaccharides are attracting fresh scientific interest as alternatives to these synthetic substances present in commercial plant fertilizers. Although there is currently more research on the use of seaweed extracts and seaweed biomass in agricultural crops, relatively few studies have focused on the influence of seaweed polysaccharides.

Seaweed polysaccharides are economically significant chemicals with several food industry requirements and well-known extraction processes [11,12,13]. Currently, the seaweed polysaccharides with the most commercial interest are alginate, agar, and carrageenan. Alginate is present in brown seaweeds, such as S. polyschides, in the form of alginic acid [14]. Agar and carrageenan are industrially very important for their gelling, thickening, and stabilizing properties, and are extracted from the red seaweed genus Gracilaria or Gelidium and Gigartinales order, respectively, from the phylum Rhodophyta [15,16]. These compounds act as elicitors in plants and can be employed as naturally occurring growth-enhancing, anti-fungal, and anti-viral agents.

On that account, this study aims to understand how polysaccharide-based solutions of alginate, from S. polyschides; agar, from G. gracilis; and three types of carrageenan, from C. crispus, will influence the growth, development, and metabolism of plants, specifically turnip greens. Turnip (B. napus L.) is a crop with recognized economic importance that is mostly cultivated in temperate regions [17]. This plant is widely cultivated as an oil and vegetable crop across the world because it produces edible roots, leaves, stems, buds, flowers, and seeds [18]. It is a member of the genus Brassica and the family Brassicaceae, one of the most significant vegetable families in agriculture [19].

As previously stated, this study attempts to present a full understanding of the action of seaweed polysaccharides in turnip cultivation (and their potential with nutritional and nutraceutical values), and thus to propose a new perspective on harnessing the potential of seaweed-based extracts as plant biostimulants, growth promoters, and potential resistance inducers against biotic and abiotic stresses in sustainable agriculture, without modifying the environment. By elucidating the intricate interplay between the effects of seaweed polysaccharides and the metabolic processes of turnip plants, this research addresses a less-explored area, offering insights that expand current understandings.

2. Materials and Methods

2.1. Harvesting and Preparation of Seaweed Biomass for Extraction

On the 14th of June 2022 (average air temperature around 24 °C), one brown seaweed, S. polyschides, and two red seaweeds, G. gracilis and C. crispus, were collected from tide pools in the intertidal zones of Buarcos Bay, Figueira da Foz (seawater temperature was a maximum of 27 °C and a minimum of 16 °C).

The seaweed species were selected and identified according to their morphological characteristics with taxonomic references [20].

S. polyschides (Phaeophyceae) (Figure 1a) is an annual, opportunistic false kelp with a large and flat stipe and characteristic marginal, undulated wings near the base. Typically, members of this species may reach lengths of 3–4 m. It lives on stony reefs in the ocean’s subtidal and low intertidal zones. Although it cannot compete with the main species in the area, such Laminaria ochroleuca and Saccharina latissima (Phaeophyceae), this opportunistic seaweed colonizes any open spaces in the sea forest, making it a seaweed of enormous economic significance [21]. Portugal, Spain, France, the United Kingdom, and Ireland make up most of its distribution areas. Even though it is an annual seaweed, its biomass is at its highest in the spring and summer [20,22].

The red seaweed G. gracilis (Figure 1b) is frequently used, all over the world, as a source for agar extraction. This seaweed may be found worldwide and lives in temperate waters between 0 and 20 m deep. The environments, such as water temperature, salinity, dissolved nutrients and other abiotic stresses, affect this seaweed variety in its life cycle, growth, and agar content [15]. The natural populations of this seaweed have been dwindling due to the strong demand for its biomass for agar extraction. Seaweed farms are being created to prevent their extinction and to safeguard this important natural resource. This seaweed reproduces readily and has a rapid growth rate [15]. Global cultivation output for the genus Gracilaria in 2019 exceeded 3.5 million tons (fresh weight), primarily in Asian nations [23].

C. crispus is a red seaweed commonly known as Irish moss. It grows in tufts from a discoid holdfast and has a compressed, thin stipe that progressively widens to a flat, repeatedly dichotomously branching frond [20]. These seaweeds can reach a maximum length of 15 cm, and some of them are iridescent underwater. It is mostly found in Portugal’s west coast, in the Faroe Islands, in West Africa, in Spain, in Canada, and in the United States, as well as in the Bering Sea from Russia to Alaska [20]. This seaweed species alternates between two isomorphic life generations that differ in cell wall phycocolloid composition: tetrasporophyte (Figure 1c) and two gametophytes, non-fructified thalli (Figure 1d) and the female gametophyte (Figure 1e) [24]. The carrageenan type extracted from C. crispus tetrasporophyte is λ-carrageenan, and the carrageenan type extracted from C. crispus non-fructified thalli and female gametophyte is a hybrid κ- and ι-carrageenan [25]. Visually, the presence of reproductive structures can differentiate the three generations: tetrasporophytes (presence of tetrasporangia), female gametophytes (presence of cystocarps), and non-fructified thalli (no reproductive structures visible, usually with blue iridescence) [26].

All seaweeds were collected with minimal epiphytes or degraded marks and transported in plastic bags in a cool box to the laboratory. Afterwards, all the seaweeds were transferred to separate trays and washed with seawater to remove sand, epiphytes, and other detritus from their biomass. Then, they were transferred again to another separate trays and washed two times with distilled water to remove the salt. Afterwards, the seaweeds were dried in an air-forced oven (Raypa DAF-135, R. Espinar S.L., Barcelona, Spain) at 60 °C for 48 h. Dried seaweeds were stored in separate silica bags to reduce moisture in the dark and at room temperature (23 °C).

2.2. Polysaccharide Extraction

Each type of polysaccharide (alginate, agar and carrageenan) was extracted according to the methods mentioned in Section 2.2.1, Section 2.2.2 and Section 2.2.3, respectively, and were performed in triplicate.

2.2.1. Alginate

The alginic acid extraction was based on the method described in [27], with modifications. The dried seaweed (S. polyschides, 7 g) (analytical scale: Highland HCB 123, Adam Equipment, Milton Keynes, UK) was milled (particles < 1 cm) with a commercial grinder (TitanMill 300 DuoClean, Cecotec, Valencia, Spain) and then added to a solution of hydrochloric acid (José Manuel Gomes dos Santos, Portugal) at 1.23% (1:30 v:v) (7 mL of hydrochloric acid: 203 mL of distilled water per 7 g of dried seaweed) and kept at room temperature (23 °C) for 24 h. The solution was filtrated, under vacuum (Laborport N820, Lisbon, Portugal), with a Gooch funnel (porosity: G2) and washed with distilled water 2 or 3 times. The obtained residue was alkali-extracted in 2% sodium carbonate (Fisher Chemicals, Porto Salvo, Portugal) (90 mL for the initial weight of the dried biomass; 1:30 m:v) and put in the ultrasound machine (ultrasonic cleaner ULTR-3L2-001, IBX instruments, Barcelona, Spain) at 50 °C for 45 min. The ultrasonication was important for optimizing this part of the process, making it faster and using less reagents than the original method. The extract was filtrated again, under vacuum, through a cloth filter (mesh type 60 approximated 0.25 mm) with a Gooch funnel (porosity: G2) to remove the residues from the alginate solution. Then, 37% hydrochloric acid (José Manuel Gomes dos Santos, Portugal) was added to the filtrate to induce the precipitation of the alginic acid (2 mL of 37% of hydrochloric acid: 30 mL of the final solution). The alginate was washed with ethanol 96% (José Manuel Gomes dos Santos, Portugal) (1:3 v:v) and placed into a cold environment. The liquid solution was discarded, and the precipitate was dried in an air-forced oven (Raypa DAF-135, R. Espinar S.L., Barcelona, Spain) at 60 °C for 48 h.

2.2.2. Agar

Agar extraction was based on the method described in [28] with modifications. The dried seaweed (G. gracilis, 15 g) (analytical scale: Highland HCB 123, Adam Equipment, UK) was added to distilled water (600 mL) and placed into an electric pressure cooker (300008IAU, Aigostar, Madrid, Spain) at a temperature of 115 °C with an air pressure of 80 Kpa for 2 h. The solution obtained was hot-filtrated under vacuum (Laborport N820, Lisbon, Portugal) through a cloth filter (mesh type 60 approximated 0.25 mm) supported in a Buchner funnel. The obtained liquid extract was filtrated again, under vacuum, with a Gooch funnel (porosity: G2). The filtrated solution was solidified at room temperature (23 °C) and frozen overnight in a plastic cup. The next day, the agar was unsolidified, washed, and purified until it became a white or translucid gel. That gel was dried in an air-forced oven (Raypa DAF-135, R. Espinar S.L., Barcelona, Spain) at 60 °C for 48 h.

2.2.3. Carrageenan

Carrageenan extraction was based on the method described in [29], with modifications. The dried seaweed (C. crispus, 1 g) (analytical scale: Highland HCB 123, Adam Equipment, UK) was milled (particles < 1 cm) with a commercial grinder (TitanMill 300 DuoClean, Cecotec, Valencia, Spain) and then pre-treated with an acetone (José Manuel Gomes dos Santos, Portugal):methanol (José Manuel Gomes dos Santos, Portugal) (1:1) solution in a final concentration of 1% (m/v) (final volume: 100 mL; 50 mL acetone: 50 mL methanol) for 16 h at 4 °C to eliminate the organic-soluble fraction. The obtained liquid solution was discarded, and the seaweed residues were dried in an air-forced oven (Raypa DAF-135, R. Espinar S.L., Barcelona, Spain) at 60 °C for about 3–5 min. The dried seaweed was immersed in 150 mL of sodium hydroxide (Applichem Panreac, Chicago, IL, USA) (2%) (1 g of initial seaweed: 150 mL of sodium hydroxide solution) in a hot water bath system (GFL 1003, GFL, Burgwedel, Germany) at 85–90 °C for 3 h. Afterwards, the solution was hot-filtrated under vacuum (Laborport N820, Lisbon, Portugal) through a cloth filter (mesh type 60 approximated 0.25 mm) supported in a Buchner funnel. The liquid extract obtained was filtrated again, under vacuum, with a Gooch funnel (porosity: G2). The filtrated solution was evaporated (rotary evaporator: 2600000, Witeg, Wertheim, Germany) under vacuum, to 1/3 of the initial volume. The carrageenan was precipitated by adding twice (1:3) its volume of ethanol 96%, and then centrifuged (Christ Universal Junior II, Martin Christ, Osterode/Harz, Germany) for 10 min at 4000 rpm. The precipitate was washed again with ethanol 96% (José Manuel Gomes dos Santos, Odivelas, Portugal) and placed in the cold (4 °C for 48 h). Finally, the extract was dried in an air-forced oven (Raypa DAF-135, R. Espinar S.L., Barcelona, Spain) at 60 °C for 48 h.

Due to carrageenan type variation, this method was performed for each sample corresponding to the different life cycle generations (tetrasporophyte, non-fructified thalli, and female gametophyte).

2.3. Physico-Chemical Characterization of Polysaccharides’ Solutions

The dried polysaccharides were milled (particles < 0.05 cm), separately, with a commercial grinder (TitanMill 300 DuoClean, Cecotec, Valencia, Spain) and then distilled water was added (1 mg/mL) under constant agitation (magnetic stirrer hot plate: H20 series, IBX instruments, Barcelona, Spain) until the complete dissolution of the polysaccharides was achieved. Afterwards, each polysaccharide solution was diluted into solutions with different concentrations, 0.5 mg/mL and 0.25 mg/mL, as described in

Table 1. Description of the treatments and their concentrations applied in the assay.

Treatment	Concentration
Negative control (Tap water)	-
Positive control (“Profertil”)	1.5% (v/v)
Alginate solution	0.50 mg/mL
Agar solution	0.50 mg/mL
Carrageenan (tetrasporophyte) solution	0.25 mg/mL
Carrageenan (non-fructified thalli) solution	0.50 mg/mL
Carrageenan (female gametophyte) solution	0.50 mg/mL

. The pH (pH meter: 3310 Jenway, Staffordshire, UK), the electric conductivity (portable conductivity meter: ProfiLine Cond 3310 WTW, Oberbayern, Germany), and the viscosity (DV-E model viscometer, Brookfield, Hadamar-Steinbach, Germany) were determined for each polysaccharide solution at room temperature (23 °C). The viscosity measurement was carried out using a spindle S02 with a speed of 100 rpm [30].

2.4. Experimental Conditions

The assay was performed in 5 L black pots (with diameters of 23 cm at the top, diameters of 16 cm at the base, and 18 cm height) in conditioned substrate (SIRO, Coimbra, Portugal), under greenhouse conditions and with a natural photoperiod, at ESAC (Escola Superior Agrária de Coimbra, Portugal). Nineteen pots were organized in a randomized block design, with two turnip seeds (Flora Lusitana, Cantanhede, Portugal) sown in each pot and a plastic bag underneath to prevent water leakage (Figure 2). The substrate in all pots was fertilized with Blaukorn Classic (Blaukorn Classic 12-8-16 (+3 + TE), Compo-expert, Lisboa, Portugal) and drip irrigation was used for 6 min (±250 mL per pot) 3 times per week.

The treatments and their concentrations applied in this experiment (Table 1) included different polysaccharides-based solutions obtained after the extraction of alginate (from S. polyschides), agar (from G. gracilis), and carrageenan (from three different life cycle generations of C. crispus, tetrasporophyte, non-fructified thalli, and female gametophyte), as described in [30]. The concentration of each polysaccharide-based solution was selected prior to this study [30]; the polysaccharides were characterized and evaluated through seed germination assays. Considering the growth parameters, the germination percentage, and especially the ratios shoot/root weight of all turnip seedlings treated with different concentrations (0.25 mg/mL, 0.50 mg/mL, and 1 mg/mL), the optimum polysaccharide-based solution concentrations for the germination of B. napus L. seeds were determined (Table 1). As a positive control, a commercial leaf biofertilizer, “Profertil” (ADP Fertilizantes, Portugal) was used, with 20% (dry matter) based on seaweed Ascophyllum nodosum, at a concentration of 1.5% (v/v), while as a negative control, tap water was used. All the polysaccharide-based solutions and the positive control were applied by spraying to the foliage (± 3 mL of extract sprayed on each plant; 18 mL per treatment in each application). Each treatment was applied to 6 plants (3 pots with 2 plants each) in 3 repetitions. This assay lasted 63 days (from sowing to plant harvesting), and the treatments were applied two times. The first treatment application was carried out 31 days after the sowing (DAS), when the plants had 3 to 4 real leaves. The second application was carried out 10 days after the first application.

2.5. Growth Parameters of the Obtained Plant Material

The evolution of the plant growth was observed throughout the experiment. Sixty-three days after the sowing, plant material was harvested, washed with tap water, and their roots and leaves were separated and evaluated. The lengths and fresh weights (FWs) of the roots and aerial parts of each sample were measured using a ruler (Shatterless 75 S.50, Molin, Portugal) and an analytical scale (PC2000 Mettler-Toledo, Zurich, Switzerland), respectively. The number of leaves in each plant sample was counted. Afterwards, plants were cut, separating the leaves from the roots, and dried in an air-forced oven (Memmert, Büchenbach, Germany) for 3 days at 65 °C until a constant weight was reached. Afterwards, each sample was cooled for about 2 h, and the dry roots and aerial parts were separately measured, defining their dry weight (DW).

2.6. Turnip Greens’ Physiological and Biochemical Characterization

All the methods described in this chapter were used for the plant material obtained from the respective treatment, as shown in Table 1, separately, and performed in duplicate.

2.6.1. Dry Matter and Ash Content

The dry matter and ash content determination were based on the method described in [31]. The dried aerial parts of the plant samples obtained in Section 2.5 were milled (particles < 1 mm) with a commercial grinder (electric coffee grinder: KG-39, DeLonghi, Treviso, Italy), and approximately 3 g of each sample (analytical scale: AB 204 Mettler-Toledo, Zurich, Switzerland) was placed into a crucible and dried in an air-forced oven (UFB 500, Memmert, Büchenbach, Germany) at 105 °C for 4 h. Then, the samples were placed into a desiccator until a constant weight was reached, then weighted again (analytical scale: AB 204 Mettler-Toledo, Zurich, Switzerland) to calculate the dry matter content.

The dry matter content (DM, g 100 g⁻¹ (%) m/m) at 65 °C was calculated according to the following formula [31]:

$$\mathrm{D}\mathrm{M} \mathrm{a}\mathrm{t} 65 °\mathrm{C} (\mathrm{\%})=\frac{{\mathrm{w}}_1}{{\mathrm{w}}_2}\times 100$$

(1)

where w₁ is the weight of the sample dried at 65 °C (g) and w₂ is the weight of the fresh sample (g).

The dry matter content (DM, g 100 g⁻¹ (%) m/m) at 105 °C was calculated according to the following formula [31]:

$$\mathrm{D}\mathrm{M} \mathrm{a}\mathrm{t} 105 °\mathrm{C} (\mathrm{\%})=\frac{{(m}_3-m_1)}{(m_2-m_1)}\times 100$$

(2)

where m₁ is the crucible weight dried at 105 °C (g); m₂ is the crucible and sample weight dried at 65 °C (g); and m₃ is the crucible and sample weight dried at 105 °C (g).

To assess the ash content, the previous samples dried at 105 °C were placed into an incineration muffle (Induzir, Leiria, Portugal) at 480–500 °C overnight and further cooled in a desiccator, then weighted again (analytical scale: AB 204 Mettler-Toledo, Zurich, Switzerland). The ash content was calculated according to the following formulas [31]:

$$\mathrm{A}\mathrm{s}\mathrm{h}\mathrm{e}\mathrm{s} (\mathrm{\%} \mathrm{d}\mathrm{b})=\frac{{(m}_3-m_1)}{(m_2-m_1)}\times 100$$

(3)

$$\mathrm{A}\mathrm{s}\mathrm{h}\mathrm{e}\mathrm{s} (\mathrm{\%} \mathrm{f}\mathrm{b})=\frac{\mathrm{A}\mathrm{s}\mathrm{h}\mathrm{e}\mathrm{s}(\mathrm{\%} \mathrm{d}\mathrm{b})\times (100-\mathrm{H})}{100}$$

(4)

where % db is the percentage of dried biomass; % fb is the percentage of fresh biomass; m₁ is the crucible weight (g); m₂ is the crucible and sample weight dried at 105 °C (g); and m₃ is the crucible and sample weight incinerated (g).

2.6.2. Total Nitrogen/Protein

The total nitrogen/protein content was determined using the Kjeldahl method [31,32]. In a Kjeldahl tube, approximately 0.5 g (analytical scale: AB 204 Mettler-Toledo, Zurich, Switzerland) of the previously dried matter obtained in Section 2.6.1 was added, and then a Kjeldahl tablet (Fisher Chemicals, Portugal) and 10 mL of sulfuric acid (Chem-Lab NV, Zedelgem, Belgium) were added. The tubes were then placed into a Kjeldahl digester (Bloc Digest 12 Rat 2, JP Selecta, Lisbon, Portugal) at 400 °C for 2 h, under “hotte”. The samples were allowed to cool in the “hotte”, and 50 mL of distilled water was added to each tube and placed into a Kjeldahl distiller (VELP Scientifica, Usmate Velate, MB, Italy). Concurrently, 20 mL of boric acid 2% (Chem-Lab NV, Zedelgem, Belgium) was placed into an Erlenmeyer (one for each sample), then further placed into the Kjeldahl distiller as well. During the distillation process, 50 mL of sodium hydroxide (NaOH) was added to the Kjeldahl tube at 40% (m/v) (Chem-Lab NV, Zedelgem, Belgium). The distilled solution was collected and titrated with hydrochloric acid (HCl) 0.1 M (Chem-Lab NV, Zedelgem, Belgium). Total nitrogen (N, % m/m_dry) was calculated according to the following formula [31]:

$$\mathrm{N}\left(\mathrm{\%}\right)=\frac{\left[\mathrm{H}\mathrm{C}\mathrm{l}\right]\times (V-V0)\times 0.014\times 100}{m}$$

(5)

where [HCl] is the hydrochloric acid concentration (M); V is the volume of HCl spent in sample titration (mL); V0 is the volume of HCl spent in control sample titration (mL); m is the sample weight (g); and 0.014 is the value (g) of N that reacts with 1 mL of HCl 1 mol dm⁻³.

The total protein content was determined by multiplication of the protein conversion factor, 6.25, to the total nitrogen content, as described in [33].

2.6.3. Mineral and Trace Element Characterization

The mineral content was analyzed through the dry mineralization extraction method and assessed using flame atomic absorption spectrometry for the determination of copper (Cu), zinc (Zn), manganese (Mn), iron (Fe), calcium (Ca), magnesium (Mg), potassium (K), and sodium (Na) [34], and molecular absorption spectrometry was used for the determination of phosphorus (P) [35]. With the ashes obtained in Section 2.6.1, acid digestion was performed with hydrochloric acid 20% (v/v) (Chem-Lab NV, Zedelgem, Belgium) in a water bath (Memmert, Büchenbach, Germany) at 100 °C for 30 min. Then, the samples were filtrated with a filter paper (Whatman filter paper cellulose-based ashless types, pore size: 8 µm and diameter of 150 mm, Whatman, Maidstone, UK) into a 50 mL volumetric flask, and the volume was adjusted with distilled water (mother liquor). To a 25 mL volumetric flask, 2.5 mL of the previous solution and 2.5 mL of lanthanum chloride (5%) (Chem-Lab NV, Zedelgem, Belgium) were added, and the volume was adjusted with distilled water (dilution: 1:10). After the necessary dilutions needed to determinate the different elements (1:100 or 1:500), the analysis was carried out using an atomic absorption spectrophotometer (PinAAcle 900T, PerkinElmer, Waltham, MA, USA) equipped with a cathode corresponding to each element. For the phosphorus analysis, 2.5 mL of the mother liquor and 5 mL of ammonium molybdate-vanadate solution in nitric medium (Chem-Lab NV, Zedelgem, Belgium) were added to a 25 mL volumetric flask and adjusted with distilled water. This solution was left overnight at room temperature (23 °C). The next day, the phosphorus analysis was carried out using the molecular absorption spectrophotometer (PYE Unicam, SP6-350, Philips, Portugal) at a wavelength of 650 nm.

2.6.4. FTIR-ATR Analysis

The Fourier-transform infrared spectroscopy—attenuated total reflection (FTIR-ATR) analysis was based on the protocol described in [36]. The dried plant samples obtained from the extraction process in Section 2.6.1 were milled (particles < 0.05 cm) with a commercial grinder (TitanMill 300 DuoClean, Cecotec, Valencia, Spain) and subjected to direct analysis (without humidity) (spectrometer: ALPHA II Compact FT-IR Spectrometer, Bruker, Germany) without any further preparation. All spectra obtained were the average of two independent measurements from 400 to 4000 cm⁻¹ with 24 scans, each at a resolution of 4 cm⁻¹.

2.6.5. Leaf Pigments Content

The detection of pigments was based on the method described in [37]. This process uses thin-layer chromatography (TLC) to separate and determinate the composition of pigments in methanolic extracts and spectrophotometry for the quantitative and qualitative analysis of those pigments.

Thin-Layer Chromatography (TLC)

The dried samples obtained in Section 2.6.1 (0.2 g of total dried leaf biomass) (analytical scale: Highland HCB 123, Adam Equipment, UK) were added to 20 mL of acetone (José Manuel Gomes dos Santos, Portugal): methanol (José Manuel Gomes dos Santos, Odivelas, Portugal) (1:1) solution (final volume: 20 mL; 10 mL acetone: 10 mL methanol), under constant agitation (magnetic stirrer hot plate: H20 series, IBX instruments, Barcelona, Spain) for 30 min. The liquid solution was filtrated, under vacuum, with a Gooch funnel (porosity: G2), and then evaporated (rotary evaporator: 2600000, Witeg, Germany) until all the pigments were adhered to the surface of the round-bottom flask. The pigments were resuspended again with 2 mL of acetone (José Manuel Gomes dos Santos, Portugal): methanol (José Manuel Gomes dos Santos, Portugal) (1:1) solution to obtain a concentrated extract. Afterwards, a silica gel TLC plate (ALUGRAM Xtra SIL G UV254, Macherey-Nagel, Düren, Germany) was activated at 120 °C for 5 min (air-forced oven: Raypa DAF-135, R. Espinar S.L., Barcelona, Spain), and then 20 μL of each concentrated extract was applied. The plate was developed in a chromatography chamber using a petroleum ether (José Manuel Gomes dos Santos, Portugal): acetone (José Manuel Gomes dos Santos, Portugal) solution (7:3 v/v) as the eluent until the front reached a height of 10 cm. The plate was then removed, and the solvent was evaporated at room temperature (23 °C). The pigments were identified by calculating the retention factor (Rf) (Rf = compound migration distance (cm)/distance traveled by the eluent) and comparing with the literature.

Spectrophotometry

The quantitative and qualitative analyses of the leaf pigments were performed via spectrophotometry. After the TLC and the necessary dilution (1:50) of the extracts, the analysis was carried out on the spectrophotometer (UV-3100PC, VWR, Lutterworth, UK), with scanning at 665.2 nm, 652.4 nm, 535 nm, and 470 nm. The following formulas were used for the quantification of the pigments (mg/100 g) chlorophyll a and b (Chl a and Chl b), as well as the carotenoids [38]:

$$\mathrm{C}\mathrm{h}\mathrm{l} a=16.75\times A_{665.2}-9.16\times A_{652.4}$$

(6)

$$\mathrm{C}\mathrm{h}\mathrm{l} b=34.09\times A_{652.4}-15.28\times A_{665.2}$$

(7)

$$\mathrm{C}\mathrm{a}\mathrm{r}\mathrm{o}\mathrm{t}\mathrm{e}\mathrm{n}\mathrm{o}\mathrm{i}\mathrm{d}\mathrm{s}=\frac{(1000\times A_{470}-1.63\times \mathrm{C}\mathrm{h}\mathrm{l} a-104.96\times \mathrm{C}\mathrm{h}\mathrm{l} b)}{221}$$

(8)

where A_665.2, A_652.4, and A₄₇₀ are the absorbance of the sample at the wavelengths 665.2 nm, 652.4 nm, and 470 nm, respectively.

The total anthocyanins (mg/100 g) were calculated according to the formula [39]:

$$\mathrm{A}\mathrm{n}\mathrm{t}\mathrm{h}\mathrm{o}\mathrm{c}\mathrm{y}\mathrm{a}\mathrm{n}\mathrm{i}\mathrm{n}\mathrm{s}=\frac{100\times A_{535}\times DF\times V}{98.2\times x}$$

(9)

where A₅₃₅ is the absorbance of the sample at the wavelength 535 nm; DF is the dilution factor; V is the volume of anthocyanin extract that was made up after extraction (ml); and x is the weight of the dried sample used for extraction (g).

2.7. Substrate Characterization

2.7.1. Substrate Density

The initial substrate (before the treatments) and final substrates (after the treatments) used for turnip potting were initially analyzed using the apparent compact density method [40] to measure the densities of the substrate samples. This step is essential for calculating the weight of the substrate sample (m_s, g), which was used in Section 2.7.2, at 60 mL. To a 1000 mL plastic graduated cylinder, the substrate sample was added without pressing it down. Then, the substrate was compacted by dropping the graduated cylinder 10 times onto a 5 mm thick rubber blanket from a height of about 10 cm. The level of the substrate was marked, and the graduated cylinder was weighted (technical scale: UFB 500, Memmert, Büchenbach, Germany). The apparent compact density of the substrate (D_s, g L⁻¹); was calculated according to the following formula [40]:

$$D_s=\frac{m_A-m_B}{V}$$

(10)

where m_A is the weight of the compacted substrate and the graduated cylinder (g); m_B is the weight of the graduated cylinder (g); and V is the final volume of the substrate in the graduated cylinder (L).

The weight of the substrate sample (m_s, g) at 60 mL, used as described in Section 2.7.2, was calculated according to the following formula [40]:

$$m_s=\frac{D_s\times 60}{1000}$$

(11)

where D_s is the apparent compact density of the substrate (g L⁻¹).

2.7.2. pH and Electrical Conductivity

The substrate samples were weighted (technical scale: UFB 500, Memmert, Büchenbach, Germany) in a 500 mL Erlenmeyer, 300 mL of distilled water was added, the cap was secured, and they were shaken for 1 h at 200 rpm (shaking machine: Rotabit, JP Selecta, Lisbon, Portugal) at room temperature (23 °C). Afterwards, the pH was determined (pH meter: 3310 Jenway, Staffordshire, UK) directly from the obtained solution. The electric conductivity (portable conductivity meter: ProfiLine Cond 3310 WTW, Oberbayern, Germany) was determined from the filtrated obtained as described in Section 2.7.3.

2.7.3. Mineral and Trace Element Characterization

The soil extract obtained in Section 2.7.2 was filtrated with filter paper (Whatman filter paper cellulose-based ashless types, pore size: 8 µm and diameter of 150 mm, Whatman, Maidstone, UK), discarding at least the first 10 mL. The rest of the filtrated extract was added to a 100 mL plastic container and stored at room temperature (23 °C). The mineral content was assessed using flame atomic absorption spectrometry for the determination of copper (Cu), zinc (Zn), manganese (Mn), iron (Fe), calcium (Ca), magnesium (Mg), potassium (K), and sodium (Na) [34], and molecular absorption spectrometry for the determination of phosphorus (P) [35] was performed as described in Section 2.6.3.

2.7.4. Organic Matter Content

The substrate samples were placed into separate aluminum trays, weighted, and then dried in an air-forced oven (UFB 500, Memmert, Büchenbach, Germany) at 75 °C for 2 days until reaching a constant weight. Then, the samples were weighted again and milled, separately, in a soil grinder (FRITSCH GmbH Pulverisette 8, Midland, ON, Canada) through a sieve of 1.5 mm, separating the thin (particles < 1.5 mm) and rough (particles > 1.5 mm) material [41,42]. Approximately 3 g of each sample (particles < 1.5 mm) (analytical scale: AB 204 Mettler-Toledo, Zurich, Switzerland) was placed into a crucible and dried in an air-forced oven (UFB 500, Memmert, Büchenbach, Germany) at 105 °C for 4 h. Then, the samples were placed into a desiccator until a constant weight was reached, then weighted again (analytical scale: AB 204 Mettler-Toledo, Zurich, Switzerland). The previous samples dried at 105 °C were placed into an incineration muffle (Induzir, Leiria, Portugal) at 480–500 °C overnight and further cooled in a desiccator, then weighted again (analytical scale: AB 204 Mettler-Toledo, Zurich, Switzerland).

The organic matter content (OM, %) was calculated according to the following formula [41,42]:

$$\mathrm{O}\mathrm{M}\left(\mathrm{\%}\right)=\frac{{(m}_2-m_3)}{(m_2-m_1)}\times 100$$

(12)

where m₁ is the crucible weight (g); m₂ is the crucible and sample weight after drying at 105 °C (g); and m₃ is the crucible and sample weight after being incinerated (g).

2.7.5. Total Nitrogen

A total nitrogen analysis of each substrate was performed as described in Section 2.6.2 with 1 g of the dried samples (at 75 °C) obtained in Section 2.7.4.

The reagent blank test was carried out in parallel with the determinations using the same procedure as that outlined in Section 2.6.1, Section 2.6.2, Section 2.6.3, Section 2.7.2, Section 2.7.3, Section 2.7.4 and Section 2.7.5, using the same quantities of all the reagents as in the determination, but omitting the test portion.

2.8. Statistical Analysis

The statistical analysis was performed using the software Sigma Plot v.14. Data were checked for normality (Shapiro–Wilk test) and homogeneity (the equal variance Brown–Forsythe test). The Holm–Sidak method was used in the analysis when the normality test was rejected. One-way analysis of variance (ANOVA) was then performed to assess statistically significant differences between the growth parameters of each polysaccharide solution. The statistical analysis was performed by comparing the different treatments, and the results were considered statistically different when the p-value was <0.05. The Tukey multiple comparison t-test was used after the rejection of the one-way ANOVA null hypothesis (Holm–Sidak method).

3. Results

3.1. Biostimulant and Biofertilizer Assay in B. napus L.

3.1.1. Biochemical Characterization of the Treatments Applied

The determination of the pH, electrical conductivity (EC), and viscosity of the biostimulant treatments applied to crops can help to anticipate how the plant will accept it and how it will affect the crop yield, quality, and pathogen resistance. The positive control was the treatment with the most neutral pH (

Table 2. pH, electrical conductivity (EC), and viscosity values of the treatments used in the biostimulant and biofertilizer assay of potted turnip. Negative control—tap water. Positive control—“Profertil”.

Treatment	Concentration (mg/mL)	pH	EC (µS/cm)	Viscosity (mPa.s)
Negative control	-	5.86	302	1.00
Positive control	1.5% (v/v)	7.30	1685	5.10
Alginate solution	0.50	3.70	109	3.60
Agar solution	0.50	5.83	73	8.40
Carrageenan (tetrasporophyte) solution	0.25	9.34	100	10.80
Carrageenan (non-fructified thalli) solution	0.50	9.56	184	9.00
Carrageenan (female gametophyte) solution	0.50	9.86	191	9.00

Measured at room temperature (20–22 °C).

). The negative control, the alginate solution, and the agar solution presented acidic pH values. The pH values of the carrageenan solutions were between 9 and 10, which is considered as alkaline. Regarding the EC, the positive control had the highest values among all treatments, whereas the agar solution had the lowest. Between all carrageenan solutions, EC increased with the rise in pH. The viscosity was the highest in the tetrasporophyte solution (the lowest pH and EC). Overall, excluding the negative control, all carrageenan solutions presented the highest viscosity values and the lowest alginate solutions.

3.1.2. Turnip Greens Parameters

By the end of the experiment (day 63), it was possible to observe differences among the potted turnip greens from each treatment (Supplementary Figures S1 and S2). Plants treated with the negative (NC) and positive controls (PC) reduced significantly in size when compared to the other treatments. The samples that presented the most robust turnip leaf development were the ones treated with the carrageenan-based solutions of C. crispus (tetrasporophyte, non-fructified thalli, and female gametophyte; CC(T), CC(NF), and CC(FG), respectively). Additionally, at day 42 (when the second application was made), the NC and PC samples started to exhibit various injuries (as holes) on the leaves, showing the existence of some parasite on the leaves (Supplementary Figure S3). The same was not observed on the treated leaves until day 63, when some injuries appeared, but in very reduced number compared to NC and PC plants (Supplementary Figure S1). Later, it was found that these leaf injuries were caused by Agrotis spp. larvae (Supplementary Figure S3).

After the harvesting of all turnip samples, the growth parameters were evaluated (leaf weight and length, root weight and length) (Figure 3), and the ratios of these parameters were analyzed (

Table 3. Ratios between the aerial part (AP) and root (R) of the fresh turnip greens from each treatment in the biostimulant and biofertilizer assay, measured after 63 days. NC—negative control; PC—positive control; SP—S. polyschides; GG—G. gracilis; CC(T)—C. crispus (tetrasporophyte); CC(NF)—C. crispus (non-fructified thalli); CC(FG)—C. crispus (female gametophyte).

Sample	AP Length/Weight	R Length/Weight	AP Weight/R Weight	AP Length/R Length
NC	1.46	23.27	36.20	2.28
PC	1.68	20.45	26.09	2.15
SP	1.03	11.91	35.04	3.02
GG	1.20	11.94	29.77	3.00
CC(T)	0.92	14.73	43.69	2.74
CC(NF)	1.12	24.01	53.54	2.49
CC(FG)	1.03	14.94	36.62	2.53

).

In Figure 3, the average values of the aerial part weight (Figure 3a), leaf length (Figure 3b), root weight (Figure 3c), and root length (Figure 3d) of the fresh turnips from each treatment are presented. The most robust samples with the statistically significant highest leaf weight (p < 0.05) and statistically significant leaf length (p < 0.05), in relation to the controls, were observed in plants treated with CC(T), CC(NF), and CC(FG) for weight, and with CC(NF) and CC(FG) for length. The samples with the lowest leaf weight (Figure 3a) and length (Figure 3b) were the control ones, particularly the PC, with 15.91 ± 15.15 g and 26.80 ± 6.70 cm, respectively. When comparing the turnip greens treated only with the polysaccharide-based solutions, the turnips that exhibited the best leaf weights were ones treated with the CC(T) (40.80 ± 5.11 g). The turnips treated with the GG presented the lowest leaf weights (29.53 ± 13.99 g). The turnip greens that exhibited the longest leaves were those treated with the CC(FG) (39.98 ± 4.00 cm). The shortest leaves were observed in turnip samples treated with the SP (35.36 ± 3.41 cm).

The heaviest statistically different (p < 0.05) roots were observed in plants treated with the CC(FG) (1.06 ± 0.24 g), but only in relation to the samples with the lightest roots: 0.61 ± 0.70 g for the PC and 0.54 ± 0.40 g for the NC. No statistically significant differences were found in the length of the roots among the treatment groups (p > 0.05). However, the obtained data showed that the samples with the longest roots were the ones treated with the carrageenan CC(FG) (15.80 ± 3.33 cm), CC(NF) (15.77 ± 6.13 cm), and CC(T) (13.76 ± 2.24 cm). Plants treated with SP and GG developed the shortest roots, measuring 11.72 ± 0.89 and 11.84 ± 2.42 cm, respectively.

Regarding the leaf number (Figure 4), plants with the lowest leaf numbers were the controls, with ± 6 leaves for the NC and ± 7 leaves for the PC. The plants that developed the highest significantly different (p < 0.05) leaf numbers (but only in relation to NC) were the ones treated with CC(T), CC(NF), and CC(FG), with ± 9 leaves.

Regarding the ratios between the growth parameters (Table 3), the ratio of aerial part weight to root weight was higher in CC(NF) (53.54), CC(T) (43.69), and CC(FG) (36.62), and lower in PC (26.09) and GG (29.77). Furthermore, the NC and PC presented the lowest ratios of aerial part length to root length, with 2.28 and 2.15, respectively.

3.1.3. Physiological and Biochemical Characterization of Turnip Greens

Mineral and Trace Element Characterization

Mineral and trace element characterization of the turnip edible section (leaves) (

Table 4. Dry matter, ashes, protein, mineral, and trace element characterization of the turnip greens within each treatment. The results are expressed as mean ± standard deviation (n = 2, dry weight basis). Significant differences found among the different samples are marked with different letters (p < 0.05). NI—no information found in the literature. NC—negative control. PC—positive control. SP—S. polyschides; GG—G. gracilis; CC(T)—C. crispus (tetrasporophyte); CC(NF)—C. crispus (non-fructified thalli); CC(FG)—C. crispus (female gametophyte).

Treatments	NC	PC	SP	GG	CC(T)	CC(NF)	CC(FG)	Literature Values	Reference
Dry matter (%)	5.30 ± 0.00 ab	5.34 ± 0.01 ab	3.42 ± 2.31 b	7.19 ± 1.68 a	5.80 ± 0.04 ab	6.66 ± 0.05 a	6.77 ± 0.43 a	6.00	[33]
Ashes (%)	22.74 ± 0.02 a	20.45 ± 0.07 a	19.52 ± 0.02 a	19.36 ± 0.28 a	20.23 ± 0.05 a	19.47 ± 0.07 a	18.48 ± 0.10 a	13.50	[33]
N (%)	5.68 ± 0.05 a	5.27 ± 0.12 ab	5.23 ± 0.06 ab	4.86 ± 0.01 b	5.37 ± 0.03 ab	5.16 ± 0.03 b	4.88 ± 0.05 b	3.23	[43]
Protein (%)	35.50 ± 0.31 a	32.91 ± 0.72 ab	32.66 ± 0.34 ab	30.34 ± 0.03 b	33.56 ± 0.19 ab	32.25 ± 0.19 ab	30.47 ± 0.34 b	33.33	[33]
P (%)	0.84 ± 0.00 ab	0.87 ± 0.01 a	0.81 ± 0.01 ab	0.73 ± 0.02 b	0.80 ± 0.00 ab	0.75 ± 0.01 b	0.76 ± 0.00 b	0.75	[33]
Ca (%)	1.46 ± 0.01 a	1.33 ± 0.01 ab	1.28 ± 0.20 ab	1.23 ± 0.05 ab	1.29 ± 0.00 ab	1.23 ± 0.03 ab	1.20 ± 0.02 b	1.67	[33]
Mg (%)	0.30 ± 0.01 a	0.27 ± 0.00 ab	0.23 ± 0.01 b	0.24 ± 0.00 b	0.26 ± 0.01 ab	0.28 ± 0.01 ab	0.27 ± 0.01 ab	0.17	[33]
K (%)	8.58 ± 0.16 a	7.31 ± 0.12 ab	8.23 ± 0.17 ab	7.43 ± 0.29 ab	8.17 ± 0.07 ab	6.88 ± 0.02 b	7.24 ± 0.17 b	5.00	[33]
Na (%)	0.39 ± 0.03 a	0.41 ± 0.00 a	0.44 ± 0.00 a	0.34 ± 0.03 a	0.39 ± 0.10 a	0.90 ± 0.52 a	0.36 ± 0.02 a	0.67	[33]
Cu (mg/kg)	35.25 ± 0.15 ab	36.80 ± 0.50 ab	33.25 ± 0.45 b	34.70 ± 0.20 ab	35.50 ± 0.80 ab	35.40 ± 1.00 ab	38.00 ± 0.30 a	NI	NI
Zn (mg/kg)	118.15 ± 1.25 a	81.00 ± 1.60 ab	77.05 ± 0.15 b	77.75 ± 1.15 b	81.10 ± 1.50 ab	77.25 ± 0.15 b	81.85 ± 0.65 ab	87.40	[43]
Fe (mg/kg)	149.40 ± 1.30 a	99.35 ± 0.25 ab	93.70 ± 6.80 ab	96.75 ± 0.75 ab	91.50 ± 3.20 b	94.95 ± 2.45 ab	91.25 ± 1.35 b	NI	NI
Mn (mg/kg)	119.75 ± 2.15 a	58.80 ± 2.60 b	89.50 ± 0.20 ab	78.15 ± 1.25 b	92.90 ± 2.30 ab	93.25 ± 1.45 ab	71.90± 1.40 b	98.70	[43]

) was important in order to understand how the treatments applied to the plants affected their nutritional quality. Significant differences between the treatments are marked with different letters (p < 0.05).

All treatments induced a slightly higher production (%) of nitrogen (N) content in all plants (Table 4) when compared to the literature value, with NC having the greatest value among all treatments. The phosphorus (P) content was slightly higher than that cited in the literature in all treatment groups, except in the plants treated with GG and CC(NF). The calcium (Ca) content was lower in all treatment groups when compared with the literature. The sodium (Na) content was lower in all treatment groups when compared to the literature, except in plants treated with CC(NF), where the Na percentage was higher than in all the other treatments. Magnesium (Mg) and potassium (K) contents were higher in all treatment groups compared to the literature.

Regarding the trace elements zinc (Zn), iron (Fe), and manganese (Mn), excluding copper (Cu), there was a considerable difference between their contents in NC plants and in the other treatments. For Zn and Mn, all values were slightly lower than those found in the literature, except for the NC. Overall, the NC exhibited the highest values in all mineral and trace elements. It should be consider that there are no published values regarding the Cu and Fe contents in any plants or whether they are safe for human consumption.

Biochemical Characterization of Turnip Greens

The FTIR-ATR spectra of the turnip greens, in the range of 4000 to 400 cm⁻¹, within each treatment are given in Figure 5 and

Table 5. FTIR-ATR band identification and characterization of the turnip greens within each treatment (dry basis). nd—not detectable. sh—shoulder. NA—not available. NC—negative control. PC—positive control. SP—S. polyschides; GG—G. gracilis.; CC(T)—C. crispus (tetrasporophyte); CC(NF)—C. crispus (non-fructified thalli); CC(FG)—C. crispus (female gametophyte).

Reference Wave Number (cm−1)	Bond	Wave Number Observed (cm−1)
		CC (FG)	CC (NF)	NC	GG	SP	CC (T)	PC
3334	Cellulose	sh	sh	sh	sh	3286	3278	3274
2917	Cellulose	2921	2921	2921	2921	2921	2920	2921
1734	Pectins with ester	1736	1736	1735	1736	1736	1736	1736
1626	Free carboxyl groups	1620	1621	1624	1619	1622	1622	1617
1520	Lignin and phenolic backbone	sh	sh	1540	sh	sh	sh	sh
1371–1314	Cellulose and xyloglucan	1377	1377	1351	1375	1376	1376	1352
1234	Proteins	1240	1240	1238	1239	1239	1239	1238
1015	Polysaccharides, sugars, and pectins	1021	1021	1023	1019	1020	1019	1016
825	NA	825.3	825	824.8	825.5	825.2	825.1	825.6
770	Phenyl groups	nd	nd	nd	nd	nd	nd	nd

. The spectra of the different treatments are aligned according to peak intensity (from highest to lowest). All the spectra (Figure 5) had similar peaks (shoulder), except the peak assigned to the lignin and phenolic backbone at 1520 cm⁻¹, which was only present in the negative control spectra. The characteristic peaks of cellulose were present in all spectra around 3280 cm⁻¹ and 2921 cm⁻¹. The bands corresponding to pectins, with ester, free carboxyl groups, cellulose and xyloglucan, and proteins, were present in all spectra around 1736 cm⁻¹, 1620 cm⁻¹, 1352–1377 cm⁻¹, and 1239 cm⁻¹. The peaks with the highest intensity in all spectra were assigned to polysaccharides, sugars and pectins, at 1020 cm⁻¹. The peak around 825 cm⁻¹ was not assigned to any specific bond, but its significant intensity was noted. None of the spectra reached a significant peak at the 770 cm⁻¹ band, corresponding to the phenyl groups.

Pigment Content

A TLC of the methanolic extracts from each treatment sample of turnip greens is presented in Figure 6. This chromatography separates the different compounds according to their molecular weight. The solvent runs upwards, from non-polar compounds (origin of the pigments) to polar compounds (solvent front). The retention factor (Rf) was used to compare and to help to identify the compounds. The Rf values observed in the different samples are demonstrated in

Table 6. Pigment identification from each treatment sample of turnip greens (dry basis). Rf—retention factor. NI—no information found in the literature. NC—negative control. PC—positive control. SP—S. polyschides; GG—G. gracilis; CC(T)—C. crispus (tetrasporophyte); CC(NF)—C. crispus (non-fructified thalli); CC(FG)—C. crispus (female gametophyte).

				Rf Observed
Nº *	Visible Color	NC	PC	SP	GG	CC (T)	CC (NF)	CC (FG)	Rf Literature	Pigment	Reference
1	light green	0.02	0.02	0.03	0.03	0.02	0.03	0.04	NI	NI	NI
2	light grey	0.07	0.08	0.09	0.10	0.10	0.12	0.14	NI	NI	NI
3	light yellow	nd	nd	0.16	0.17	0.20	0.20	0.22	0.18	Neoxanthin	[44]
4	bright yellow	0.57	0.57	0.57	0.57	0.68	0.68	0.69	0.15–0.35	Xanthophyll	[45,46]
5	light green	0.62	0.62	0.62	0.62	0.73	0.73	0.75	0.32–0.42	Chlorophyll b	[45,46]
6	faded green	0.66	nd	0.66	0.66	0.77	0.77	0.79	NI	NI	NI
7	dark green	0.74	0.74	0.74	0.74	0.84	0.83	0.84	0.44–0.59	Chlorophyll a	[45,46]
8	light grey	0.83	nd	0.83	nd	0.91	0.91	nd	0.49	Pheophytin b	[45]
9	dark grey	0.91	0.91	0.91	0.91	0.95	0.95	0.95	0.60	Pheophytin a	[45]
10	light grey	0.95	0.95	0.95	0.95	nd	nd	nd	NI	NI	NI
11	golden	0.99	0.99	0.99	0.99	0.98	0.98	0.98	0.95–0.98	β-carotene	[44,45,46]

* Corresponding numbers in Figure 6.

, as well as a comparison of Rf values from the literature and the referred pigment. From the wavelength absorbance of each sample in the corresponding pigments (chlorophyll a, chlorophyll b, anthocyanins, and carotenoids), it was possible to quantify them (

Table 7. Pigment quantification (mg/100 g) from each treatment sample of turnip greens. The results are expressed in mean ± standard deviation (n = 3, Dry weight basis). Significant differences between the treatments are marked with different letters (p < 0.05). NC—negative control. PC—positive control. SP—S. polyschides; GG—G. gracilis; CC(T)—C. crispus (tetrasporophyte); CC(NF)—C. crispus (non-fructified thalli); CC(FG)—C. crispus (female gametophyte).

Pigments (mg/100 g)	NC	PC	SP	GG	CC(T)	CC(NF)	CC(GF)
Chlorophyll a	4.346 ± 0.01 b	4.458 ± 0.01 ab	5.233 ± 0.01 ab	4.303 ± 0.01 b	6.916 ± 0.01 a	5.516 ± 0.01 ab	5.914 ± 0.01 ab
Chlorophyll b	1.503 ± 0.01 ab	1.399 ± 0.01 b	1.729 ± 0.01 ab	1.361 ± 0.01 b	2.301 ± 0.01 a	1.841 ± 0.01 ab	1.851 ± 0.01 ab
Anthocyanins	0.011 ± 0.01 a	0.010 ± 0.01 a	0.011 ± 0.01 a	0.009 ± 0.01 a	0.016 ± 0.01 a	0.012 ± 0.01 a	0.014 ± 0.01 a
Carotenoids	0.936 ± 0.01 b	1.013 ± 0.01 b	1.231 ± 0.01 ab	1.056 ± 0.01 ab	1.448 ± 0.01 a	1.230 ± 0.01 ab	1.426 ± 0.01 ab

). After comparing the usual order of the pigments in a TLC and their characteristic colors, pigments were assigned to each number (Table 6).

Samples from CC(T), CC(NF), and CC(FG) moved more than the other samples (Figure 6), and, therefore, had generally higher Rf values (Table 6). Additionally, the pigment marked as “3” in Figure 6 corresponding to neoxanthin (Table 6), did not appear in the TLC of the NC and PC. On the other hand, the pigment marked as “10” only appeared in the TLC of the NC, PC, SP, and GG. The pigment marked as “8” in Figure 6, corresponding to pheophytin b (Table 6), was very difficult to identify clearly due to its brighter color, and it was almost absent in the TLC of the PC, GG, and CC(FG). There is no information published regarding pigments 1, 2, 6, and 10.

CC(T) presented the highest significantly different (p < 0.05) values in Chlorophyll a and Chlorophyll b in turnip greens (Table 7). Regarding the values in Anthocyanins, no significant differences were found, but plants threated with CC(T) and the carotenoids had the highest values in relation to the control-treated plants (NC and PC). Overall, all the treatments from C. crispus increased the pigments in the turnip greens.

3.2. Substrate Characterization

The initial substrate (negative control) and final substrates (after the treatments) used for the potting of the turnip greens was analyzed (

Table 8. Apparent compact density (Ds), weight of the sample (ms) at 60 mL, organic matter content (OM), and nitrogen content (N) of the substrates used for potting of turnip greens for each treatment. The results are expressed as the mean ± standard deviation (n = 2). Significant differences between the treatments are marked with different letters (p < 0.05). NC—negative control. PC—positive control. SP—S. polyschides; GG—G. gracilis; CC(T)—C. crispus (tetrasporophyte); CC(NF)—C. crispus (non-fructified thalli); CC(FG)—C. crispus (female gametophyte).

Soil Sample	NC	PC	SP	GG	CC(T)	CC(NF)	CC(FG)
Ds (g/L)	945.19 ± 29.58 a	804.97 ± 14.01 ab	767.59 ± 10.83 b	771.10 ± 4.21 b	836.16 ± 68.54 ab	802.81 ± 4.18 ab	837.85 ± 16.07 ab
ms at 60 mL (g)	56.71 ± 1.77 a	48.30 ± 0.84 a	46.06 ± 0.65 a	46.27 ± 0.25 a	50.17 ± 4.11 a	48.17 ± 0.25 a	50.27 ± 0.96 a
OM (%)	34.97 ± 1.33 a	24.17 ± 0.73 b	27.22 ± 1.89 ab	23.52 ± 0.50 b	19.54 ± 1.16 b	27.67 ± 7.40 b	23.53 ± 0.84 b
N (%)	0.42 ± 0.01 a	0.40 ± 0.02 a	0.44 ± 0.02 a	0.41 ± 0.02 a	0.35 ± 0.00 a	0.33 ± 0.01 a	0.37 ± 0.03 a

, Supplementary Figures S4–S10). A clear difference was observed between the NC (initial substrate) and the final substrates. The NC had the highest statistically different (p < 0.05) Ds and OM values when compared with the substrates from the other treatments (Table 8). Also, NC produced the heaviest sample weight (ms), but there was no significance between the treatments. The substrate sample with the highest N content resulted from SP treatment, and the lowest N contents were observed in CC(GF), CC(T), and CC(NF), without any significant differences between them. When comparing the initial substrate (NC) with the final substrate of each treatment, there was a slightly decrease in the pH (Supplementary Figure S4), but an increase in the EC values (Supplementary Figure S5). The NC had a neutral pH, and the other substrates had more acidic pH values. The EC of the NC had no saline effects, whereas the other substrates had slight saline effects, with PC having the highest value.

All figures (Supplementary Figures S6–S10) are divided into fertility classes (very low to very high) according to (Laboratório Químico Agrícola Rebelo da Silva, 1977) [46]. Regarding the P₂O₅ content, NC and CC(NF) demonstrated low fertility levels, and PC exhibited high fertility levels. The other treatments presented moderate levels of soil fertility. Considering the CaO and MgO contents, all treatments exhibited very high fertility levels, except the NC. The CC(GF) showed the highest values for CaO and MgO. Regarding the K₂O content, all treatments presented very high fertility levels.

4. Discussion

In agriculture, clear determination of the right values from pH and electrical conductivity (EC) of the biostimulants applied to a crop can aid in the absorbance of their nutrients from the plants, anticipating crop yield, quality, and pathogen resistance [47]. The typical pH for an alginate in solution has values between 2.0 and 3.5 [12]. The pH of the carrageenan solutions is usually between 8 and 11 [11]. There has not been any information published regarding the ideal pH for agar solutions. In this work, the positive control exhibited an increased EC, which can be explained by the composition of the commercial leaf biofertilizer “Profertil” (ADP Fertilizantes, Portugal), containing 20% (dry matter) of the seaweed A. nodosum. The most abundant elements in A. nodosum are potassium, sodium, and calcium [48]. These elements can increase the salinity of the solution and, therefore, increase the EC.

Viscosity, i.e., the fluid’s resistance to flow, is considered one of the most important physical properties used to assess the gelling capabilities of polysaccharides [49]. This property depends on the degree of polymerization, temperature, concentration, molecular weight, and the presence of polyvalent metal cations in the polysaccharide structure [12]. According to EFSA (Panel on Food Additives and Nutrient Sources added to Food), the viscosity of alginate solutions can vary from 4 to 1000 mPa.s, and the gelling capability can be affected when the solution has a pH lower than 4 [12], at which point the viscosity increases sharply [50]. According to our results, as our alginate concentration was very low (0.5 below 4), there was a slight decrease in the viscosity of the alginate solution. Additionally, the viscosity levels of carrageenan solutions should range from 5 mPa.s to 800 mPa.s, but an ideal viscosity is less than 100 mPa.s, as this would maintain the desirable properties of the carrageenan [11,16]. There are no published data regarding the relation between pH, EC, and viscosity of agar solutions. Agar viscosity is usually influenced by the temperature of the solution: at higher temperatures (>80 °C), agar-based solutions have lower viscosity, and at lower temperatures (<50 °C), the viscosity increases [16]. In our case, the agar viscosity was measured at room temperature (20–22 °C), but the concentration of agar in the solution was very low, increasing the viscosity as expected.

The polysaccharides’ structures, their biochemical characterization (mineral content), and their optimum solutions concentrations were analyzed prior to this study [30].

Polysaccharides, such as alginate, agar, and carrageenan, act as elicitors to enhance plants’ metabolisms and resistance against environmental stresses [51]. When polysaccharide-based solutions are sprayed onto the foliage, the plant’s cell wall reacts quickly to this interaction and binds with these molecules to induce local resistance. In this study, some potential polysaccharide activity was observed in relation to Agrotis sp. larvae on the turnip greens. As this was not the purpose of this study, it may be of interest as a potential subject of research in the future.

As seen in other studies [52], during this resistance process, plants can exhibit a biochemical response by enhancing their metabolic pathways and the synthesis of secondary metabolites, like phenolic compounds. These biochemical responses trigger other morphological responses related to nutrient uptake and, consequently, growth and development [53].

In this study, the turnip greens demonstrated clear differences in growth parameters among the treatments. The turnip greens treated with the polysaccharide-based solutions exhibited the best results in terms of both leaf weight and length when compared to the negative (tap water) and positive controls (“Profertil”). Plants that presented the best results were the ones treated with carrageenan from C. crispus, particularly the tetrasporophyte generation, with higher ratios of aerial part weight to length.

The observed ratios allowed us to make an association between the development of the aerial part of the plant and the roots. A lower aerial part weight vs. root weight indicated that the turnip greens had spent more energy on the growth of the root biomass than of the aerial part, and on the other hand, greater ratios indicated that the plants had spent more energy on leaf biomass (Supplementary Figure S2e–g). The lowest aerial part length: root length ratio was observed in the negative and positive controls (Supplementary Figure S2a,b), and the highest in SP and GG (Supplementary Figure S2c,d). As demonstrated in Supplementary Figure S2, the roots from plants treated with polysaccharide-based solutions (Supplementary Figure S2c–g), were more robust than the ones obtained from the control treatments (Supplementary Figure S2a,b). However, despite these ratios, major root biomass development (observed in NC and PC) did not lead to the development of, better root systems or more efficient nutrient absorption from the soil.

The mineral profiles of the turnips from each treatment were compared with the reference values of [33], which provide safety and quality standards of turnip greens for human consumption in Portugal. The differences between the treatment groups and the literature values can be also related to genetic differences and environmental factors. The mineral and trace element contents of each treatment can affect the dry matter percentage and ashes. A low ash content usually indicates that the plant is denser; therefore, this should be taken into consideration when analyzing the mineral percentage [54]. For instance, N improves the quality and quantity of dry matter in leafy vegetables, such as turnips [54]. Amino acids, building blocks of proteins, are created when N is joined to C, H, O, and S. Therefore, there is a direct correlation between the protein content and the nitrogen content. N is required for all enzymatic processes in plants and for photosynthesis [54].

The NC turnip greens exhibited the highest values of ashes, protein, minerals, and trace elements. These can be considered as genetic characteristics that were not influenced by the environment (i.e., treatments), so their phenotypes were stable. Treated plants presented very similar mineral contents compared with the published standards for human consumption in Portugal, which indicates that these solutions are not toxic and can even improve the nutritional quality of turnip greens.

In addition to mineral and trace element characterization, analyzing the cell wall and its components can help us to understand and characterize the effects that the treatments had on the turnip greens, as the plant cell wall has a significant role in plant metabolic processes. FTIR-ATR has been used for fast cell wall characterization [55]; however, due to the complexity and variability of the cell wall composition, it is not always possible to assign each FTIR-ATR band to its respective functional chemical group or compound. By comparing the obtained spectra with bibliographically supported data [55], contrary to the other samples, we found that the NC was the only one to exhibit a peak in the lignin and phenolic backbone area at 1520 cm⁻¹. Numerous studies have reported variations in the quantity of lignin and other polyphenols when plants are under a stressed environment [56].

FTIR analysis is relevant in order to understand how the polysaccharide-based solutions affected the turnip greens at a metabolic and molecular level [57]. If we compare the FTIR-ATR similarity rate between the samples treated with polysaccharide solutions and the negative or positive controls, it corroborates all the previous results (biochemical characterization) with numbers. For example, when comparing the different samples to the negative control, we obtained CC(FG) with 118,660% (most similar but with higher content, most nutritious); CC(NF) with 110,220%; CC(T) with 87,218% (less similar); GG with 94,983%; and SP with 92,348%. When comparing the different samples to the positive control, we obtained CC(FG) with 136,430%; CC(NF) with 126,570%; CC(T) with 100,170%; GG with 109,210%; and SP with 105,970%.

Apart from the mineral and cell wall characterization of the turnip greens, the detection and quantification of their pigments is a crucial step in further understanding the effects of the treatment groups in these plants during the experiment, especially in terms of photosynthetic activity. Overall, the Rf values observed in all treatments were greater than the Rf values found in the literature, except in the case of neoxanthin and β-carotene. This difference could be related to oxidation of the pigments, the type of silica plate used, the eluent, the plant species, and the quantity of the solution applied to the TLC [37]. The absence of pigment marked as “10” in the C. crispus’ TLC could be explained by the pigment’s entrainment at the end of the silica plate, preventing us from differentiating the pigments clearly. Overall, the turnip greens treated with the carrageenans extracted from C. crispus exhibited the greatest pigment content among all treatments. An increase in a pigment such as chlorophyll can indicate an increase in photosynthetic activity and, consequently, an increase in plant growth and development. This means that plant carbohydrate production boost could be related to the application of carrageenans from C. crispus in the early growth stages of the plants.

Soil/substrate is an extremely complex and important ecosystem that directly influences plants’ growth and development. The soil density (Ds) could be influenced by several physical and chemical properties, such as porosity, texture, soil organic matter, and mineral content. This information is essential for soil management and the application of the best farming techniques [58]. The Ds of all samples had very high standard deviations, which could be explained by the variation in porosity in the substrate, since its property was very difficult to control in the experiment. In addition, organic matter (OM) and nitrogen (N) content had significant influences on plant growth. Soils with high contents of OM and N usually enhance the photosynthetic processes and, consequently, the plant development [59]. However, the availability of OM and N in the substrate does not imply their absorption by the plant’s roots or their use in plant photosynthesis. As shown in our results, the turnips from the negative control, with the highest OM and N contents in the substrate, did not take any advantage from these for their growth.

The application of polysaccharide-based solutions to turnip greens was very efficient in improving the plants’ growth, biomass, and root systems and in enhancing photosynthetic activity, essential nutrient uptake, and soil fertility when compared to the PC and NC. Turnip greens treated with the carrageenan from C. crispus presented the best results in terms of crop production compared to plants treated with alginate (S. polyschides) and agar (G. gracilis). This was particularly noticeable in turnip greens treated with CC(T). The type of carrageenan extracted from this generation of C. crispus was λ-carrageenan, which is usually more sulphated (32–39% of sulfate group) than κ-carrageenan (20–30% of sulfate group) and ι-carrageenan (28–35% of sulfate group). It is a hybrid type of carrageenan extracted from the non-fructified thalli and female gametophyte of C. crispus [60,61]. The degree of sulfation can directly influence the bioactivity of the polysaccharides. Typically, alginophytes, such as S. polyschides, show the lowest sulfate group content, whereas carrageenophytes show the highest [21]; this was supported by this study. λ-carrageenan was the polysaccharide with the most bioactivity and greatest positive effect in turnip greens.

5. Conclusions

The application of polysaccharide-based solutions to turnip greens was very efficient in terms of improving their growth; increasing the plant biomass and root system; and enhancing photosynthetic activity, essential nutrient uptake, and soil fertility when compared to a commercial seaweed leaf biofertilizer (“Profertil”). Altogether, the turnip greens treated with the carrageenan from C. crispus presented the best results in terms of improving the crop productivity compared to the alginate (S. polyschides) and agar (G. gracilis). This was particularly noticeable in turnip greens treated with the carrageenan from C. crispus (tetrasporophyte), where λ-carrageenan was the polysaccharide with major bioactivity and positive effects, improving the turnip plant growth, biomass, and root systems and enriching the photosynthetic pigments, nutrients, and soil nutrient uptake. Therefore, providing polysaccharides from C. crispus seaweeds from different life cycle generations (which synthesize different types of carrageenan) and using them in plant growth and development could be extremely beneficial for the future of sustainable agriculture.

More studies should be conducted to understand the potential that seaweed polysaccharides have in agriculture.