**Morphological and Biochemical Responses of** *Glycine max* **(L.) Merr. to the Use of Seaweed Extract**

### **Sławomir Kocira 1,\*, Agnieszka Szparaga 2, Maciej Kubo ´n 3, Ewa Czerwi ´nska <sup>4</sup> and Tomasz Piskier <sup>2</sup>**


Received: 21 December 2018; Accepted: 15 February 2019; Published: 18 February 2019

**Abstract:** Currently, modern agriculture aims to improve the quantity and quality of crop yield, while minimizing the negative impact of treatments on the natural environment. One of the methods to increase plant yield and quality, especially after the occurrence of both abiotic or biotic stress factors, is the application of biostimulants. The aim of the study was to determine the effect of *Ecklonia maxima* extract on plant growth, and the yield, nutritional, and nutraceutical properties of soybean seeds. A field experiment was conducted in three growing seasons (2014–2016). Soybean seeds of Atlanta cultivar were sown in the third 10-day period of April. *Ecklonia maxima* extract was applied in the form of single or double, spraying in the concentrations of 0.7% and 1.0%. Determinations were conducted for: biometric traits, seed yield, seed number, thousand seeds weight, contents of lipids, and proteins in seeds. Further analyses included the contents of total polyphenols, flavonoids, anthocyanins, and reducing power. The number of seaweed extract applications and its concentration modified biometric traits, yield, and quality of crop, while also also altering the nutraceutical and antioxidative potential of soybean. The application of this preparation improved the growth and yield of soybean without any negative effect on the nutritive value of seeds.

**Keywords:** antioxidant activity; growth; nutrients; nutraceutical potential; soybean; yield

#### **1. Introduction**

Soya (*Glycine max* (L.) Merrill.) is one of the most important leguminous plants that are cultivated around the world because it is a precious source of both protein and fat [1,2]. Its use for production of food, oil, and fodder means that the demand for this plant is continuously growing [3]. Due to its broad use, it is called a "wonderful crop" [4]. However, this plant is sensitive to unfavourable climatic conditions [5]. Thus, to ensure its effective protection against biotic and abiotic factors, it is recommended to use it in the cultivation of biostimulants, which may improve the biochemical, morphological, and physiological processes that take place in a plant [6–8].

Biostimulants as plant-growth promoters were defined for the first time in the world literature by Kaufman [9]. In turn, Du Jardin [10] claims that "a plant biostimulant is any substance or microorganism, in the form in which it is supplied to the user, applied to plants, seeds or the root environment with the intention to stimulate natural processes of plants to benefit their nutrient use efficiency and/or their tolerance to abiotic stress, regardless of its nutrients content, or any combination of such substances and/or microorganisms intended for this use". The European Biostimulants Industry Council (EBIC) was established in order to develop legal regulations regarding the registration of biostimulants, according to the specificity of their action. However, currently, the registration of these preparations is still based on legal regulations that are set for fertilizers and plant protection products [11–13]. According to Colla et al. [14,15] and Battacharyya et al. [16], among the entire group of those preparations, extracts from seaweed and protein hydrolysates constitute the two most-important categories of substances of natural biostimulants. According to Aguilar, brown algae are the most often used in agriculture [17]. The most popular are, among others, *Ecklonia maxima* (Osbeck) Papenfuss and *Ascophyllum nodosum* (L.) Le Jolis. Brown-algae extracts include various phytohormones, such as auxins, gibberellins, cytokininis, abscisic acid, ethylene, betaine, and polyamins, and other growth promoters as well as trace elements and microelements [18]. Seaweeds also include a varied range of organic compounds, among others, aminoacids, such as asparaginic acid, glutamine acid, and alanine. While, alginic acid, laminarin, and mannitol constitute almost half of the total content of carbohydrates in such biostimulating preparations. Seaweeds also contain a wide range of vitamins that can be used by plants, such as C, B2, B12, D3, E, K, niacin, panthotenic acid, and folic acid. Although, vitamin A does not occur in algae extracts, the presence of its precursor—carotene and another possible precursor, fucoxnathin—was determined [18–20].

According to Ecoforce [21] and Van Oosten et al. [22], seaweed extracts that are used as biostimulants increase the yield and its quality in two ways: a. they stimulate hormone synthesis, influence absorption, and translocation of nutrients; b. they condition the soil, improving its ability to retain moisture and stimulate the activity of favourable microorganisms. Medjdoub estimates that the use of biostimulants, which include extracts from seaweeds, has greater meaning in agriculture. That is because plant growth and development is controlled by plant hormones, which directly or indirectly control the course of various physiological reactions and their integration with the total metabolism [23]. Many studies on seaweed extract indicate that they may increase: a. plant growth, b. activity of photosynthesis, c. resistance to fungi, bacteria and viruses, d. tolerance to ground frost, drought, and salt content, and e. yield and productivity of many cultivations [24–26], mainly by activation of protective mechanisms of plants [27]. Foliar application of biostimulants that are based on seaweeds is an agrotechnical treatment that brought many advantages in numerous cultivations, including grapevine, watermelon, strawberry, apple, tomato, spinach, onion, bean, pepper, carrot, potato, wheat, corn, barley, rice, and turf grass. The results show that plants treated with lower concentrations of extract indicated a stronger growth, higher yield, and higher mineral and nutritive elements content relative to the control [28–33]. Positive reactions also included an improved flowering and fructification ability, product quality and efficiency, and resistance to abiotic stress [31,34,35]. Studies that were performed on a wide group of crops proved that the application of sea-algae-based biostimulants stimulates the primary and secondary metabolisms in plants through the absorption and assimilation of nutrients [36–44]. Growth of productivity of crops induced by the use of such biostimulants in optimal and suboptimal conditions may be related to several direct and indirect mechanisms, including the stimulation of enzymatic activities that are related to carbon, nitrogen metabolism, Krebs cycle, and glycolysis. Such use may also induce activity similar to hormones, especially the one that is assigned to auxins and gibberellins, and improve the nutrition of treated plants by the modulation of the root system [14–16,45].

However, other results indicate that the application of such biostimulants, despite its numerous advantages, like faster germination and earlier growth [46,47], may inhibit the growth and development of many plants. This calls for greater care in the use of seaweeds extracts [48]. Therefore, the inhibition of plant growth after application of biostimulants is a potential problem in plant production. The concentration of these products is an important factor in this regard [49]. This issue may be caused directly by elements that are included in the extract [50], or it might be a consequence of modifications in the regular physiological growth of the plant [51,52]. Improvement of commercial-product formulas, and knowing the mechanisms of active substances in plants and their persistence, should mitigate such

negative effects. The exact effect of various elements (e.g., nutrients, betaines, oligomers, polymers) from seaweeds on improving plant growth, vigour, and fractioning of extracts is not fully known. Therefore, a detailed analysis of composition and the fractioning of elements on plant physiology, together with a better ability to monitor the impact of such extracts on those variables and on the expression of genes, would shed light on some of the performance mechanisms [53,54].

Among biostimulants that include seaweeds, Kelpak is particularly interesting. It is extracted from the species *Ecklonia maxima* Osbeck and then harvested along the shores of Africa. Kelpak contains phytohormones, such as auxins (11 mg dm−3) and cytokinins (0.031 mg dm−3), and also alginates (1.5 g L<sup>−</sup>1), amino acids (total 441.3 mg 100 g−1), mannitol (2261 mg L−1), neutral sugars (1.08 g L−1), and small amounts of macro- (mean composition: N 0.09%, P 90.7 mg kg−1, K 7163.3 mg kg<sup>−</sup>1, Ca 190.4 mg kg<sup>−</sup>1, Mg 337.2 mg kg−1, Na 1623.7 mg kg−1) and microelements (mean composition: Mn 17.3 mg kg−1, Fe 40.7 mg kg−1, Cu 13.5 mg kg−1, Zn 17.0 mg kg−1, B 33.0 mg kg−1) [55,56]. Moreira Sisalema [57] indicates that Kelpak, due to the unique extraction process, contains a very-high auxins-to-cytokinins ratio. The dominance of auxins stimulates the dynamic growth of plant roots, which increases the absorption of indispensable nutrients and minerals, and consequently, of plant production. Activity of this biostimulant may also increase plant resistance to drought and enable faster plant regeneration after water stress. Cals [58] shows that Kelpak should be used in leguminous plants before flowering in the dose of 2.0 L ha−1, in relation to the condition of plant nutrition. The concentration of these preparations in foliar applications is usually from 0.2% to 1% and it rarely exceeds these concentrations. Depending on the specific cultivation, cultivar, and climatic conditions, farmers are usually recommended to use these biostimulants in the form of two-week spraying in the stage of intense plant growth [28,59,60].

Because of the varied reactions of many plants to the application of biostimulants from seaweeds, and due to small number of studies on their influence in soya cultivation, a three-year log field study was carried out. Its main aim was to assess the impact of the application of *Ecklonia maxima* extract (Kelpak) on plant growth, yield size, and the quality and nutraceutical potential of genetically non-modified seeds of Atlanta cultivar. The initial hypothesis was that the introduction of agrotechnical treatment to soya cultivation in the form of plant spraying with Kelpak preparation would modify the plant growth, yield, and chemical composition of soya seeds. To test this hypothesis, the yield and structural elements of soya cropping were assessed, as well as the protein, fat, and anti-oxidant potential of seeds in relation with the applied doses and concentrations of the tested preparation. In order to know the morphological and biochemical plant reaction on seaweed extract performance, the responses of treated and untreated control plants in the same environmental conditions were compared. It was expected that the observation of plant reaction would considerably increase knowledge regarding the manner of seaweed extract performance, particularly in leguminous plants cultivation, which are sensitive to biotic and abiotic stresses. The present work is a concrete step towards broadening the understanding of the advantages of the application in agricultural practice of *Ecklonia maxima* seaweed extracts for the improvement of the size and quality of crops.

#### **2. Materials and Methods**

#### *2.1. Plant Materials and Growth Conditions*

The field experiment was carried out in 2014–2016 in Perespa (50◦66' N; 23◦63' E, Poland). It was established in a randomized block design in four replications on experimental plots with an area of 10 m2. Soybean was cultivated on the soil belonging to the Gleyic Phaeozems, which was characterized by alkaline pH (pH in 1M KCl: 7.4–7.5). The soil content in the assimilable nutrients was at the medium level, as follows: P (12.6–14.2 mg P2O5 in 100 g soil), K (15.3–17.1 mg K2O in 100 g soil), and Mg (6.2–6.8 mg Mg in 100 g soil). Each year, winter wheat was used as a forecrop. Soybean seeds (*Glycine max* (L.) Merr.) of Atlanta cultivar (Agroyoumis, Poland) were sown on the 25 of April in 2014 and 2015, and 23 of April in 2016 in rows every 30 cm at a raw spacing of

3.5 cm. The weeds were mechanically and manually removed. No pesticides were used (pests did not exceed the thresholds of harmfulness). In the growing season, the plants were sprayed with biostimulant (water solutions) that was based on the *Ecklonia maxima* extract (Kelpak). Kelpak contains phytohormones (mostly auxins 11 mg kg<sup>−</sup>1, cytokinins 0.03 mg kg−1, and auxin: cytokinin ratio 367:1), carbohydrates (16.9 g kg−1), amino acids (2.5 g kg−1), vitamin B1 (0.9 mg kg−1), B2 (0.1 mg kg−1), C (20 mg kg−1), and E (0.7 mg kg<sup>−</sup>1). The elemental profile of the biostimulant is: N 3.6 g kg−1, P 8.2 g kg<sup>−</sup>1, K 7.2 g kg−1, Ca 0.8 g kg−1, Mg 0.2 g kg−1, Fe 13.6 mg kg−1, Mn 8.4 mg kg−1, B 0.24 mg kg−1, Zn 4.2 mg kg−1, and Cu 0.2 mg kg−<sup>1</sup> [14,61]. The scheme of doses, developmental stages of plants, and terms of spraying are presented in Table 1.


**Table 1.** Plant developmental stages and dates of biostimulants application.

Plants sprayed with water served as the control. The biostimulant (or water) was sprayed with a GARLAND FUM 12B battery field sprayer (Lechler LU 120-03) at a pressure of 0.30 MPa, using 300 l liquid per hectare. The average temperature and rainfalls in the soybean growing season are shown in Table 2.


**Table 2.** Temperature (T) and rainfalls during the soybean growing season 2014–2016.

#### *2.2. Plant Growth, Yield, and Nutritional Value Determination*

After the pods have matured, when the seeds have obtained a typical color and hardness (BBCH 89), the plant height, the internode number on the main shoot, and the first pod height were recorded. In addition, after harvesting, the number of pods per plant, the number of seeds per 1 m2, the weight

of seeds, and the weight of thousand seeds were determined. Subsequently, the seeds were dried and then grinded. The flour was used for further analysis.

Protein content was determined with the Kjeldahl method, whereas the content of lipids was based on the acid hydrolysis method [62].

#### *2.3. Nutraceutical Potential*

Seed extract was prepared following the methodology that was proposed by Swieca et al. [ ´ 63]. The ground soybean seeds were extracted with a mixture of acetone, water, and hydrochloric acid (70:29:1; v/v/v). Afterwards, the samples were centrifuged for 10 min (6800× *g*) and the resulting supernatant was collected and then used for further analyses.

#### 2.3.1. Phenolics Determination

#### Determination of Total Phenolic Compounds (TPC)

The content of total phenolic compounds (TPC) was determined with the method of Singleton and Rossi using the Folin–Ciocalteau reagent [64]. Absorbance of the samples was measured with a UV-vis spectrophotometer at a wavelength of 725 nm, then TPC was computed and expressed as gallic acid equivalents (GAE) in mg per g of dry matter (DM).

#### Determination of Flavonoid Content (TFC)

The total content of flavonoids was determined acc. to the method that was presented by Lamaison and Carnet [65]. The prepared soybean extract was mixed with a methanolic solution of AlCl3 × 6H2O. After incubation, the absorbance was measured with a UV-vis spectrophotometer at the wavelength of 430 nm. The total flavonoid content was expressed as quercetin equivalents (QE) in mg per g DM.

#### Determination of Anthocyanins (TAC)

Using the method that was proposed by Fuleki and Francis using potassium chloride and sodium acetate buffer at two pH values (1.0 and 4.5), the content of anthocyanins was assayed [66]. After 15 min, absorbance of each sample was measured at wavelengths of 520 nm and 700 nm. Subsequently, anthocyanin content was calculated as cynidin-3-glucoside equivalents (Cy3-GE) in mg per g DM.

#### 2.3.2. Reducing Power

Reducing power was measured following the method that was provided by Pulido et al. [67]. The soybean extract was mixed with a phosphate buffer (200 mM, pH 6.6) and 1% solution of K3[Fe(CN6)]. Next, the samples were incubated at 50 ◦C for 20 min. The reaction was stopped with trichloroacetic acid and the samples were centrifuged (6800× *g*, 10 min). The resulting supernatant was mixed with distilled water and FeCl3. Afterwards, absorbance was measured at the wavelength of 700 nm. Reducing power was expressed as Trolox equivalents in mg per g DM.

#### *2.4. The Index of Biostimulant Effect*

The index of biostimulant effect (ABT-C) was determined as the difference between the mean result that was obtained after biostimulant application (ABT) and the control (C), which enabled the evaluation of the effect of biostimulant type on the analyzed traits. The mean value for each treatment has been obtained clustering the means of lower concentration single spraying (LSS), lower concentration double spraying (LDS), single application of the higher concentration (HSS), and higher concentration double spraying (HDS) from different years all together. The standard deviation value (SD) was determined for all reported mean values of ABT-C [5].

#### *2.5. Statistical Analysis*

The obtained results were statistically elaborated with Statistica 13 software (StatSoft, Inc.). The materials were collected over three seasons (2014–2016). Laboratory analyses were performed in triplicate. Normality of data distribution was assessed with the Shapiro–Wilk test. The significance of differences between the evaluated mean values was estimated with the Tukey test at a significance level of *p* < 0.05.

#### **3. Results**

#### *3.1. Effect of Biostimulants on Biometric Traits*

#### 3.1.1. Plant Height

The single application of the higher concentration (HSS) of Kelpak biostimulant ensured better effects in increasing soybean plant height (increased by 35% as compared to the control) (Table 3). The highest plants were obtained in the growing season 2016 after their single spraying with the higher concentration of Kelpak. In contrast, the smallest plants were produced in the 2015 season and their height differed significantly from the values noted in seasons 2014 and 2016. The biostimulant increased the height of plants, which was indicated by a value of the Kelpak effect index (ABT-C) of 28.2 cm for this trait (Table 4).



Abbreviations: C, control; LSS, lower concentration single spraying; LDS, lower concentration double spraying; HSS higher concentration single spraying; HDS, higher concentration double spraying. Means in the columns, concerning the selected traits, followed by different small letters are significantly different at *p* < 0.05.


**Table 4.** The index of biostimulant effect (ABT-C).

#### 3.1.2. Number of Internodes in the Main Shoot

Internode number decreased regardless of the Kelpak concentration and the number of its applications, although the differences were insignificant (Table 3). The highest number of internodes on the main shoot was obtained in the first and third season. The highest number of internodes on the main shoot was obtained in the first season and it differed significantly from the number that was determined in 2015. The value of the ABT-C index computed for Kelpak was negative (Table 4).

#### 3.1.3. Location Height of the First Pod

Biostimulant treatment increased the height of the first pod as compared to the control. Significant differences were observed between the double application of the lower concentration of Kelpak and the control (increased by 17%) (Table 3). The tallest heights of the first pods were observed in the 2015 season, however they did not significantly differ from the values that were reported in the two other seasons. Values of the ABT-C index demonstrate that the height of the first pod was larger with the application of Kelpak preparation (Table 4).

#### 3.1.4. Number of Pods per Plant

Double foliar application of the lower concentration of Kelpak permitted achieving the highest number of pods per plant (increased by 45% as compared to the control) (Table 3). The study demonstrated that the mean number of pods determined in particular growing seasons was at a similar level and did not significantly differ among seasons. In turn, biostimulant increased the pod number per plant because the value of Kelpak effect index was 5.9 pods/plant after spraying with this preparation (Table 4).

#### *3.2. Effect of Biostimulants on Soybean Yield*

#### 3.2.1. Number of Seeds

Double spraying soybean plants with the higher concentrations (HDS) of Kelpak had the largest effect on the increase in seed number per m2 (increased by 43% as compared to the control) (Table 5). The analysis of growing seasons demonstrated the largest value of this trait in 2016 and the smallest one in 2015 (lower by 5% than that noted in 2016). The application of seaweed extract increased this number, which was indicated by values of the ABT-C index that were calculated for this trait (Table 4).


**Table 5.** Effect of *Ecklonia maxima* extract (Kelpak) treatment on yield and nutritional properties of soybean (average from 2014–2016).

Abbreviations: C, control; LSS, lower concentration single spraying; LDS, lower concentration double spraying; HSS higher concentration single spraying; HDS, higher concentration double spraying. Means in the columns, concerning the selected traits, followed by different small letters are significantly different at *p* < 0.05.

#### 3.2.2. Seed Yield

The most positive response of plants to the use of biostimulant was observed after double spraying with the higher concentration of Kelpak preparation, as indicated by their seed yield increase by 36% when compared to the control (Table 5). The highest mean seed yield for Atlanta cv. was obtained in 2016. In contrast, the seed yield of 2015 season turned out to be the lowest among the studied seasons (lower by 4% than that noted in 2016). Foliar application of Kelpak increased the seed yield of soybean of Atlanta cv., which was indicated by positive values of the ABT-C index that were calculated for this trait (Table 4).

#### 3.2.3. Thousand Seed Weight

Foliar application of Kelpak decreased 1000 seed weight. Its lowest value was determined after double application of Kelpak in the lower concentration (decrease by 7% as compared to the control) (Table 5). The least decrease of 1000 seed weight was achieved after double plant spraying with the

higher concentration of Kelpak biostimulant. The highest mean 1000 seed weight was reported in the 2014 growing season. The values of the biostimulant effect index calculated for this trait were negative, which points to the negative impact of Kelpak preparation on 1000 seed weight (Table 4).

#### *3.3. Effect of Biostimulant on the Nutritional Properties*

#### 3.3.1. Total Protein in Soybean Seeds

Depending on concentration and number of applications, Kelpak increased or decreased the protein content in a dry matter of seeds. However, the statistical analysis demonstrated that differences in the effects of biostimulant on this trait were insignificant. Increased protein content was determined in seeds of plants single-sprayed with the lower concentrations of Kelpak (Table 5). Concerning growing seasons, the highest protein content of seeds was noted in 2015. Values of the ABT-C index that were calculated for this trait were positive for this preparations (Table 4).

#### 3.3.2. Total Fat in Soybean Seeds

Regardless of the number of sprayings and concentration of biostimulant, its use decreased the fat content in dry matter of soybean seeds, with the greatest decrease (by 14% as compared to the control) being noted after double spraying the plants with the lower concentration of Kelpak (Table 5). In contrast, the smallest decrease in fat content of the seeds as compared to the control was determined after double spraying with the higher concentrations of Kelpak. The highest fat content of soybean seeds was noted in season 2014 and the lowest in 2015. The values of the ABT-C index that were calculated for this preparation were negative (Table 4), which is indicative of its negative effect on fat content in of Atlanta cv. soybean seeds.

#### *3.4. Effect of Biostimulants on the Antioxidant Potential in Soybean Seeds*

#### 3.4.1. Total Phenolic Content

The use of Kelpak in soybean cultivation caused changes in contents of total polyphenols (TPC) in seeds (Table 6), which varied depending on both the number of applications and the concentration of this preparation. The use of the biostimulant based on *Ecklonia maxima* extract caused an increase in phenolics compounds content in soybean seeds. However, significant differences were only demonstrated in plants that were single-sprayed with 1% Kelpak (HSS). The TPC content that was determined for this combination was over twofold higher, when compared to the control combination. This nutraceutical property of soybean was influenced by meteorological conditions that occurred in a given growing season. The highest significant differences were observed in 2014 and 2016. A positive value of the difference between contents of phenolics in combinations that were treated with Kelpak biostimulant and the control samples (ABT-C) was calculated for soybean seeds (Table 4).

#### 3.4.2. Total Anthocyanins Content

The presence of anthocyanins was detected in seven out of the 15 analyzed combinations of Kelpak biostimulant use in soybean cultivation. These compounds were not detected in the control samples in any of the growing seasons studied.

The use of Kelpak affected the content of anthocyanins in soybean seeds. However, their presence was only detected in 17% of the analyzed combinations. The number of applications and concentration of the biostimulant were the factors that determined anthocyanins content. The highest value of which was noted after plants spraying with the higher concentration of Kelpak. In this case, significant differences were also observed as influenced by conditions that occurred during the plant growth stage (Table 6). The values of biostimulant effect ABT-C index calculated for this trait were positive (Table 4).


**Table 6.** Effect of *Ecklonia maxima* extract (Kelpak) treatment on the antioxidant potential in soybean seeds (average from 2014–2016).

Abbreviations: C, control; LSS, lower concentration single spraying; LDS, lower concentration double spraying; HSS higher concentration single spraying; HDS, higher concentration double spraying. Means in the columns, concerning the selected traits, followed by different small letters are significantly different at *p* < 0.05.

#### 3.4.3. Total Flavonoid Content

Flavonoid content analysis showed a significant effect of the application of biostimulant on its values. The foliar application of Kelpak resulted in the increased content of flavonoids in seeds. Significantly, the highest content of these compounds was noted after plant spraying with 1% solution of this preparation, regardless of the number of applications.

The analysis of the effect of biostimulants with different composition revealed that their foliar application resulted in an increased content of flavonoids when compared to the control samples (a positive value of the ABT-C difference) (Table 4).

#### 3.4.4. Reducing Power

The evaluation of the effect of applying biostimulants with different compositions on the antioxidant activity of soybean included the determination of the reducing power, the value of which was increased by almost all combinations of this biostimulant.

Significant differences in reducing power values were observed upon the application of Kelpak biostimulant (Table 6). A tendency for an increase of reducing potential was noted after the application of this preparation in the higher concentration and after single spraying the plants with its 0.7% solution. In the second study year, the value of reducing power was the lowest when compared to the other analyzed years (over twofold decrease of RP value). Foliar application of *Ecklonia maxima* extract

increased its reducing power values, which was indicated by positive values of the ABT-C index that were calculated for this trait (Table 4).

#### **4. Discussion**

Biostimulants induce the growth and development of plants, from seed germination throughout the entire ontogenesis. They affect the metabolic processes that occur in the plant by enhanced activity and synthesis of phytohormones, by stimulating the growth of the root system, and by improving the uptake, translocation, and retention of nutrients, which determines quantity and quality of crop yield [6,68].

Our study demonstrates a significant increase in the growth of soybean plants after the foliar application of biostimulant that is based on *Ecklonia maxima* extract. An earlier study also showed the growth stimulation of soybean treated with a biostimulant (Fylloton) containing *Ascophyllum nodosum* extract and free amino acids [8]. The marked growth responses in soybean plants are possibly due to *Ecklonia maxima* extract (Kelpak) composition, especially the PGRs (Plant Growth Regulator) that were identified (cytokinins, auxins, polyamines, gibberellins, brassinosteroids) and the mineral content in this biostimulant [55,69,70]. Additionally, the stimulatory role of Kelpak in the production of phytohormones has been demonstrated. For example, it increased the content of cytokinins in *Eucomis autumnalis* [70].

However, despite the observed favourable effects related to the application of biostimulants, including seaweeds, the precise mechanism of their activity still remains mostly unknown [26]. It should be emphasised that a full explanation of the principle or principles of their operation may cause a potential increase in the use of these preparations. According to Crouch and Van Staden [71] and Craigie [72], a wide scope of reported physiological responses of plants in cultivations where seaweed extracts were used is related to the fact that those products include numerous active compounds. Cytokinins, auxins, gibberellins, brassinosteroids, and other activating particles, like, for example, oligomers and polysaccharides are included [73]. According to Depuydt and Hardtke [74], cytokinines, together with auxins, function as regulators of various physiological processes, including those that are related to plant growth and development [75–79]. Thus, each change in the concentration of endogenic cytokinines influences the regulation of many physiological processes and as a result impacts the growth of the entire plant [80,81]. Studies by Aremu et al. [70] proved that the total content of cytokinines increases in plants after the application of the Kelpak preparation. The qualitative composition of the listed compounds is changed, which is related to their functional and physiological role, in particular, during plant morphogenesis. According to Strnad [76], isoprenic cytokinines determine the growth processes that include a continuation of the cell cycle. On the other hand, aromatic cytokinines model growth processes, such as morphogenesis and ageing. Aremu et al. [70] even assumed that the quantification of the endogenic content of cytokinins might provide information regarding possible physiological mechanisms that are related to the application of Kelpak biostimulant. However, researchers stress that, due to numerous active substances and compounds that are contained in Kelpak, the observed, favourable impact on the growth and development of plants may not only be assigned to cytokinins, but instead be the result of possible cross reactions of those compounds with other biologically active particles that are included in seaweed biostimulants. Therefore, further research concerning those fields is indispensable in order to obtain a full explanation for the Kelpak performance [70].

Still, in the literature, the prevailing hypothesis is that the majority of responses of plants that were cultivated with biostimulants' application, including seaweeds, results from the presence of compounds from the group of plant hormones, namely cytokinins [18]. The assumption stems from the fact that these compounds, isolated from seaweed extracts and individually tested in cultivations, mitigates the stress that is caused by free radicals through direct capturing and the prevention of reactive oxygen forms (ROS). This is done through the inhibition of xanthine oxidation [18,34,82–84]. Khan et al. [85] and Panda et al. [18] additionally indicate that extracts from seaweeds, such as Kelpak, support plant tolerance to stress, influencing the increase of K<sup>+</sup> capture in plants.

Additionally, in the literature, we may find hypotheses regarding modelling the growth and development of plants through the application of biostimulants from seaweeds as an effect of the presence of substances that are similar to gibberellins [86]. Research by Stephenson [19] indicates that these extracts include at least two compounds that behave like gibberellins (GA3 and GA7). However, they also show the presence of terpenoids and α-tokopherol, the performance of which may imitate gibberellins' activity on plants [18,87,88].

Recent theories indicate that the activity of seaweed extracts may be the result of their content of betains. Panda et al. shows that these compounds have a similar impact on plants as the aforementioned cytokinins [18]. It was proved that seaweed extracts include, among others, gamma aminobutyric acid betaine, 6-amino valeric acid betaine, and glyco betaine. Mancuso et al. [89] suggested that, due to the presence of those active compounds, extracts influence the mitigation of the osmotic and oxidization stress in plants, which may lead to the damage of DNA lipids, carbohydrates, and proteins, and also disturb correct cell signalling. Genard et al. [90] and Blunden et al. [91] even assigned an improvement of plant yield to the presence of betaines in extracts, since that led to the increased concentration of chlorophyll. According to Naidu et al. [92], betaines constitute a source of nitrogen when they are provided in low doses, or act as osmolites in higher concentrations. Many studies also show that betaines play a role in the correct formation of somatic germs from cotyledons tissues and mature seeds [18,93,94].

The stimulation of plant growth and development may also result from the occurrence of polyamins in biostimulants based on seaweed extracts, since these compounds may act as plant growth regulators. However, it should be emphasised that they are not classified as plant hormones. Several amino groups that usually replace hydrogen in the alkaline chain (putrescine, spermidine, and spermine) are characteristic of polyamins' structure. Research by Haman et al. [95] proves that polyamins determine the stability of various RNA and DNA conformation states. These compounds are often related to important stages of the cell division cycle. They also ensure the stability of a membrane to various cell membranes. Thus, due to the fact that polyamins affect a wide scope of physiological growth processes, their occurrence in biostimulating products that are made of seaweed may influence plant growth [18].

In our study, the use of biostimulant increased the fat and protein content in soybean seeds. The stimulating effect of biostimulants on the nutritional composition of various plants is mainly due to the number of PGR contained in the solutions [89]. In addition, the increased nutritional content in *C. triloba* is probably related to the ability of biostimulators to improve the slow release of nutrients and their uptake by plants [68,96]. A stimulating activity of seaweeds extract is also found in the presence of abscisic acid (ABA) [97,98]. However, the ABA function remains not fully characterized. Nevertheless, it is known that this acid induces protein synthesis, which are needed by plants in dealing with stress factors during water deficiencies [99,100]. Davies [101] shows that, during drought, this compound in plants caused numerous physiological reactions, including the closing of stoma, increase of a trend for accumulation of protein in seeds, gene transcription for proteinase inhibitors, as well as inhibition of sprouts growth or initiation of some states of seeds dormancy.

Unfortunately, the precise mechanisms that are activated by those biostimulants are still difficult to be identified, despite even greater knowledge on the composition of extracts from seaweeds. This is also due to the fact that these preparations constitute an abundance of many biologically active chemical compounds. They also include bioactive secondary metabolites, vitamins, and vitamin precursors [102,103]. Many authors underline the meaning of their synergetic cooperation, which stimulates that growth of plants assuming a mechanism that has not been fully known yet [24,72,104]. One of the main components of seaweed extracts are polysaccharides, including alginians, fucoidans, and laminarans [85,105]. Fucoidans have various structures due to a varied degree of methylation, sulphurization, and branching [72]. Alginians are polymers of mannuronic and

galuronic acid, with a confirmed activity of plant-growth promotion [106]. Finally, laminarins that are included in extracts are registered compounds that increase plant resistance to fungi and bacteria pathogens [107].

Improvement of a nutritive value of soya seeds observed in our research, as expressed in protein and fat content, could also be caused by the fact that seaweeds are rich in phenolic compounds (complex chloroclucinol, eckol, and dieckol polymers). Phenols belong to secondary metabolites synthesized in plants under the influence of stress. Their task is to protect cells and the components of cell nuclei [108,109]. The ability to chelate metal ions [110] is a significant role of these compounds, besides their antioxidant activity. Research by Raj et al. [60] confirms that phenolic compounds with dihydroxybenzene or trihydroxybenzene groups show strong chelating activity. Rengasamy et al. [111] also shows that eckol belonging to phenolic compounds proves to have a strong auxinosimilar activity. According to the authors, the impact of these polyphenols, included in seaweed extracts, on the endogenic content of auxins is a key element that is necessary for understanding basic mechanisms of these preparations. Korasick et al. [112] reached similar conclusions. They conclude that auxins have a strong impact on many important stages of physiological growth in the life cycle of a plant. Researchers emphasise that maintaining proper concentration of active auxin in cells is of key significance for controlling almost all aspects that are related to the plant growth. The concentration of cell auxin is affected by the speed of anabolism, catabolism, transport, and conjugation [113]. In relation to the type of concentration, polyphenols may inhibit or stimulate the development of vegetative plants. It mainly takes place due to their abilities to modulate the metabolism and concentration of active auxin forms in plants [114–117]. Gaspar et al., [118] proves that the phenolic inhibitors of oxidase IAA, such as chlorogenic acid, influence the activity of auxins. Some of the mentioned compounds even constitute alternative substrates for the oxidizing enzyme, which in turn is related to the protection of auxins before oxygen decomposition. Wilson and Van Staden [119] prove that some of the phenolic acids that protect auxins before decarboxilation increase the concentration of active forms of auxins, which are indispensable for the stimulation of growth and development of crop roots. However, according to Arem et al., attempts to explain mechanisms that are responsible for a positive response of plants to application of extracts from seaweeds should take into account the possible cross reactions between phytohormones included therein and the quantitative concentration of auxins, which may justify the observed morphological differences [120].

In the literature, one may find hypotheses that assume that the increased growth and yield of plants that are treated with seaweed extracts resulted from a positive impact of those preparations on the activity of esterase enzymes. This enzyme is considered to be a marker of plant growth processes due to its role in organogenesis. It also works as an index of somatic embryogenesis [121–124]. According to Aremu et al., a higher activity of esterase in plants that were treated with seaweed extracts indicated their stimulating impact on the increase of plant biomass production [120].

Plant metabolism may be modelled through the use of biostimulants. According to Nardi et al., this group of active preparations affects most of all carbon and nitrogen metabolism, which is associated with an enhanced activity of enzymes participating in, among others, the process of glycolysis, Krebs cycle, or nitrogen assimilation [125]. Oboh at al. and Ertani et al. demonstrated that biostimulants application yielded metabolic pathways that are linked with secondary metabolites, like e.g. phenolic compounds [126,127]. It should also be emphasized that the synthesis of secondary metabolites proceeds as an element of chemical defense [128]. Already, in 1959, these compounds were no longer treated as ballast substances [129]. Today, they are believed to play a significant role in plant protection against adverse factors [130]. The most common indicator of plants resistance to biotic factors is the content of phenolic compounds [131], which are precursors of more complex phenolic structures, like flavonoids or lignins [132].

In our study, the foliar application of Ecklonia maxima extract (Kelpak) caused a significant increase in polyphenols content. Ertani et al. and Lakhdar et al. showed that the application of biostimulants in plant cultivation enhanced the synthesis of antioxidative compounds, which are indicators of increased plant resistance to biotic and abiotic stress factors [38,133]. The physiological response of plants to the use of biostimulants results from the presence of active substances in them, such as phytohormones, amino acids, proteins, phenols, or triacontanol [38,127,134].

A positive impact of compounds that are included in seaweed extracts on the total content of phenolic compounds in soya seeds has a significant meaning in the attempt to explain the mechanisms of operation of those biostimulants. The antioxidant potential of plants is inseparable from the amount and quality of phenolic compounds [135]. In real environmental conditions, the regulation of phytochemical synthesis includes a range of advanced mechanisms that enable a precise control of production of specific particles in a suitable place and time, and also in response to outside signals [136]. Such compounds include those in seaweed extracts that may activate specific biochemical pathways that are responsible for the synthesis of secondary metabolites in plants [136–138]. According to Cheynier et al., eckol that is contained in biostimulants influences the phenylpropanoid pathway in the biochemical synthesis of phenolic acids [136]. Researchers assume that this impact is caused by the regulation of the enzymatic activity of ammonia lyase of pheyloalanin and chalconic sythase. However, only the approach that is based on the genetic analysis will enable the observation of gene regulation engaged in those pathways [16,120]. Research results that were carried out by Jannin et al. proved that cysteine protease, related to the process of synthesis of phenolic compounds, were regulated downwards, while the expression of genes that are related to photosynthesis, cell metabolism, response to stress, and nitrogen metabolism were significantly raised in the case of plants treated with seaweed extracts [139]. Roupahel et al. observed an increased concentration of phenolic compounds after the application of such biostimulants assigned to their main components, such as polysaccharides (alginians, fucoidans, and laminarins) [140]. These compounds influence endogenic hormonal homeostasis [63,64]. Additionally, processes of synthesis and accumulation of secondary metabolites may be related to the activity of enzymatic groups that are engaged in phytochemical homeostasis (the so-called direct effect) [127,140]. They also depend on the plant nutrition condition and potassium and magnesium concentration (direct effect) [41]. Roupahel et al. [140] also search for the growth of the concentration of bioactive compounds in the activation of key enzymes, such as chalkone isomerase, which is engaged in the biosynthesis of flavon precursors [141].

According to Azcona et al. [142] and Ertani et al. [127], the high effectiveness of biostimulants in plant crops is also influenced by the number of treatments at the appropriate stages of plant development. The first treatment of plant with these preparations resulted most of all in the increased number and weight of leaves, which is referred to as "short-time effect". Another dose of biostimulants, applied at the plant blooming stage, led to the long-term effect, which was manifested by changes in crop size and quality. In the case of fruits, it results in, among others, an increase in their number and weight when compared to control samples that were not treated with biostimulants [125,127]. The increased content of polyphenols in the crop may indeed result from the use of biostimulants at the appropriate growth stages of plants. Experiments that were conducted by Oboh et al. [126] and by Zhang and Hamauzu [143] confirmed that the first application of these preparations led to an increased content of phenolic compounds in leaves, and that this increase was smaller after the second application of biostimulants. According to Ertani et al. [127], changes in the total polyphenolics content resulting from different numbers of applications of biostimulants are also linked with changes in contents of individual phenolic acids.

This was since the increasing total content of phenolic acids led to an increased number of their functional groups, which are sequesters of free radicals [144]. It must be emphasized that the increased content of polyphenols in plant tissues, as evoked by the action of biostimulants, is a beneficial phenomenon, not only because of the increased plant resistance to stress factors, but also because of significant importance to consumers, since such plant products are rich sources of antioxidative compounds being valuable to the human body [145,146]. Phenolic acids, such as caffeic, gallic, and ferulic, are claimed to exhibit anticarcinogenic and antimicrobial activities [147,148].

To sum up, biostimulants that contain seaweed extracts enable many opportunities to improve plant growth. Their use in agriculture is considered to be favourable for cropping. However, the operation mechanism of such products is not completely described [149]. That is because the impact of the application of growth regulators on plants is not only a consequence of their direct ability to control metabolic pathways, since their activity may be multidirectional. Limited knowledge on the mechanisms of the preparations' activity are still mainly based on assumptions and hypotheses highlights the need of further research within this scope [5]. So far it has been proved that seaweed extracts influence the plant physiology through changes in their general profile of transcriptome, and also in metabolism [139,150]. Research by Fan et al., concerning the analysis of gene expression, expanded the understanding of the possible mechanisms that regulate the activity of these preparations [141]. Researchers indicate that, after the application of extracts from seaweed, increases in the amount of transcripts of regulatory enzymes that are related to the nitrogen metabolism (cytolase glutamine synthesis), antioxidant ability (glutathione reductase), and glycine betaine synthesis (betaine aldehyde dehydrogenesis and choline monooxigenesis) were observed [16,18,22,72].

Although biostimulants are extensively used in agricultural practice, presently the most significant research on those preparations requires a better understanding of the mechanism of their influence [22,151]. According to Van Osten et al. [22] and Povero et al. [152], only after obtaining a complete explanation of those mechanisms, can the design and production of new generation biostimulants can take place. Due to their complex composition and interactions between particular compounds, mechanisms of operation of preparations based on seaweed extracts are slowly and successively discovered, with applications of molecular biology, metabolomics, and genomics techniques. However, according to many researchers, observed favourable biological effects of extracts activity is caused by the activity of small organic particles, as well as polymers that are included in products that have an ability to regulate genes' operation responsible for ensuring and modelling plant resistance systems [16,71,139].

#### **5. Conclusions**

The number of biostimulant applications and its concentration modified the biometric traits, crop size, and yield, as well as the nutraceutical and antioxidative potential of soybean seeds. The study demonstrated that the foliar application of *Ecklonia maxima* extract improved the growth and yield of soybean without any negative effect on the nutritive value of its seeds. Our experiment showed a positive effect of double foliar application of the higher concentrations of this biostimulant on soybean seed number and yield. The application of *Ecklonia maxima* extract increased the antioxidative activity of soybean seeds, and content of total phenolic compounds, flavonoids, and anthocyanins. The results of our study indicate the need for continuing investigations and extending their scope with the aim to identify responses of different cultivable plants on the use of biostimulants that are based on various biologically-active compounds.

**Author Contributions:** S.K. and A.S. conceived and designed the research. S.K., A.S. and E.C. performed the experiments. S.K., A.S., E.C., M.K. and T.P. prepared the materials. S.K., A.S., and M.K. analyzed the data. S.K. and A.S. wrote the paper. M.K. and T.P. revised the manuscript. All authors read and approved the final manuscript.

**Funding:** This research received no external funding.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **Comparison between Chemical Fertilization and Integrated Nutrient Management: Yield, Quality, N, and P Contents in** *Dendranthema grandiflorum* **(Ramat.) Kitam. Cultivars**

#### **Beniamino Leoni, Danilo Loconsole \*, Giuseppe Cristiano and Barbara De Lucia**

Department of Agro-Environmental and Territorial Science (DISAAT), University of Bari, "Aldo Moro", Italy Via Amendola 165/A, 70120 Bari, Italy; beniamino.leoni@uniba.it (B.L.); giuseppe.cristiano@uniba.it (G.C.); barbara.delucia@uniba.it (B.D.L.)

**\*** Correspondence: danilo.loconsole@hotmail.it; Tel.: +39-080-544-3039

Received: 27 February 2019; Accepted: 17 April 2019; Published: 19 April 2019

**Abstract:** To assess the effects of a new integrated nutrient management protocol on yield and cut stem quality, root morphology, N accumulation, nitrogen utilization efficiency (NUE), and P content in tissue, a biennial (2011 and 2012) chrysanthemum cut flower cultivation was carried out. In both years, two nutrition management (CNM: conventional NM and INM: integrated NM) treatments and two *Dendranthema grandiflorum* (Ramat.) Kitamura cultivar ("White CV1" and "Yellow CV2") treatments were compared. The treatments were arranged in a split-plot design with three replicates. CNM was fertilized using a recommended dose fertilization of mineral NPK; INM treatment was fertilized using a half dose (50%) of CNM plus a combined usage of N organic fertilizer, seaweed extract (*Ascophyllum nodosum*), and microrganism consortium (*Glomus* sp. and *Bacillus* sp.). Yield at harvest (+19%), number of leaves (+33%), leaf area (+46%), number of flower heads (+27%), and total aboveground dry weight (+40%) were significantly increased by the INM application compared to the control. In terms of the root system, the increase was evident in terms of length (+174%), volume (+167%), projected area (+166%), and surface area (+165%), tips (+175%), forks (+285%), and crossings (+464%). The greatest N accumulation, in both years, was registered by INM treatment at harvest: +94% in 2011 and +55% in 2012. Differences in the NM were evident in the NUE, which was highest in CNM (on average 162) compared to INM (on average 142). In both years the P content in above-ground chrysanthemum tissues was in the order of head > leaves > stems, which was maintained in both INM and CNM treatments. A higher yield (138 stems m<sup>−</sup>2) was obtained in "CV2 Yellow" compared to "CV1 White" (120 stems m<sup>−</sup>2). Based on our findings, applying INM to chrysanthemum improves yield, cut flower quality, and plant nutrient uptake, in an agro–environmentally sustainable way. A basic economic analysis on fertilizers, cost gross production, and takings difference obtained, was carried out.

**Keywords:** N organic fertilizer; seaweed extract; mycorrhizal inoculants; phosphate-solubilizing microorganisms; biofertilizers; microorganism consortium

#### **1. Introduction**

Fertilization is essential for optimizing crop productivity [1]. Mineral fertilizers, particularly nitrogen (N) and phosphorus (P), are important for plant nutrition [2,3]. However, when used in overly large doses they are also a potential source of environmental pollution [4–6]. Nutrient overapplication has introduced major challenges in terms of soil infertility [7], N and P runoff [8,9], environmental degradation [10], and climate change [11,12].

Today there is an increasing need for a balanced fertilization strategy, minimizing the use of mineral fertilizers to enhance both crop production and quality and nutrient uptake under low input conditions [13]. Mineral fertilizers can be replaced by organic fertilizers [14], plant biostimulants [15], and beneficial microbial inoculants [16].

Possible interventions in conservation agriculture include the combined use of inorganic and organic fertilizers, as well as biostimulants and biofertilizers in order to increase a balanced nutrient supply [17]. Integrated nutrition management (INM) focusing on the optimization of the biological potential improves fertilizer input efficiency, reduces environmental risks, and increases crop productivity, through root/rhizosphere management [18].

In terms of biostimulants, seaweed extracts are used in sustainable agriculture in order to increase growth, quality, and shelf life [19–21]. Many studies have demonstrated the positive effects of seaweed extracts on a wide range of crops, including cereals [22], ornamental and flowering plants [23], vegetables [24], and field crops [25].

Biofertilizers are also an important alternative source of plant nutrients and are key components of integrated nutrient management in crop production. The use of microbial inoculants with P solubilizing activities in soils is an environmental-friendly alternative to further applications of chemical-based P fertilizers [26,27]. Various studies have examined the potential of different bacterial species to solubilize inorganic phosphate compounds. *Bacillus* spp., and in particular *B. subtilis* and *B. megaterium*, may provide the available forms of P to plants, thus considerably improving plant growth performance [28–31].

Other microbial inoculants, such as arbuscular mycorrhiza fungi (AMF), increase the P availability through the expansion of the root surface area by extraradical hyphae formation [32,33].

The various benefits of AMF include increased growth and nutrient uptake (especially N, P, and K) and crop yields [34–38]. The AMF also produce a heat-stable protein called glomalin, which is a glycoprotein that enhances soil aggregation and helps in soil carbon sequestration. Together, glomalin and mycorrhizal hyphae lead to a stable soil structure.

The combined use of N organic fertilizers, biostimulants, and biofertilizers is therefore a new approach that has not been widely investigated in ornamentals, which entails developing many efficient formulations with low mineral inputs, with positive impacts on crops and environment.

Chrysanthemum (*Dendranthema grandiflorum* (Ramat.) Kitamura) is a commercial cut flower, belonging to the *Asteraceae* family, with nearly 200 cultivars. It is one of the top ten elite cut flowers globally, due to its different shapes, dazzling colors, varying sizes, and excellent vase life. In Italy, where our research was carried out, there is a considerable demand in both domestic and export markets.

Extracts of the plants (stems and flowers) have many potential medicinal properties, including anti-HIV, antibacterial, and antimycotic [39]. N, P, and K play a vital role in the production of good quality flowers. N is essential for the creation of biomass as well as for the biosynthesis of enzymes in chrysanthemum leaves [40].

The N requirements of chrysanthemums are known to be higher during the first seven weeks of growth, and during this time, deficiencies are more difficult to correct than in later stages of development [41]. Chrysanthemums take up N at an even rate from the time of planting until the flower bud differentiation stage where after N uptake decreases [42]. In chrysanthemums, the need for P is significantly lower than that of nitrogen [43]. K requirements are high, and its presence in the plant favorably affects growth and flower color [44].

To the best of our knowledge, there are no available data on how the INM system based on mineral and organic N fertilizers, seaweed extracts, plus a consortium between AMF (arbuscular mycorrhiza fungi) and PSB (Phosphate solubilizing bacteria), affects yield and quality in chrysanthemums.

The goal of this research was to evaluate the effects of an innovative INM compared to conventional nutrient management, in chrysanthemum cut flower cultivation, on: (1) yield and cut stem quality, (2) N concentration, accumulation, and utilization efficiency and P uptake, (3) root architecture, and (4) soil fertility.

#### **2. Materials and Methods**

#### *2.1. Experimental Conditions*

Two field experiments were carried out in 2011 and 2012, from August to December, at a floricultural farm located in Sannicandro di Bari (southern Italy: 40◦59 24" N, 16◦47 01"E, 181 m a.s.l.). The local climatic conditions are characterized by hot dry summers and mild rainy autumns and winters, typical of the Mediterranean climate. During the plant growth period under natural photoperiod, the mean air temperature was 17.2 ◦C and 18.2 ◦C in 2011 and 2012, respectively; minimum air temperature was 3.7 ◦C in December 2011 and 5.4 ◦C in December 2012; maximum air temperature was 32.4 ◦C in August 2011, and 32.6 ◦C in August 2012.

Seasonal chrysanthemum cuttings (Minstreel Serie, Straathof Plants BV, The Netherlands), ideal for blooming from November to late December, were obtained from a local commercial propagator, with the following characteristics: stem length, 11.6 cm; number of leaves, 8; leaf area, 81.1 cm2; plant fresh weight, 3.1 g; and plant dry weight, 0.3 g. In both years, plants were transplanted on 6 August into an uncovered tunnel. In the first week of October in both years, the tunnel was covered by ethylene vinyl acetate (EVA) film.

The main soil characteristics (taken from 0 to 25 cm depth) are described in Table 1. Soil pH was determined with a pH meter (P9991, Hanna Instruments, Italy) in a settling suspension on a 60 g sample mixed with 150 mL of deionized water, after shaking for 60 min at room temperature (22 ◦C). The soil used for our experiment was slightly sub-alkaline (pH = 7.34, near to neutrality) and it was representative of Apulian soils in which chrysanthemum was cultivated with remarkable production results. Chrysanthemum plants generally grow with a pH ranging between 6 and 7.2 [45].

The electric conductivity (EC) was measured on water extract (1:5 *v*/*v*) with a conductivity meter (HI 4321, Hanna Instruments, Italy). Soil organic carbon (SOC) was determined by wet oxidation. Based on USDA classification, experimental soil was classified as clay loam soil. Experimental soil was moderately provided with organic matter and CEC was also classified as moderate [46].

The total Kjeldahl N (TKN) was measured using 1 g samples of both growing media and plant tissues using the Kjeldahl method after 96% H2SO4 hot digestion. Total phosphorus was determined (P) by the colorimetric molybdovanadate phosphoric acid method. Exchangeable K, Ca, and Mg were determined using 0.2 g of dry sample (105 ◦C for 24 h) after acid digestion in a microwave oven (CEM Mars Xpress, Cologno al Serio, IT). Substrate digests were filtered, diluted, and analyzed by atomic absorption spectrometry (Perkin-Elmer Aanalyst 200, Waltham, MA, USA). The analyses were carried out in triplicate.

The soil was sandy clay with a slightly alkaline pH of 7.3 (IUSS), EC of 1.77 dS m<sup>−</sup>1, and moderately high CEC (cation exchange capacity) of 23.8 Meq 100 g<sup>−</sup>1.


**Table 1.** Initial soil physico–chemical characteristics (mean ± standard error). Data are the means of three samples.

#### *2.2. Treatments and Experimental Design*

In both years, four treatments in total consisting of two nutrition management (NM) and two *Dendranthema grandiflorum* (Ramat.) Kitamura cultivar (CV) were compared as follows:


Treatments were carried out using a split-plot design with three replicates, with NM as the main plot and CV as the subplot. The surface of each experimental plot measured 2.2 m2.

CNM treatment was applied through a fertigation system using a recommended dose of mineral NPK: 17 g m−<sup>2</sup> N, 16 g m−<sup>2</sup> P2O5, and 17 g m−<sup>2</sup> K2O plus microelements, starting one week after transplanting, every week, for 12 weeks, the last one during the second week of November (flower bud differentiation).

INM treatment was applied by fertigation at a half dose (50%) of CNM plus a mixture of an N organic fertilizer, seaweed extract and microorganism consortium as shown in Table 2, starting from transplantation. Commercial products were applied at the manufacturer's recommended rates.

NPK doses added with INM fertilization were the following: 11.8 g m−<sup>2</sup> N, 8 g m−<sup>2</sup> P2O5, and 12 g m−<sup>2</sup> K2O. N organic fertilizer added to the mineral NPK dose mentioned above, was derived by hydrolyzed animal epithelium, beet molasses extract, and brown seaweed extract.

In the second year, the same treatments were repeated.

In both years, the growing density was 34 plants m<sup>−</sup>2.


**Table 2.** Combined use of N organic fertilizer, seaweed extract, and microorganism consortium applied in two experiments (2011 and 2012).

(\*) by Eurovix SpA, Entratico (BG), Italy; (\*\*) fertigation from transplant (week 1) to flower bud differentiation (week 12).

During the experiments, all field management procedures (e.g., irrigation and pest control) were the same among treatments. The irrigation system was a micro drip; each drip line was placed between two plants rows with an emitter (pressure compensating) discharge rate of 2.0 L h<sup>−</sup>1. Except for nutrition, production was carried out using the grower's standard practices. Cut flowers were harvested when 50% of flower heads had opened.

Morpho–biometric measurements were carried out at the Department of Agro–environmental and Territorial Sciences (DISAAT), University of Bari, Italy. Plants were sampled for aboveground and ground biomass and N and P content (%) at 55, 93, and 131 DAT (days after transplant) in both growing periods.

The growth and yield observations were recorded on twelve randomly selected plants from each treatment.

In both years, at harvest (second ten days of December), the soil was washed from roots, and aboveground plants were divided into stems, leaves, and flowers, which were oven dried at 70 ◦C until they reached a constant mass to measure the respective dry weights.

At flower harvest, the measurements involved: yield (secondary branches = stems m−2), stem length (cm), inflorescence (*n* and diameter, cm), leaves (*n*), and leaf area (cm2), Chlorophyll SPAD

(Single-photon avalanche diode) index (Minolta Chlorophyll Meter SPAD-502), dry and fresh weight of leaves, stems, inflorescences, roots, and whole plants. In order to perform root morphology analysis, only in the first year, roots were spread out, washed, and then scanned at 300 dpi on an HP DeskScan II scanner (HEWLETT PACKARD C6261A, Palo Alto, CA, USA). Root analysis was performed using the WinRHIZO® image analysis system (V 4.1c Régent Instruments, Quebec, Canada); measurements involved total root length, average root diameter, projected and surface area, tips, forks, and crossings.

The total Kjeldahl N (TKN) content was measured, both in the first and second years, using 1 g samples of foliar and radical tissues, using the Kjeldahl method after 96% H2SO4 hot digestion. On the other hand, the P-Olsen measurement was only used during the first year.

Nitrogen utilization efficiency (NUE) was estimated by the ratio of dry biomass to plant N accumulation at harvest.

#### *2.3. Economic Analysis*

A basic economic analysis about fertilizer costs (for CNM and INM), gross sealable production, and profit raised was developed.

#### *2.4. Statistical Analysis*

The data were analyzed by three-way ANOVA using CoStat-Statistics Software. Treatment means were separated with Student–Newman–Keuls (SNK) (*p* ≤ 0.05).

#### **3. Results**

The overall aims of this research were to evaluate the effects of an innovative INM compared to CNM, in a biennial chrysanthemum cut flower cultivation, on (i) yield and cut stem quality; (ii) root morphology; and (iii) N accumulation, NUE, and P content in plant tissue.

The main effect of NM was found to be highly significant for most of the parameters investigated. Yield at harvest, as determined by the harvestable number of cut stems per plant (Table 3),

increased significantly in INM (140 stems m<sup>−</sup>2, +19%) compared to those under CNM (118 stems m2). Genotype influenced marketable yield: CV2 registered the highest value (138 stems m−2),

surpassing that of CV1 by 15% (on average 120 stems m<sup>−</sup>2).

Concerning the Y factor, the yields were not different (133 stems m−<sup>2</sup> on average).

**Table 3.** Main effects of nutrient management, cultivar on yield, stem height, leaf number, leaf area, chlorophyll index, and number of flower heads in chrysanthemum plants over the two years of application.


Different letters within each column indicate significant differences according to SNK test (*p* ≤ 0.05). NS not significant \* *p* < 0.05 and \*\* 0.01, indicate level of significance.

Table 3 also shows the influence of the treatments on the commercial quality parameters of the cut stems at harvest. The stem height is an important parameter that is used for the classification of the stems for marketing and sales, and in fact, customers often prefer flowers with a longer stem. Stem height was not found to be significant between both NM and CV treatments; however, it showed significant differences among Y: in 2012 it was 20% lower (92 cm) than 2011 (116 cm).

Regarding the number of leaves per plant, the INM treatment led to an increase of 33% (80 leaves/stem) compared to CNM (60 leaves/stem); in 2012 the number of leaves (59) showed an average decrease of 28% (82 leaves) compared to 2011.

The INM treatment also showed a significant increase of 46% (3017 cm2) in the leaf area value compared to CNM (2064 cm2).

The chlorophyll index SPAD was not significant in any of the treatments.

The number of flower heads per stem was highest (8.4) with an increase of +27% when plants were treated with INM, compared to CNM (6.6). No differences were found between the cultivars and years.

Concerning the leaves, stems, flower heads, and aboveground dry weight, Table 4 shows the statistically significant differences in favor of INM compared to CNM. Leaf values showed a 38% increase, stem value a 37% increase, and flower heads a 55% increase, which were reflected in the increase of aboveground dry weight (+40%). No difference was found between the cultivars.

Table 4 also shows that 2011 had the highest aboveground dry weight value, which decreased to 25% during 2012.


**Table 4.** Main effects of nutrient management, cultivar on dry weight of various organs, and above-plant on chrysanthemum over the two years of application at harvest time.

Different letters within each column indicate significant differences according to SNK test (*p* ≤ 0.05). NS not significant \* *p* < 0.05 and \*\* 0.01, indicate level of significance.

In 2011 the root morphology (Table 5) was evaluated. Parameter values for the plants under INM treatment were higher than CNM as follows: root length (+174%), area projection (+166%), surface area (165%), root volume (+167%), tips (+175%), forks (+285%), and crossings (+464%).

Regarding the CV, the best performing root system was White (CV1) compared to Yellow (CV2): root length (+63%), area projection (+37%), surface area (+38%), root volume (+19%), tips (+100%), forks (+109%), and crossings (+197%).


**Table 5.** Main effects of nutrient protocol management and cultivar on total root length (TRL), area projection (AP), surface area (SA), root volume (RV), root tips (RT), root forks (RF), and root crossings (RC) at 2011 harvest period in chrysanthemum plants.

Different letters within each column indicate significant differences according to SNK test (*p* ≤ 0.05). NS not significant \* *p* < 0.05 and \*\*0.01, indicate level of significance.

Concerning the plant N accumulation (gm<sup>−</sup>2) at every DAT in both years (Table 6), the maximum value was obtained under INM, which was the result of a simultaneous increase in dry weight (Table 4). The highest N accumulation, in both years, was at harvest (131 DAT), in 2011 with an increase of 94%, and in 2012 with an increase of 55%. No significant difference was found between the CVs, except for the flower head value at 131 DAT in both years.

**Table 6.** Main effects of nutrient management and cultivar on N accumulation (g m<sup>−</sup>2) at three different days after transplant (DAT) in chrysanthemum plants over the two years of application.


Different letters within each column indicate significant differences according to the SNK test (*p* ≤ 0.05). NS not significant \* *p* < 0.05 and \*\*0.01, indicate level of significance.

Table 7 shows that in both years CNM treatment statistically influenced N accumulation (gm<sup>−</sup>2) in all plant epigeal organs and on all sample dates. In the first year, the highest N accumulation was observed compared to CNM in the leaves (+48%) at 55 DAT, stems at 93 DAT (+85%), and flower buds (+79%) at 131 DAT. Regarding INM, in the second year, the highest value was recorded in leaves (+28%) at 55 DAT, stems (+46%), and flower buds (+117%) at 131 DAT. In both years the CVs did not influence N accumulation.


**Table 7.** Main effects of nutrient management and cultivar on N accumulation (g m2) in different organs at three different DAT in chrysanthemum plants over the two years of application.

Different letters within each column indicate significant differences according to SNK test (*p* ≤ 0.05). NS not significant \* *p* < 0.05 and \*\*0.01, indicate level of significance.

Figure 1 shows that in both years the N utilization efficiency (NUE) value was highest in CNM (on average 162) compared to INM (on average 142); no significant difference was found between the CVs.

In both years, the P content (%), at harvest, in above-ground vegetative tissues (leaves, stems, and heads) of INM plants was higher than those of CNM plants (Table 8). In the first year, the increase in INM compared to CNM was 11% in the leaves, 20% in the stems, and 21% in the flower heads. In the second year, the increase in P content in the leaves under INM was similar to that recorded in the first year (12%), while it was lower for stems (+12%) and flower heads (+14%). In both years the P content in above-ground vegetative tissues were in the order of head > leaves > stems, which was maintained in both INM and CNM treatments.

**Figure 1.** Main effects of nutrient management and cultivar on N utilization efficiency (NUE) in first (A) and second (B) year in chrysanthemum plants at harvest time (\* indicates the level of significance at *p* < 0.05).


**Table 8.** Main effects of nutrient management and cultivar on phosphorus content (%) in different organs at harvest in chrysanthemum plants over the two years of application.

Different letters within each column indicate significant differences according to SNK test (*p* ≤ 0.05). NS not significant \* *p* < 0.05 and \*\*0.01, indicate level of significance.

Concerning an economic point of view, the increased yield obtained with INM (140 stems m<sup>−</sup>2) compared with CNM (118 stems m<sup>−</sup>2) led consequently to an increase of gross production of exactly €50,600.00 (takings difference). This amount was much greater than the cost increases needed for INM compared to CNM (Table 9).

**Table 9.** Basic economic analysis of fertilizers cost, gross production, and takings difference obtained.


\* Chrysanthemum price calculated at 2012–2013: 0.23 €/stem (ISMEA/2012–2013).

#### **4. Discussion**

In our study, mineral nutrient management (CNM) and integrated nutrient management (INM) were compared in chrysanthemum cultivation. The INM protocol, which combined the application of half the rate of CNM and seaweed extract, organic and biofertilizer (AMF + PSB), improved yield, cut stem quality traits, root morphology, as well as N accumulation and P content in tissues. Based on other research about INM practices [47–49], this protocol seems to be suitable in order to obtain advantages on profits and sustainability. Our aim was to verify a new mixture in order to reduce mineral fertilizer application, making chrysanthemum cultivation more sustainable, as well as highly profitable.

Compared to CNM, the INM protocol led to a significantly higher yield in terms of the number of secondary branches per m−<sup>2</sup> (Table 3). This could be attributed to a better nutrient translocation in the plant, which led to the production of a greater number of axillary buds and therefore of secondary axes, in line with Kale et al. [50] in *Salvia* and Nethra [51] in the Chinese aster. In other studies regarding biostimulant applications, yield also increased in seaweed treated plants influenced by cytokinin content, which enhances nutrient mobilization in plant organs [52].

Regarding cut stem quality traits (Table 3), our results are in agreement with Verma et al. [53], who applied an INM on chrysanthemum CV Roja. The treatment that consisted of *Azospirillum*, PSB, vermicompost, and 50% of recommended mineral NPK recorded the highest plant height, number of branches, and flowers per plant. Similar results were reported in *Crossandra* [54] and Dahlia [55]. The combination of biofertilizers with the recommended NPK dose yielded a higher flower production in *Limonium* [56] and *Calendula* [57].

In our study, a higher leaf area value was found in chrysanthemum plants under INM. According to De Lucia and Vecchietti [58], the application of seaweed extract (*A. nodosum*) in *Lilium* CV Brindisi, greatly affected these parameters (12.3 cm<sup>2</sup> of treated plants, compared with 10.3 cm<sup>2</sup> of untreated plants). This was potentially due to the direct effect of the biostimulant containing betaine. The nutrient concentration present in both the N organic fertilizer and seaweed extract biostimulant cannot on its own explain the positive response as an increase in aerial organ dry weight (Table 4). In fact cytokinins have a considerable influence on nutrient mobilization in vegetative and reproductive organs [59].

Microbial inoculants are also good supplement with half the recommended mineral dose of fertilizer. Wu et al. [60] reported that *G. mossae* plus *B. megaterium* on maize increased plant growth and NPK assimilation. As regards the effect of applying INM, the chrysanthemum root development exhibited a remarkable increase in all parameters compared to CNM (Table 5). The root growth promoting activity has been observed in snapdragon, when a biostimulant was applied [61]. Previous research has shown that the brown seaweed extract, rich in auxin, improved lateral root formation when applied to mung bean [62]. A study carried out by Mancuso et al. [63] on potted *Vitis vinifera* under seaweed extracts, showed an increase in the total volume of the root system.

Concerning the nutrient uptake, Biswas et al. [64] and Adesemoye et al. [65] showed that PGPR (plant growth-promoting rhizobacteria) also influences this parameter through a more pronounced development of the root surface area. The INM seems to encourage a better uptake of mineral nutrients by plants, which results in a higher number of branches as well as leaf area, and more flowers [66].

The N uptake by chrysanthemum plants may be enhanced by the use of biofertilizers, possibly because they stimulate better root architecture or due to the influence of growth hormones contained in seaweed extracts. These substances can increase the ability of nutrient absorption as well as enzymatic activity, in agreement with Kumari et al. [67].

The N accumulation (g m−2) value in the INM treatment could be caused by the better availability and uptake of nutrients facilitated by the application of both mineral and organic fertilizers, biostimulants, and biofertilizers (Tables 6 and 7). Mahadik et al. [68] showed that the increase in N and P uptake by chrysanthemum plants was the highest with the application of *Azotobacter* plus PSB, 50% of RDF (Recommended Dose of Fertilizers) (100:100:100 kg ha−<sup>1</sup> NPK), and 10 t ha−<sup>1</sup> of vermicompost. Regarding the P content in plant tissue (Table 8), our findings are validated by similar results found in a number of earlier studies on bacteria.

Shirmardi et al. [69] reported that PGPR solubilizes the inorganic phosphate and produces IAA, thus improving plant growth by increasing P-uptake from the soil and its transport to plant shoots. A significant increase in sunflower growth parameters, including plant P content, was found in inoculated plants after inoculation with *Bacillus* sp., possibly due to the P-solubilizing, IAA-synthesizing, and root-colonizing of these strains [70], which increase nutrient uptake.

Richardson (1994) and Rodríguez and Fraga [71] studied the influence of several soil bacteria on the supply of P to plants as a consequence of their capacity for inorganic or organic P solubilization and, therefore, for improving plant growth performance. In addition, in a 1994 study, Garbaye [72] postulated that some PSB behave like mycorrhizal helper bacteria with a synergistic interaction.

Compared to the non-treated control, the combined application of mycorrhizal fungus and rhizobacteria significantly increased growth parameters, i.e., total fresh weight, aerial dry weight, shoot length, and leaf area, in bananas. The leaf mineral content, i.e., N, P, and K, also increased significantly following the combined application of both microorganisms [73].

Finally, integrated nutrient management practices could be viable for sustainable floriculture on a commercial and profitable scale. Our data on the economics of chrysanthemum flowers are in agreement with those Verma, who showed that the cost of fertilizer can be saved with inoculation of both Azospirillum and PSB, obtaining higher flower yield compared to CNM. Angadi too, carried out a study that shed light on the combination of Azospirillum, PSB, 50% vermicompost, and 1/2 recommended NPK dose, giving the maximum net returns per euro invested.

#### **5. Conclusions**

The quality and quantity of fertilizers are the key factor affecting the growth, yield, and quality of cut flowers. Since chrysanthemum is an energy-intensive ornamental crop with a very high input of fertilizers, several experiments have been aimed at using alternative methods, reducing mineral fertilizers, and in particular the INM.

Our results shows that the INM protocol, 50% mineral RDF with N organic fertilizer plus biostimulant (seaweed extract) plus biofertilizer (microbial consortium of *Glomus* sp. and *Bacillus* sp.), is effective in enhancing yield, quality, root morphology, and nutrient uptake compared to RDF. This indicates the possibility of the sustainable, eco-friendly cultivation of chrysanthemum. In order to discern the influence of each component of INM mixture on yield and quality traits, future research is needed.

**Author Contributions:** B.L.: substantial contribution to the experimental work and data collection; D.L.: bibliographic research and support for drafting; G.C.: support with data analysis, interpretation, and critical revision of the manuscript; B.D.L.: experimental design and coordination of the work, interpretation of the data, and critical revision of the manuscript.

**Funding:** This research was funded by Chimicaverde Flor (Province of Bari).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Arbuscular Mycorrhizal Fungi Modulate the Crop Performance and Metabolic Profile of Sa**ff**ron in Soilless Cultivation**

**Matteo Caser 1,**†**, \*, Sonia Demasi 1,**†**, Íris Marisa Maxaieie Victorino 2,3,4, Dario Donno 1, Antonella Faccio 2, Erica Lumini 2, Valeria Bianciotto <sup>2</sup> and Valentina Scariot 1,2**


Received: 7 March 2019; Accepted: 7 May 2019; Published: 8 May 2019

**Abstract:** Saffron (*Crocus sativus* L.) is cultivated worldwide. Its stigmas represent the highest-priced spice and contain bioactive compounds beneficial for human health. Saffron cultivation commonly occurs in open field, and spice yield can vary greatly, from 0.15 to 1.5 g m−2, based on several agronomic and climatic factors. In this study, we evaluated saffron cultivation in soilless systems, where plants can benefit from a wealth of nutrients without competition with pathogens or stresses related to nutrient-soil interaction. In addition, as plant nutrient and water uptake can be enhanced by the symbiosis with arbuscular mycorrhizal fungi (AMF), we also tested two inocula: a single species (*Rhizophagus intraradices*) or a mixture of *R. intraradices* and *Funneliformis mosseae*. After one cultivation cycle, we evaluated the spice yield, quality (ISO category), antioxidant activity, and bioactive compound contents of saffron produced in soilless systems and the effect of the applied AMF inocula. Spice yield in soilless systems (0.55 g m−2) was on average with that produced in open field, while presented a superior content of several health-promoting compounds, such as polyphenols, anthocyanins, vitamin C, and elevated antioxidant activity. The AMF symbiosis with saffron roots was verified by light and transmission electron microscopy. Inoculated corms showed larger replacement corms (+50% ca.). Corms inoculated with *R. intraradices* performed better than those inoculated with the mix in terms of spice quality (+90% ca.) and antioxidant activity (+88% ca.). Conversely, the mixture of *R. intraradices* and *F. mosseae* increased the polyphenol content (+343% ca.). Thus, soilless systems appeared as an effective alternative cultivation strategy for the production of high quality saffron. Further benefits can be obtained by the application of targeted AMF-based biostimulants.

**Keywords:** biostimulants; *Crocus sativus*; *Funneliformis mosseae*; glasshouse; protected cultivation; *Rhizophagus intraradices*; substrate

#### **1. Introduction**

*Crocus sativus* L. (saffron) is a flowering plant belonging to the Iridaceae family [1], grown for its red scarlet stigmas that represent the world's highest-priced spice. The market price for high quality saffron can reach 15,000–20,000 € kg−<sup>1</sup> [2]. This species is widely cultivated in several countries, such as Iran, Italy, Spain, Morocco, France, Greece, China, India and Mexico [3], with an annual spice production that exceeds 220,000 kg [4].The importance and notoriety of saffron, used since ancient times as a dye, ingredient for the preparation of spirits, and condiment for food, is due to the substances contained in the spice, primarily crocins, picrocrocin and safranal [5,6]. These compounds confer the saffron's unique colour, taste, and aroma, and can also have positive biological effects. Saffron active constituents, such as carotenoids (i.e., crocins), polyphenols, and vitamins showed significant antioxidant activity [7–12]. Furthermore, saffron extracts exhibit anti-carcinogenic, anti-depressive, anti-hyperglycemic, hypoglycemic, and memory-enhancing effects [3,13]. *Crocus sativus* is a highly hand labour-intensive crop, mainly during flower harvesting and stigma separation. It is traditionally cultivated in small and flat plots, wherein mechanisation is not economically sustainable due to the harvest type and short flowering period [5,8]. Five hundred hand labour hours are needed to obtain 1 kg of dried saffron [4,5]. Saffron cultivation can be carried out on an annual or multi-year cycle [14,15]. Annual cultivation guarantees the effective control of plant diseases with a more accurate corm selection. On the contrary, in a multi-year cycle (e.g., 3–4 years in Spain, 4–5 years in Italy, and 6–8 years in India and Greece) [14], corm multiplication and the size of replacement corms in the ground can decrease drastically over the third year [15]. Environment and cultivation management affect flower induction in *C. sativus* [5,16–18]. In Mediterranean environments, flower induction occurs from early spring to mid-summer, while flower emergence occurs from early- to late-autumn. Differences in the time required for flower initiation have mostly been attributed to the corm size [19]. To produce flowers, the *C. sativus* corm diameter needs to be greater than 1 cm [20]. As the corm increases, flowering increases [16,21] and occurs in advance [22]. Commercially, a 2.5–3.5 cm diameter corm appears to be the most common size used to have full flowering already during the first cultivation cycle [23]. To increase saffron yield and quality, and to reduce production costs, flowering modulation through cultivation in soilless systems has been proposed [6,19,24]. In this cultivation system, plants are grown without the use of soil as a rooting medium and are supplied of inorganic nutrients via the irrigation water [25], and thus can benefit of a wealth of nutrients without competition with pathogens or stresses related to nutrient-soil interaction [26]. However, at present, only limited and controversial reports of saffron soilless cultivation under protected conditions are present in the literature. Molina et al. [18] reported that, in a glasshouse, temperatures may be responsible for production differences in terms of flower induction and flowering duration. Maggio et al. [19] showed that, in southern Italy, cultivation in a cold glasshouse on vermiculite and perlite-based substrates positively affected the yield and number of replacement corms. Similarly, Helal Beigi et al. [27] found that cocopeat and perlite substrates enhanced corm dry weight. While Souret and Weathers [28] and Mollafilabi et al. [24] concluded that soilless cultivation in experiments carried out in France and Iran, respectively significantly decreased the spice yield, in comparison to open field cultivation.

Plant performance in soilless systems may be improved through use of biostimulants, i.e., any natural substance or microorganism applied to plants with the aim to enhance nutrition efficiency, abiotic stress tolerance and/or crop quality traits, regardless of its nutrients content [29], with a consequent decrease of chemicals and increase of sustainability of the production system [30]. Soil microorganisms such as arbuscular mycorrhizal fungi (AMF) are collecting growing interest as biostimulants. They can form mutualistic symbiosis with about 80% of land plant species, including several crops [31]. Across the interface between the plant and the fungus, carbohydrates and mineral nutrients (i.e., N, P, Zn and B) are exchanged [32]. Thus, AMF can alleviate the limitation in plant growth caused by an inadequate nutrient supply and can improve tolerance to biotic and abiotic stress [33]. Additionally, there is evidence to indicate that AMF symbiosis may have a positive impact on crop quality [34]. Increased yield of essential oils, terpenes and polyphenols, and enhanced antioxidant activity were induced by AMF symbiosis in several medicinal and aromatic plants (MAPs) [12,35–38]. This higher concentration of bioactive molecules makes AMF-hosted plants generally more attractive for the pharmaceutical and food industries [39].

The positive effects of AMF on corm growth, spice yield, and the nutraceutical compound content of *C. sativus* have already been reported in open field trials [12,40–42]. However, so far little is known of the proper saffron AMF inocula application and effects in soilless conditions, where plants are cultivated in pots filled with sterilised substrates that are free of AM fungal propagules or highly reduced in AMF diversity [43]. In the meta-analysis performed by Berruti et al. [31], it has been observed that the fungal colonization gain in inoculated plants was significantly more frequent in the greenhouses than in the open-field conditions, even if the effectiveness of AMF inoculation on shoot biomass and yield was equally successful.

Thus, in the literature, saffron cultivation on soilless systems has been proposed for spice production, but no comparison with open field has been reported. While, the effects of AMF-based biostimulants have been investigated only in open fields. To evaluate if saffron cultivation in soilless systems and AMF application may improve crop performance, spice yield and quality, and modulate bioactive compounds content, we cultivated saffron on soilless systems, applying two AMF inocula, and we compared results with those obtained in a previous open field-based trial [12].

#### **2. Materials and Methods**

#### *2.1. Plant Material and Soilless Cultivation*

*Crocus sativus* corms with horizontal diameters of 2.5–3.5 cm, provided by the Azienda agricola "Les épices Vda" di Alessandro Putzolu (Chatillon, AO, Italy), were planted during the last 10 days of August 2017 in the experimental heated glasshouse of the Department of Agricultural Forest and Food Sciences (DISAFA) of the University of Torino (Italy, 45◦06 23.21"N Lat, 7◦57 82.83"E Long; 300 m a.s.l.). Corms were cultivated in pots (4 L, 14 cm diameter and 17 cm height; two corms per pot; density of 91 corms m<sup>−</sup>2) filled with sterile quartz sand (2 L per pot; bulk density of 1.2 kg m−3) on a layer of sterilised expanded clay (1 L per pot; bulk density of 300 kg m<sup>−</sup>3) for a total weight of about 1.5 kg. During the flowering period, the average temperatures were 22 ◦C during the day and 14 ◦C during the night.

Two inocula (MycAgro Lab, Breteniére, FR) were used in this experiment: one composed of a single fungus *Rhizophagus intraradices* (Ri) and one composed of *R. intraradices* and *Funneliformis mosseae* (Ri+Fm). Both inocula consisted of AMF spores and inorganic substrate (calcined clay, vermiculite and zeolite). Inocula treatments were compared to a control without any formulation (AMF-). Ten grams of each inoculum were inserted into each vase. The treatment was placed under each corm in order to guarantee contact between the inoculum and the roots, therefore, favouring mutualistic symbiosis. Corms were not treated for fungal pathogens and cultivation lasted one cycle (August 2017–April 2018).

A complete randomised block design was used, with a total of 48 pots in two experimental plot units (24 pots per unit) and three treatments (8 pots per treatment). Irrigation water (pH 7.4, EC 505 μS cm) was added weekly (250 mL per pot) with a drip system. The corms were fertilised by fertigation (N:K 13:46; VIGORFLOR, AL.FE. srl, MN, Italy) every 2 weeks starting from the emergence of the spate, in quantities of 1.5 g L−<sup>1</sup> of water.

#### *2.2. Determination of Flower Production, Stigma Yield and Corm Growth*

At flowering (October and November 2017), the number of flowers produced daily per corm and the yield of spice (i.e., stigmas dried at 40 ◦C for 8 h in an oven) were measured. The spice yield was calculated by weighting the mg of saffron produced per pot (area equal to 196 cm2) and comparing the values to g of spice per square meter (m2). At the end of the vegetative period (April 2018), corms were lifted, rid of topsoil, cleaned and de-tunicated, then the number, size and weight of replacement corms were determined.

#### *2.3. Preparation of the Sa*ff*ron Extract*

The saffron aqueous extracts were prepared according to Caser et al. [12]. Briefly, 50 mg of powdered saffron were suspended into 5 mL of deionised water. After stirring (1000 rpm) for 1 hour at room temperature (circa 21 ◦C) in the dark, the solution was filtered with polytetrafluoroethylene (PTFE, VWR International, Milano, Italy) filters with a 25 mm diameter and 0.45 μm pore size. The saffron extract was then diluted 1:10 with deionised water to obtain the working solution. Each sample was prepared in triplicate.

#### *2.4. Determination of Sa*ff*ron Quality by ISO 3632*

Saffron aqueous extracts were analysed with a spectrophotometer (Ultrospec 2100 Pro, Ultrospec 2100 pro, Amersham Biosciences, Uppsala, Sweden) to determine the content of picrocrocin, safranal, and crocin to have the information on the bitterness, the flavouring strength, and the colouring strength [44]. Data were related to the dry matter percentage and expressed as the absorbance of a 1% aqueous solution of dried saffron at 257, 330 and 440 nm respectively, using a 1 cm pathway quartz cell [A1% 1 cm (λ max)] and calculated according to the following formula [45]:

$$\text{A1\%} \text{1cm} \left(\lambda \text{ max}\right) = \text{D} \times 10000 \text{/m} \times \left(100 - \text{wMV}\right) \tag{1}$$

where D is the specific absorbance; m is the mass of the evaluated solution in grams; and wMV is the moisture expressed as a percentage mass fraction of the sample.

Moisture content (wMV) was determined using the following formula:

$$\text{rawMV} = (\text{m0} - \text{m1}) \times (100/\text{m0})\% \tag{2}$$

where m0 is the mass, in grams, of the saffron portion before drying; and m1 is the mass, in grams, of the dry residue after incubation, performed in an oven for 16 h at 103 ± 2 ◦C.

All analytical steps were conducted in the dark to prevent analyte degradation.

#### *2.5. Determination of Bioactive Compounds by HPLC*

Bioactive compounds were determined by means of four high performance liquid chromatography-diode array detection (HPLC–DAD) methods (Table 1; [46]) using an Agilent 1200 High-Performance Liquid Chromatograph coupled to an Agilent UV-Vis diode array detector (Agilent Technologies, Santa Clara, CA, USA). Phytochemical separation was achieved with a Kinetex C18 column (4.6 × 150 mm, 5 μm, Phenomenex, Torrance, CA, USA) using several mobile phases for compound identification and recording UV spectra at different wavelengths, based on HPLC methods, as previously tested and validated [47], with some modifications. UV spectra were recorded at 330 nm (α), 280 nm (β), 310 and 441 nm (γ), and 261 and 348 nm (δ). All single compounds were identified by a comparison and combination of their retention times and UV spectra with those of authentic standards under the same chromatographic conditions.


**Table 1.** Characteristics of the HPLC methods applied to analyse the bioactive compounds present in the studied saffron samples.

CT = conditioning time; Method α—gradient analysis: 5% B to 21% B in 17 min + 21% B in 3 min + 2 min of conditioning time—wavelength: 330 nm; Method β—gradient analysis: 3% B to 85% B in 22 min + 85% B in 1 min + 2 min of conditioning time—wavelength: 280 nm; Method γ—gradient analysis: 5% B to 95% B in 30 min + 95% B to 5% B in 5 min + 10 min of conditioning time—wavelengths: 310 nm + 441 nm; Method δ—isocratic analysis: 10 min + 5 min of conditioning time—wavelengths: 261 nm + 348 nm.

#### *2.6. Phytochemical Characterisation*

The phytochemical characterisation of each sample was performed as previously described by Caser et al. [48,49]. Briefly, the total anthocyanin content (TAC) was determined using the pH-differential method. Saffron solution was added to pH 1 and pH 4.5 buffer solutions. The absorbance of samples was determined at 515 and 700 nm after 15 min of equilibration. The results were expressed as milligrams of cyanidin 3-O-glucoside (C3G) per 100 grams of dry weight (mgC3G 100g−<sup>1</sup> DW). The total phenol content (TPC) was measured using the Folin–Ciocalteau phenolic method at 765 nm. The results were expressed as mg of gallic acid equivalents (GAE) per 100 g of dry weight (DW; mgGAE 100g−<sup>1</sup> DW). The antioxidant activity (AOA) was determined at 595 nm using the ferric reducing antioxidant power (FRAP) method and at 734 nm using the 2,2 -azinobis (3-ethylbenzothiazoline-6-sulphonic acid; ABTS) method. Results were expressed as millimoles of ferrous iron (Fe2+) equivalents per kilogram of dry weight (mmol Fe2<sup>+</sup> kg−<sup>1</sup> DW) and as μmol of Trolox equivalents per gram of dry weight (μmol TE g−<sup>1</sup> DW), respectively. All analyses were performed in three replicates and the absorbances were read using a spectrophotometer (Ultrospec 2100 Pro, Ultrospec 2100 pro, Amersham Biosciences, Uppsala, Sweden).

#### *2.7. AMF Evaluation*

On the base of saffron highly mycorrhization level (70 to 90% mycorrhizal intensity) previously reported [12], we randomly selected saffron roots in April 2018. Then, the root segments were processed for observation in light and under transmission electron microscopy. Root segments were excised under a stereomicroscope and quickly fixed in 2.5% glutaraldehyde in 0.1 M cacodilate buffer (pH 7.2) for 2 hours at room temperature and overnight at 4 ◦C. The samples were then post-fixed in 1% OsO4 in the same buffer and dehydrated in an ascending series of ethanol to 100%, incubated in two changes of absolute acetone and infiltrated in Epon-Araldite resin [50]. The resin was polimerised for 24 h at 60 ◦C. Semi-thin (1 μm) sections were then stained with 1% toluidine blue and ultra-thin (70 nm) sections were counter-stained with uranyl acetate and lead citrate [51], and used for electron microscopy analyses under a Philips CM10 transmission electron microscope.

#### *2.8. Chemicals and Reagents*

Sodium carbonate, Folin–Ciocalteu phenol reagent, sodium acetate, citric acid, hydrochloric acid, iron (III) chloride hexahydrate, 2,4,6-tripyridyl-S-triazine (TPTZ) and 1,2-phenylenediamine dihydrochloride (OPDA) were purchased from Sigma Aldrich (St. Louis, MO, USA), whereas acetic acid was purchased from Fluka Biochemika (Buchs, Switzerland). Ethylenediaminetetraacetic acid (EDTA) disodium salt was purchased from AMRESCO (Solon, OH, USA), whereas sodium fluoride was purchased from Riedel-de Haen (Seelze, Germany). Ethanol, acetone, sodium citrate and lead nitrate were purchased from Fluka Biochemika. Analytic HPLC grade solvents, methanol and formic acid were purchased from Sigma Aldrich and Fluka Biochemika, respectively; potassium dihydrogen phosphate, ammonium dihydrogen phosphate and phosphoric acid were also purchased from Sigma Aldrich. Milli-Q ultrapure water was produced by Sartorius Stedium Biotech mod. Arium (Sartorius, Goettingen, Germany). Cetyltrimethylammonium bromide (cetrimide) was purchased from Extrasynthése (Genay, France), whereas 1,2-phenylenediamine dihydrochloride (OPDA) was purchased from Sigma Aldrich. All polyphenolic and terpenic standards were purchased from Sigma Aldrich. The organic acids were purchased from Fluka Biochemika, whereas ascorbic acid and dehydroascorbic acid were purchased from Extrasynthése. All chemicals specific for electron and optical microscopy were purchased from Electron Microscopy Sciences (Newark, PA, USA), i.e., glutharaldeyde, cacodylate buffer, osmium tetroxide, epon/araldite resin, toluidine "O" and uranyl acetate.

#### *2.9. Statistical Analysis*

An arcsin transformation was performed on all percentage incidence data before statistical analysis in order to improve the homogeneity of the variance (Levene test). All the analysed data were checked for the normality of variance. For all the analysed parameters, mean differences were computed using a one-way ANOVA with a Tukey *post hoc* test (*p* ≤ 0.05). Mean comparisons between data obtained in soilless and those from the first growing season of a previous work conducted in open field [12] cultivations were performed using an independent samples t-test. All analyses were performed using SPSS 24.0 Inc. software (SPSS Inc., Chicago, IL, USA).

#### **3. Results and Discussion**

#### *3.1. Crop Performance, Quality and Secondary Metabolite Content of Sa*ff*ron in Soilless Cultivation*

Soilless cultivation in a glasshouse has been recently proposed as an alternative method to open field cultivation for saffron. Maggio et al. [19] and Gresta et al. [6] reported that, by controlling growth conditions, flowering could be modulated, extended and considerably increased, compared with open field cultivation. In the present study, under protected conditions, flowering had the same duration (ca. 22 days) compared to cultivation of the same corms planted on the same days in a northwestern Italian open field [12], but the saffron flowering moved forward about 20 days (from 5 October 2017 to 23–30 October 2017), in agreement with Gresta et al. [6]. Since, for the flower emergence, corms required to be transferred from 23–27 ◦C to 17 ◦C [18], the most likely reason for this results is related to the fact that, in a glasshouse, the lowering of seasonal temperatures takes place more slowly than in an open field. In addition to the temperature lowering, Gresta et al. [52] indicated the soil water content as another environmental component that can trigger flowering. However, as in these two studies object of comparison, the cultivation occurred in different substrates (quartz sand vs soil), it appears not possible to make speculations.

Saffron yield can vary from 0.15 to 1.5 g m−2, based on planting density, plantation age (from one to six year crop cycles), and climatic conditions during the crop season [1]. In this study, an average of 0.55 g m−<sup>2</sup> was obtained, indicating a profitable production already during the first year. This yield was similar to what obtained cultivating the same corms at a density of 39 corms m−<sup>2</sup> in a northwestern Italian open field [12] and superior to that obtained in south Italy under similar glasshouse conditions by Gresta et al. [6] (corm density equal to 40 corms m<sup>−</sup>2; 0.46 g m−2) with corms

coming from Sardinia (Italy). With similar corm density to our work, Cavusoglu and Erkel [53] and Maggio et al. [19] obtained much higher yields (0.88 g m−<sup>2</sup> and 2.34 g m<sup>−</sup>2, respectively) in glasshouses located in Turkey and south Italy. In Iranian open fields, at a corm density similar to our study, Mollafilabi et al. [24] and Koocheki and Seyyedi [54] obtained an average spice yield of 0.48 g m−2. As affirmed by Gresta et al. [52], to trigger saffron flowering, a not yet fully understood combination of temperature and soil water content is needed.

In addition to the spice yield, another economically important attribute of saffron is the number of replacement corms. The obtained values (2.63 replacement corms corm<sup>−</sup>1) are lower of those obtained by Maggio et al. [19] in soilless cultivation in a cold glasshouse in south Italy, by using peat and perlite (1:1) substrates, where corms produced from 3.0 to 4.5 replacement corms per corm. In addition to a different substrate, these authors also incubated corms in the dark for 83 days before planting. Thus, the combination of these two factors could have guarantee a superior result. Comparing to open field experiments that used corms with similar size to our study, results were in agreement with those from our trial in northwest Italy [12], and the trials performed by Turhan et al. [55] in Turkey (2.32 replacement corms corm<sup>−</sup>1), while superior to those obtained by Koocheki and Seyyedi [54] in Iranian fields (1.32 replacement corms corm<sup>−</sup>1).

Guidelines for the analyses of the main compounds that contribute to the sensory profile of saffron have been established by ISO 3632 regulations [44]. These regulations define procedures to determine these compounds by spectrophotometric analyses and have established the limits by which saffron quality is classified into three different categories (first, second and third). Specifically, the saffron produced under soilless conditions belongs to the highest quality, i.e., first category, for all the studied parameters.

The evaluation of antioxidant activity is generally considered as an important method to evaluate the nutraceutical properties of food, as indicated in other previous studies [30]. Apart from crocins, Karimi et al. [56] and Asdaq and Inamdar [57] highlighted that phenols and flavonols are responsible for the antioxidant potential of saffron. Overall, the saffron produced in soilless systems showed a very high TPC (4445.4 mgGAE 100g−<sup>1</sup> DW), more than the saffron cultivated in other sites in the Alps (range between 1340 and 2355 mgGAE 100g−<sup>1</sup> DW) [12], Lebanon (160 mgGAE 100g−<sup>1</sup> DW) [58], and India (828 mgGAE 100g−<sup>1</sup> DW) [8]. In terms of antioxidant activity, FRAP values were superior to those of Iranian and Italian samples (circa 570 and 1250 mmol Fe2<sup>+</sup> kg−1) [12,56] and ABTS values were comparable to those found in Italian and Greek saffron by Caser et al. [12] and Ordoudi et al. [59].

#### *3.2. AMF Colonisation*

In our study, the presence of AMF and their colonisation of saffron roots were confirmed by observations using light microscopy (Figure 1) and transmission electron microscopy (TEM; Figure 2) on semi-thin and thin sections, respectively. Observations on semi-thin sections, stained in blue, show that the saffron roots are mycorrhised when inoculated with both inocula (Figure 1A–C), confirming the mycorrhizal intensity described in Caser et al. [12]. At the level of the cortical root parenchyma, the typical mycorrhizal arbuscular fungal structures have been highlighted (insets Figure 1A,C). Figure 1 shows the presence of intercellular and intracellular hyphae (Figure 1C) and arbuscules (Figure 1A,B). No fungal structures were found in the roots of the control treatments (Figure 1D).

**Figure 1.** Light microscope images of semi-thin sections of *Crocus sativus* roots inoculated with *Rhizophagus intraradices* and *Funneliformis mosseae* (Ri+Fm, **A**), *R. intraradices* alone (Ri, **B** and **C**) or the control (arbuscular mycorrhizal fungi (AMF)-, **D**), stained with toluidine blue. At the level of the cortical cells, note the presence of intercellular and intracellular hyphae (i) and arbuscules (a). Magnification in insets A and C shows details of the intracellular hyphae. Cortical parenchyma (PC) cells with nucleus (N) are indicated. No fungal structure is present between and inside the root cells in AMF-roots (D). Note the central cylinder (cc) and the endodermide (en). Bars are 20 μm in A, C and D, and 10 μm in B.

Here, the host plasma membrane invaginates and proliferates around all the developing intracellular fungal structures, and cell wall material is laid down between this membrane and the fungal cell surface. The exchange of molecules between the fungal and plant cytoplasm takes place both through their plasma membranes and their cell walls; a functional compartment, known as the symbiotic interface, is thus defined. At the electron microscope level, as seen in Figure 2A,C (arrows), this new apoplastic space, based on membrane proliferation, is evident around the intracellular and arbusculated hyphae of the AMF penetrated inside the saffron root cortical cells. On the basis of TEM observations, we can conclude that the mycorrhizae, formed between saffron roots and the two species of AM fungi in the inocula used in pot experiments, are alive and functionally active.

**Figure 2.** Transmission electron microscopy images of thin sections of saffron roots colonised by *Rhizophagus intraradices* and *Funneliformis mosseae* (Ri+Fm, **A**), *R. intraradices* alone (Ri, **B** and **C**) or the control (AMF-, **D**). In details, a: fungal arbuscule; N: nucleus; M: mithocondria; P: plastids; i: fungal hyphae; PCW: plant cell wall; FCW: fungal cell wall; arrow: plant plasmamembrane; arrowhead and inset: Golgi apparatus. The bar is 1 μm in A, B, C and D.

#### *3.3. Impact of AMF on Sa*ff*ron in Soilless Cultivation*

#### 3.3.1. Crop Performance and Quality Classification

In the present study, slight differences in flowering time and production were detected between treated corms (Figure 3 and Table 2). Both applied inocula (Ri and Ri+Fm) anticipated saffron flowering time of one week, compared to untreated corms (AMF-; 23 October vs. 30 October), whereas the flowering peaks and end of flowering occurred in about the same number of days (6–9 November and 11–13 November, respectively).

No significant differences were observed between the treatments in terms of the number of flowers corm−<sup>1</sup> and the obtained mg of spice flower−<sup>1</sup> (Table 2). Very few reports about the effective role of AMF in saffron flowering and yield are available in the literature, and only under open field conditions. Aimo et al. [40] and Caser et al. [12] indicated a positive role of AMF on the saffron productive performance, with an increase in flower production (+68% and +138%, respectively, compared to the untreated corms) using AMF species belonging to the genus *Glomus*.



the second (II) quality category are: picrocrocin >55; safranal 20–50; crocins >170. ISO3632 limits for the third (III) category are:

picrocrocin>40;

 safranal 20–50; crocins>120.

**Figure 3.** Effects of AMF inoculum composed of *Rhizophagus intraradices* alone (Ri), *R. intraradices* and *Funneliformis mosseae* (Ri+Fm) or the control (AMF-) on the flowering calendar of *Crocus sativus* corms and the daily number of picked flowers m−<sup>2</sup> during soilless cultivation.

Both of the AMF inocula increased the size of replacement corms in comparison to untreated corms (Table 2), suggesting a positive effect on flower production for the following cultivation cycle, in agreement with Aimo et al. [40] and Mohebi-Anabat et al. [39]. Corm size is indeed a major factor in bulbous plants to determine the flowering capacity and production of new replacement corms [5,42].

Saffron quality greatly depends on the growing conditions [12,60]. In the present study, among the AMF inocula, *R. intraradices* alone significantly increased the content of picrocrocin (bitterness), safranal (flavouring strength) and crocins (colouring strength), in comparison to the other treatments. On the contrary, Ri+Fm significantly reduced the content of these molecules and, thus the quality of the spice, in particular by lowering the crocin content to the third category of ISO 3632. To the best of our knowledge, this is the first report indicating the effect of AMF on the quality (ISO) of saffron obtained by soilless cultivation. The positive role of Ri on the increase of the saffron quality, especially on the content of picrocrocin, was highlighted also in northwestern Italian open field [12]. Thus, the corm inoculation with Ri could further increase the already high quality saffron produced in the Italian Alps [45,61].

#### 3.3.2. Saffron Metabolic Profiling Comparing to Other Foods

In addition to the peculiar organoleptic characteristics, the stigmas of the *C. sativus* flower contain many secondary metabolites with demonstrated pharmacological effects [3,11,62–64]. The identification and quantification of bioactive compounds in saffron and the evaluation of their biological activities are important to gauge their potential efficacy in food and pharmaceutical industries [65]. The range of all chemicals can vary greatly as a result of growing conditions, such as in response to the application of biostimulants [63]. Inoculation with AMF is known to alter the production of secondary metabolites in MAPs, both in roots, shoots, and flowers, even if is not consistent among plant organs [66]. The effects of AMF inocula on the biosynthesis of secondary metabolites in saffron are presented in Table 3. This more in-depth analysis confirmed the results obtained by assessing the spice quality according to ISO3632 guidelines. The single species inoculum Ri significantly increased the content of crocins (crocin I and II), whereas the mix Ri+Fm decreased it; these findings are in agreement with those obtained by Caser et al. [12] under field conditions in a temperate mountain area (north-west Italy), where the saffron obtained by corms inoculated with Ri resulted in superior quality (i.e., quality compared to the ISO standards). Regarding antioxidant activity (AOA), inoculation with Ri resulted in superior values in both used methods (FRAP and ABTS). The AMF inoculum composed of Ri+Fm significantly increased the contents of isoquercitrin and the total phenolic (TPC) compared to Ri, while of ellagic acid in comparison to Ri and AMF-. Differences in results according to the AMF inoculum composition were also observed in other plant species cultivated on different substrates. Among the reviewed studies, it has been found that the single inoculation of *R. intraradices* tend to be more successful for bioactive compounds increase than inoculation experiments with more than one species applied at the same time. In *Echinacea purpurea* Moench. [67] cultivated in a sand and soil (1:1) substrate, *R. intraradices* alone increased more the content of polyphenols than the mixed inoculum, while in *Cynara cardunculus* L. cultivated in sandy soil [68] and *Lactuca sativa* L. cultivated in a mixture of peat, sandy loam soil and calcinated clay (1:1:1) [69] *R. intraradices* enhanced more the antioxidant activity. However, it has not been observed any effect on the accumulation of polyphenols in *Ocimum basilicum* L. cultivated in a sterilised sand and soil (3:1) substrate [70] and in *Salvia o*ffi*cinalis* L. in sand, soil, and expanded clay (1:1:1) [71,72].

**Table 3.** Bioactive compounds, anthocyanins, total polyphenol content and antioxidant activity (ferric reducing antioxidant power (FRAP) and 2,2 -azinobis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS); antioxidant activity (AOA)) of the saffron produced via glasshouse cultivation with AMF inocula composed of *Rhizophagus intraradices* alone (Ri), *R. intraradices* and *Funneliformis mosseae* (Ri+Fm) or a control (AMF-).


Mean values with the same letter are not statistically different at *p* ≤ 0.05, according to a Tukey *post hoc* test. The statistical relevance of 'Between-Subjects Effects' tests (\* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001, ns = not significant).

Karimi et al. [56] and Rahaiee et al. [63] indicated that the antioxidant capacities of saffron might be due to the presence of total phenolics and flavonoids. Based on the obtained results, the content of the bioactive compounds detected in saffron could be compared to other commonly eaten fruits with highly advantageous nutritive properties. Saffron had a higher total phenol content (TPC) and antioxidant activity (AOA) than fresh *Ribes nigrum* L. berries (circa +1000% and +493%, respectively), and fresh (circa +2000% and +1800%, respectively) and dried (circa +900% and +1650%, respectively) *Lycium* spp. fruits [65,73], analysed with the same method. Since saffron showed an antioxidant activity superior than 500 mgGAE 100g−<sup>1</sup> it could be also listed within the health beneficial fruits such as *Rubus glaucus* Berth. and *Prunus serotina* var. Capulì as suggested by Vasco et al. [74]. Its content of vitamin C was similar to what found in *Actinidia deliciosa* (A.Chev.) C.F.Liang & A.R.Ferguson and *Citrus sinensis* (L.) Osb., and even higher than in *Lycium* spp. (+150%) and *Vaccinium* spp. (+580%). Also, the coumaric acid content was superior (+85%) than in *Morus nigra* L. fruits [75] while lower than in *Lycium* spp. fruits, that showed also higher content of gallic acid, ellagic acid, catechin, and epicatechin

was generally lower in saffron (on average circa −75%, −70%, −92%, and −95%, respectively) [73,75]. Lastly, the content of anthocyanins, that are suggested to have neuroprotective properties [76], was up to 11654.5 mgC3G 100g−<sup>1</sup> DW, i.e., a value very high in comparison to fresh fruit extracts from *Morus nigra*, *Rubus idaeus* L., and *Fragaria ananassa* D. (80.0, 33.7, and 35.2 mgC3G 100g<sup>−</sup>1, respectively) [75].

#### 3.3.3. Soilless Cultivation vs. Open Field

Saffron root colonisation by AMF could be affected by the cultivation conditions related to the substrate composition, root temperature or the presence of antagonistic fungi naturally occurring in the soil [31,40,41,76]. In our recent studies, AM fungal colonisation was noted in *C. sativus* roots inoculated with Ri and Ri+Fm, both in soilless (Figures 1 and 2) and in open field conditions [12]. Figures 4 and 5 report the comparisons of the results obtained by these studies. Compared to open field, in soilless conditions not-inoculated corms (AMF-) showed similar spice yields but with higher quality while, referring to AMF treatments, Ri-inoculated corms produced less spice but with a higher quality, whereas Ri+Fm inoculated corms produced less spice, with a lower quality (i.e., reduction in crocin content).

**Figure 4.** Effects of AMF inoculum consisting of *Rhizophagus intraradices* alone (Ri), *R. intraradices* and *Funneliformis mosseae* (Ri+Fm) or a control (AMF-) on (**A**.) mg of saffron corm<sup>−</sup>1, (**B**.) picrocrocin, (**C**.) safranal, and (**D**.) crocin of *Crocus sativus* corms cultivated in soilless (black bars) and open field (grey bars, [12]) conditions. Mean comparisons of each treatment in the two cultivation types were performed using an independent samples t-test.

**Figure 5.** Effects of AMF inoculum consisting of *Rhizophagus intraradices* alone (Ri), *R. intraradices* and *Funneliformis mosseae* (Ri+Fm) or a control (AMF-) on the content of (**A**.) isoquercitrin, (**B**.) quercitrin, (**C**.) ellagic acid, (**D**.) epicatechin, (**E**.) crocin I, (**F**.) crocin II, (**G**.) ascorbic acid, (**H**.) vitamin C, (**I**.) total polyphenol content (TPC), and (L.) antioxidant activity (FRAP assay) of saffron produced in soilless (black bars) and open field (grey bars, [12]) conditions. Mean comparisons of each treatment in the two cultivation types were performed using an independent samples t-test.

With respect to the nutraceutical compounds, the comparisons are presented in Figure 5. No differences were reported between the untreated corms (AMF-), whereas the application of Ri in the soilless condition induced an increase in the contents of epicatechin, crocin I, and antioxidant activity (+80%, +435%, and +675%, respectively), while a decrease in the contents of isoquercitrin, quercitrin, ellagic acid, ascorbic acid, vitamin C, and TPC. Fewer differences were induced by Ri+Fm, which positively stimulated both the total phenolic content and antioxidant activity (+210% and +325%, respectively), but caused a decrease in quercitrin, crocin II, ascorbic acid, and vitamin C.

#### **4. Conclusions**

Soilless cultivation in a glasshouse appeared as an effective strategy for the cultivation of saffron with a first-year cultivation spice yield that is comparable with open field production sites. Moreover, the high quality saffron produced via soilless cultivation presented an elevated content of several health-promoting compounds with highly advantageous nutritive properties, such as polyphenols and elevated antioxidant activity. Further studies are needed to define better the methodologies to modulate time and duration of flowering, to improve yield, and to efficiently schedule harvest practices.

Arbuscular mycorrhizal-based products have received great interest in agriculture for their potential to improve crop productivity, nutritional quality, as well as resistance to plant pathogens and numerous environmental stresses. The literature highlights that AMF must be chosen by evaluating different aspects, such as the inoculum type, host plants, and the environmental and growing conditions.

Here, AMF successfully colonised *C. sativus* roots; their effects varied on the basis of inoculum type and cultivation conditions. Among the studied AMF inocula, *R. intraradices* appeared to give more benefits to *C. sativus* than the mix of *R. intraradices* and *F. mosseae*. Specifically, the *R. intraradices* inoculation appeared successful in open field to increase spice yields while in soilless systems to increase the spice quality.

Thus, soilless systems appeared as an effective alternative cultivation strategy for the production of high quality saffron. Further benefits can be obtained by the application of targeted AMF-based biostimulants. A cost-benefit analysis should be performed to assess the economic sustainability.

**Author Contributions:** M.C., E.L., V.B. and V.S. contributed to the experimental design. M.C., S.D., Í.M.M.V., D.D., A.F. and V.S. acquired and interpreted the data. M.C. drafted the manuscript. V.B. and V.S. conceived the study, coordinated the work and critically revised the manuscript.

**Funding:** This research was funded by the project titled 'SaffronALP—Lo zafferano di montagna: tecniche sostenibili per una produzione di qualità' - Fondazione Cassa di Risparmio di Torino (RF=2017.1966) and by the program Interreg V-A Francia Italia Alcotra "Attività innovative per lo sviluppo della filiera transfrontaliera del fiore edule–Antea' n. 1139.

**Acknowledgments:** The authors acknowledge Dario Sacco for statistical assistance and Alessandro Putzolu for technical assistance.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **Exogenous Application of Amino Acids Improves the Growth and Yield of Lettuce by Enhancing Photosynthetic Assimilation and Nutrient Availability**

**Shumaila Khan 1,**†**, Hongjun Yu 1,**†**, Qiang Li 1,**†**, Yinan Gao 2, Basheer Noman Sallam 1, Heng Wang 1, Peng Liu <sup>1</sup> and Weijie Jiang 1,\***


Received: 19 April 2019; Accepted: 16 May 2019; Published: 26 May 2019

**Abstract:** As natural plant growth stimulators, amino acids are widely used to improve the yield and quality of crops. Several studies have illustrated the effects of different amino acids on lettuce plant parts. However, the effects of applying single amino acids on root growth remain elusive. The objective of this study was to evaluate the effect of root application of L-methionine on the growth of lettuce. In this study, two successive experiments on butterhead lettuce were conducted under hydroponic conditions. Three amino acids, L-methionine (20 mg/L), L-glycine (210 mg/L), and L-tryptophan (220 mg/L), were applied separately. L-methionine significantly increased the growth performance by 23.60%, whereas growth using L-tryptophan and L-glycine decreased by 98.78% and 27.45%, respectively. Considering the results of the first experiment, a second experiment was established with different concentrations of L-methionine (2200 mg/L, 220 mg/L, 22 mg/L, 2.2 mg/L, 0.2 mg/L, and 0.02 mg/L). The plants were allowed to grow for four weeks. Leaf width, plant area, leaf area, chlorophyll contents, etc., were evaluated. The results show that plant growth significantly improved by applying L-methionine at the lowest concentrations of 0.2 mg/L and 0.02 mg/L, which can, therefore, improve hydroponic production of lettuce and, accordingly, human nutrition.

**Keywords:** L-methionine; L-tryptophan; L-glycine; lettuce; nitrogen

#### **1. Introduction**

Lettuce (*Lactuca sativa* L.) is one of the main vegetable crops widely cultivated in China and consumed as a salad throughout the world. Due to its high nutritional value provided by mineral elements, vitamins, and folate, which play significant roles in human nutrition and diet, lettuce has become the focal point of several studies [1–4].

Nitrogen is an essential element for lettuce plants, an integral component of protein, phospholipid, and chloroplast [5,6]. Nitrogen uptake, assimilation, and utilization play essential roles in plant growth and development [7,8]. The plants mainly take up nitrogen in the form of nitrate (NO3 −) and ammonium (NH4 <sup>+</sup>) or N2 from the atmosphere through nitrogen-fixing bacteria [9–11]. The application of a large amount of chemical fertilizer to ensure high crop yield causes serious issues for agricultural products [12] and the environment [13,14]. Hence, there is a need to look for sustainable horticultural

practices to counteract chemical-based agribusiness. In this respect, the application of amino acids, as a type of growth-promoting substance, supplies plant nutrients but also improves plant quality, which ultimately boosts the yield and commercial output of crops [15]. Therefore, it has become popular in sustainable agriculture [16–20]

Amino acids as biostimulants (substances that promote plant growth, improve nutrient availability, and enhance plant quality) [20,21] are not only getting popular for mitigating injuries caused by abiotic stresses [22] but also serve as hormone precursors [20,23–25]; signaling factors of different physiological progressions, such as glutamate receptors (GRLs) [24–26]; and regulators of nitrogen uptake [27], root development [25,28–30], and antioxidant metabolism [25,30–32]. Better root development supported by the addition of amino acids can boost nitrogen fixation, which induces an enhanced root surface for nutrient uptake [29,33].

Direct application of amino acids and their products could modulate N uptake and assimilation; this phenomenon is mediated by enzymes engaged with N assimilation [15,25,30]. In addition, it could be a follow-up to the flagging pathway that controls N securing amino acids in roots, which are mostly accessible as supplements [30]. Additionally, application of amino acids was also found to increase K<sup>+</sup> in plants both in the presence of salt stress and without salt application [25,30,32]

A recent study [31] showed that seed treatment or foliar application of amino acids had different effects on soybean crops. An amino acid applied individually acts as a signaling component, i.e., increases antioxidant enzyme activity and causes efficient nutrient uptake [25,34,35]. Different investigations have demonstrated a positive impact of foliar application of amino acid mixtures on plants, for example, increased production in *Solanum lycopersicum* L. [36] and accumulation of dry matter, chlorophyll [37], starch, and polysaccharides in *Vicia faba* L. [38].

Previous studies have indicated an association of L-methionine with the biosynthesis of growth regulating substances such as cytokinins, auxins, and brassinosteroids in plants [39,40]. L-methionine functions as a precursor of a significant number of essential biomolecules such as vitamins, polyamine, cofactors [41], and antioxidants such as glutathione, which are considered to be significant determinants of cellular redox homeostasis and many defense compounds [19]. All of these biomolecules contain sulfur moieties that act as functional groups and are derived from L-methionine. In plants, L-methionine biosynthesis plays a central role in fixing inorganic sulfur from the environment, providing the only metabolic sulfide donor for the generation of glutathione, phytochelatins, iron–sulfur clusters, vitamin cofactors, and multiple secondary metabolites [42,43].

However, there is little data on the impact of isolated amino acids, particularly in root application. Additionally, the majority of investigations have been carried out utilizing a mixture of amino acids and other methods of application, such as foliar application and seed treatment. Hence, this study is based on the hypothesis that the root application of individual amino acids can improve the uptake of nitrogen and other growth-related factors, which can lead to increased productivity of lettuce plants. Therefore, the objective of the present work was to evaluate the effect of the separate application of L-tryptophan, L-glycine, and L-methionine in nutrient solution on the growth, yield, and physiology of lettuce plants.

#### **2. Materials and Methods**

The research study was carried out at the Vegetables and Flowers Institute, Chinese Academy of Agricultural Sciences, Beijing, China, in 2017–2018 to determine the regulation of lettuce plant growth response under different amino acids and concentrations.

#### *2.1. Plant Material and Growth Conditions*

Sowing of butterhead lettuce seeds was done under controlled conditions using a mixture of peat moss with an average of 2–3 seeds per hole. All cultural practices were maintained in order to have a good plant stand. Average minimum and maximum monthly temperatures were set to 24 ◦C and 34 ◦C. Plants were provided with natural sunlight with a light intensity of approximately 900–1000 μmoles m−2/s. At pre-emergence stages, the nutrient solution was applied once a week. Plants with at least 2 fully expanded leaves 30 days after sowing were transferred to a closed-loop hydroponic system (Figure 1).

**Figure 1.** The hydroponic system, mesh basket, and other materials used to grow lettuce plants.

Briefly, the hydroponic system consisted of 3 growing nutrient trays (used as 3 replicates) with 10 holes each, with a distance of 21.2 × 20.5 cm between the holes. The height of the hydroponic growing stand was 160 cm from the ground, while the length and width of the growing tray were 102 cm and 38 cm, respectively. The capacity of the water reservoir was 80 L with 6 cm depth and recyclable. Plastic mesh used to cover the plants was 6 cm long and 2 cm wide.

A pH of 6.0–6.3 and electrical conductivity (EC) of 1.5–2.0 mS cm−<sup>1</sup> of nutrient solution were maintained regularly for optimal plant growth.

Plantlets were grown with 75% strength Hoagland nutrient solution containing the following nutrients (in mg/L), as previously described [44]: Ca(NO3)2 = 1122; KNO3 = 910; KH2PO4 = 272; NH4NO3 = 40; MgSO4 = 247; EDTA (Ethylenediamine tetraacetic acid Ferric Sodium Salt) = 16.80; ZnSO4 = 1.20; Na2B4O7 = 0.28; Na2MoO4 = 0.20; CuSO4 = 0.10; and MnSO4 = 0.86.

#### *2.2. Application of Three Amino Acids on Lettuce*

The experiment was conducted from December 2017 to February 2018 with 3 replications. The concentration of 3 amino acids, L-methionine, L-tryptophan, and L-glycine, was kept at 20 mg/L, 210 mg/L, and 220 mg/L, respectively. This gave 4 treatment combinations (3 amino acids and 1 control treatment). The amino acid treatment was started 8 days after transplanting into the nutrient solution to prevent plants from undergoing nutrient shock. Data were recorded every week and the crop was harvested after 30 days of treatment.

#### *2.3. Application of L-Methionine Concentrations on Lettuce*

The second experiment was conducted from January to March 2018 with a single amino acid, L-methionine (selected from experiment 1), in 6 concentrations, as 3 treatments and 3 replications. Plants were transplanted to the nutrient solution and treated with the amino acid after 8 days. The concentration of L-methionine applied was 2200 mg/L, 220 mg/L, and 22 mg/L for the treatment and 2.2 mg/L, 0.2 mg/L, and 0.02 mg/L for the control. All other experimental conditions were the same as in the first experiment. The tanks of nutrient solution were refreshed weekly.

#### *2.4. Data Collection and Analysis*

Data were recorded for the following morphological and physiological parameters: root length, leaf length, leaf width, leaf area, plant area, chlorophyll content, and fresh and dry mass of root and shoot.

#### 2.4.1. Vegetative Growth Parameters

The number of leaves, plant height, plant diameter, and leaf area were measured every 7 days following standard procedures as proposed in [45–47].

$$AF\left(cm^2\right) = 0.7 \times Length\left(cm\right) \times Width\left(cm\right) - 2.4$$

Leaf length and width were measured by using a measuring tape/scale.

Root length was measured by separating roots from plants and placing them on paper and blotting them, then using a measuring tape (recorded in cm). Fresh weight per plant was square root transformed to normalize the error distribution before the analysis, as described [48], using an electronic balance (S = 0.1 g) (Acculab V-1200). The harvested plants were rinsed with distilled water, then the roots were blotted on filter paper and dried completely in an oven at 60–65 ◦C to determine dry weight [49]. The following formulas were used to calculate the index of growth traits [45].

Relative growth rate (RGR) was calculated by the following formula:

$$\text{RGR} = \left(\text{lnW}\_2 - \ln \text{W}\_1\right) \left(\text{t}\_2 - \text{t}\_1\right)$$

where W2 and W1 denote the plant's dry mass (g) at time t2 and t1, respectively.

The net assimilation rate was calculated by the following formula:

$$\text{VAR} = \text{dW}/(\text{A} \times \text{dt})$$

where A is the area of assimilation organs (cm2), dW is the dry mass increment (g), and dt is the time of cultivation (days). Root mass ratio (RMR; root mass per unit total plant mass) was calculated as described in [49].

#### 2.4.2. Physiological Measurements

Total chlorophyll content was estimated by using a portable The Soil Plant Analysis Development (SPAD) chlorophyll apparatus (SPAD-502 Plus, Konica Minolta, Tokyo, Japan). The leaf net photosynthesis rate for 3 independent lettuce seedlings per experimental replicate was determined using a portable LI-6400 photosynthesis system (Li-Cor 6400-18, Lincoln, NE, USA) [40,50]. The set values used were as follows: photosynthetic photon flux density, 500 <sup>μ</sup>mol·m−2·s<sup>−</sup>1; air flow rate inside the sample chamber, 400 <sup>μ</sup>mol·s<sup>−</sup>1.

The nutrient contents were measured in dried leaves ground by an electric mortar (multipurpose high-speed disintegrator, Dingia), sieved, weighed out to 0.2 g, and digested by concentrated nitric acid (HNO3, 5–6 mL) carefully under a laminar flow hood cabinet. All nutrients were analyzed using an optical emission spectrometer (Optima 5300 DV Spectrometer, Shelton, CT, USA).

The total N was determined by the Kjeldahl method [51].

#### *2.5. Statistical Analysis*

The recorded data were subjected to analysis of variance (ANOVA) and fixed-factor models [52], and Duncan's multiple range test was used to assess the significance of treatment differences by means of IBM SPSS Statistics for Windows (version 20.0, IBM Corp., Armonk, NY, USA).

#### **3. Results**

#### *3.1. Application of Three Amino Acids on Lettuce*

The effects of different amino acids were studied for vegetative growth. The plants grown in the modified nutrient solution (Hoagland and amino acids) showed varied responses for metric traits.

The vegetative indicators responded positively and significantly to all applied L-methionine concentrations (Figure 2A–F). Leaf length, width, and the number increased in response to L-methionine application, and decreased with L-glycine and L-tryptophan. The leaf length of L-methionine treated plants increased by 11.41% compared to control plants, while it decreased by 13.76% and 61.92% in L-glycine and L-tryptophan treated plants, respectively (Figure 2A). A significant increase in leaf width (17.46%) was also found with L-methionine, but there was an 18.25% and 63.49% decrease in response to L-glycine and L-tryptophan, respectively (Figure 2B). Similarly, leaf area and leaf numbers also increased under L-methionine treatment (31.41% and 50.4%, respectively), while leaf area decreased under L-glycine and L-tryptophan (29.67% and 86.25%, respectively) and leaf numbers decreased under L-tryptophan (50.36%) compared to control (Figure 2C,D).

Furthermore, plant height and area also had an encouraging response to L-methionine application (Figure 2E,F). The results revealed that there was an abrupt change in plant height and area, which were reduced by 82.91% and 90.78%, respectively, upon L-tryptophan application.

**Figure 2.** (**A**) Leaf length, (**B**) leaf width, (**C**) leaf area, (**D**) plant height, (**E**) leaf number, and (**F**) plant area with L-methionine (20 mg/L), L-glycine (220 mg/L), and L-tryptophan (210 mg/L). Means followed by the same lowercase letters do not differ significantly from each other in the comparison between amino acid treatments each week using Duncan's multiple range test (*p* < 0.05).

Likewise, root length, shoot-to-root ratio, relative water content, net assimilation rate, and fresh and dry biomass were positively affected by L-methionine treatment and negatively by L-tryptophan and L-glycine (Figure 3). The results revealed that lettuce plants showed a significant increase in root length (Figure 3A), relative water content (Figure 3D), and net assimilation rate (Figure 3C) in response to L-methionine application, and a decrease with the other two amino acids. Interestingly, the shoot-to-root ratio was found to be higher in response to L-tryptophan, which suggests that amino acids other than L-methionine also have an important role in plant growth. Moreover, a relative increase in fresh and dry biomass was observed with L-methionine application, by 20.88% and 15.71%, respectively, and a decrease with L-tryptophan and L-glycine treated plants (Figure 3D). Taken together, these results indicate that biostimulants, specifically the amino acid L-methionine, play a critical role in the growth and development of lettuce plants.

**Figure 3.** Effects of amino acids on (**A**) root length, (**B**) net assimilation rate (NAR), (**C**) plant fresh weight (FW), (**D**) relative water content (RWC), (**E**) shoot-to-root ratio (S:R), and (**F**) plant dry weight (DW). Means followed by the same lowercase letters do not differ significantly from each other according to Duncan's multiple range test (*p* < 0.05).

#### 3.1.1. Photosynthetic Measurements

Photosynthetic measurements of lettuce leaves (Table 1) included amino acid application effect, rate of net photosynthesis, stomatal conductance, and transpiration rate. They were significantly affected by applied amino acids compared to control, except for total chlorophyll content. Transpiration rate and intracellular CO2 were relatively higher among all amino acid treated plants. The high levels of photosynthesis and chlorophyll content suggest that amino acids are important regulators of photosynthesis in lettuce, ultimately leading to higher yield and biomass.


**Table 1.** Effects of amino acids on physiological indicators of lettuce plants.

L-Meth (20 mg/L), L-Try (220 mg/L), and L-Gly (210 mg/L). n.s., not significant. Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test (*p* < 0.05).

In addition, relative growth parameters including leaf area index, leaf dry matter content, root mass ratio, specific leaf area, and leaf area ratio were higher in response to L-tryptophan treatment (Table 2). The relative growth rate was found to be higher with L-glycine application. L-methionine and L-glycine treated plants had a nonsignificant association with all other traits compared to control plants. These results show that along with L-methionine, L-tryptophan is also an important player, especially for relative growth enhancement in lettuce.


**Table 2.** Effects of amino acids on growth indices of lettuce plants.

L-Methionine (20 mg/L), L-Tryptophan (220 mg/L), and L-Glycine (210 mg/L). LAI, leaf area index; LDMC, leaf dry matter content; RMR, root mass ratio; SLA, specific leaf area; LAR, leaf area ratio; RGR, relative growth rate; n.s. not significant. Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test (*p* < 0.05).

However, vitamin C content was not significantly affected by the applied concentrations, although there was a tendency for it to be higher in L-methionine and L-glycine compared to control (Table 3). In contrast, dry matter percentages were higher in L-tryptophan treated plants.



L-Methionine (20 mg/L), L-Tryptophan (220 mg/L), and L-Glycine (210 mg/L). n.s., not significant. Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test (*p* < 0.05).

#### 3.1.2. Nutrient Contents

Nutrient content analysis (Table 4) revealed a dynamic change in response to all amino acid concentrations. The content of essential elements such as nitrogen, phosphorus, and potassium varied among treatments.


**Table 4.** Effects of amino acids on essential elements of lettuce.

L-methionine (20 mg/L), L-tryptophan (220 mg/L), and L-glycine (210 mg/L); Macronutrients: N, nitrogen; P, phosphorus; K, potassium; S, sulfur; Ca, calcium; Mg, magnesium. Micronutrients: Fe, iron; Cu, copper; Mo, molybdenum; Na, sodium; Zn, zinc; Al, aluminum. Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test (*p* < 0.05).

#### *3.2. Application of Di*ff*erent L-Methionine Concentrations on Lettuce*

In the second trial, L-methionine at a higher concentration had a reduced effect on plant growth and physiology. The results show that lower levels of L-methionine significantly contributed to enhancing the number of leaves, plant height, and leaf length and width (Figure 4). Remarkably, 0.22 mg/L concentration of L-methionine resulted in a gradual increase in vegetative growth compared to control plants during all weeks. In contrast, higher levels were negatively associated with the corresponding measurements. Lettuce plants had short stature and fewer leaves in response to 2200 mg/L and 220 mg/L of L-methionine (Figure 4B,D). Concomitantly, more than 80% and 50% decreases in these two traits were found with increasing amino acid levels.

**Figure 4.** Effects of L-methionine concentrations on (**A**) leaf length, (**B**) plant height, (**C**) leaf width, and (**D**) the number of leaves per plant. Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test at *(p* < 0.05).

Similarly, relatively decreased leaf length (42.24% and 23.3%, respectively) and width (54.5% and 25.23%, respectively) were observed with increased treatment levels (Figure 4A,C). A strong increasing trend was also found for both traits in response to 0.2 mg/L of L-methionine. Overall, these results indicate that higher levels of L-methionine have an inhibiting effect on plant growth.

Leaf and plant area, root length, and fresh and dry weight of lettuce plants were improved by lower L-methionine concentrations (especially 0.2 mg/L) in advanced growth stages (Figure 5). However, mixed growth patterns were also present with different amino acid levels at each plant stage. A maximum reduction in leaf and plant area (75.5% and 74.653%, respectively) was found in 2200 mg/L treated plants, and a minimum (15.6% and 4.03%, respectively) in 0.2 mg/L plants (Figure 5A,B). Moreover, root length was found to be reduced at all levels except 0.2 mg/L, which caused a relative increase by 14.8% (Figure 5C). At the same time, the fresh and dry weight of roots were also lower with more concentrated treatment (Figure 5D), which suggests that amino acids are essential elements required as micronutrients for plant growth. An increased concentration leads to restricted plant biomass. Moreover, the decreased fresh and dry weight of lettuce plants indicates that nutrient stress and reduced photosynthetic activity were responsible for the lower accumulation of leaf organic material and growth rate.

**Figure 5.** Effects of L-methionine concentrations on (**A**) leaf area, (**B**) plant area, (**C**) root length, (**D**) root fresh weight, (**E**) and root dry weight. Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test at (*p* < 0.05).

#### 3.2.1. Relative Growth Measurements

Relative growth parameters measured in the second trial revealed that most of the traits were substantially increased by decreased L-methionine concentrations (Table 5). Surprisingly, root mass and leaf area ratios increased (*p* < 0.05) under higher concentration, which shows that plants can respond to a stress environment by maintaining their growth patterns. However, all other measured parameters had a decreasing tendency under a nutrient stress environment.


**Table 5.** Effects of concentrations of L-methionine on growth indices of lettuce plants.

LAI, leaf area index; RMR, root mass ratio; SLA, specific leaf area; LAR, leaf area ratio; RGR, relative growth rate; RWC, relative water content. Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test *(p* < 0.05).

#### 3.2.2. Photosynthetic Measurements

Enhanced photosynthetic activity, transpiration, and total chlorophyll content (Table 6) at lower L-methionine concentrations suggest that plants require this nutrient only in small amounts. More moderate transpiration activity occurred with all applied treatments, and reduced accumulation of chlorophyll in leaves causes restricted photosynthetic activity. There was no significant difference observed for stomatal conductance but it was relatively higher with 22 mg/L of L-methionine.

**Table 6.** Effects of L-methionine levels on net physiological and growth indicators of lettuce leaves.


Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test (*p* < 0.05).

#### 3.2.3. Nutrient Contents

Data in Table 7 describe the effects of L-methionine on macro- and microelements. The increasing trend of essential element accumulation including N, P, and K at reduced L-methionine levels indicates that these elements affect plant metabolism and help to adapt to modified environmental cues, which directly or indirectly affects plant metabolism. For example, a significant increase in nitrogen (N) content in leaf tissues increases photosynthesis efficiency, which is key to increasing crop yield. Plant metabolism is maintained by these elements with lower fractions of amino acids to regulate plant growth and development. Mixed fractions of other elements at different concentrations signify their importance in plant health regulation. For example, in addition to essential elements, S, Mg, Fe, Cu, Mn, and Na accumulation was higher at all levels.

Any change in amino acid concentration leads to stress conditions, and plants respond differently at different levels by changing their growth patterns.

Moreover, a significant (*p* < 0.05) decrease in vitamin C content and leaf dry matter content and percentage also highlights the importance of micronutrients in plant metabolism.

Table 8 shows that vitamin C content decreased significantly (*p*<0.05) with 2200 mg/L, but increased by 14.21% with 0.22 mg/L as compared to control. In contrast, a significant increase in leaf dry matter content was found with 2200 mg/L and in dry matter percentage (*p* < 0.05) with L-methionine application of 2200 mg/L and 220 mg/L. Moreover, a decrease in the fresh and dry weight of lettuce plants indicates that reduced photosynthetic activity was responsible for the lower accumulation of leaf organic material and reduced growth.


**Table 7.** Effects of L-methionine concentrations on essential macro- and microelements of lettuce leaves.

Macronutrients: N, nitrogen; P, phosphorus; K, potassium; S, sulfur; Ca, Calcium; Mg, magnesium. Micronutrients: Fe, iron; Cu, copper; Mo, molybdenum; Na, sodium; Zn, zinc; Al, aluminum. Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test at (*p* < 0.05).

**Table 8.** Effects of L-methionine concentrations on vitamin C content, leaf dry matter content (LDMC), dry matter percentage (DM%), fresh weight (FW), and dry weight (DW) of lettuce leaves.


Means followed by the same letters are not statistically different from each other, according to Duncan's multiple range test (*p* < 0.05).

As indicated by the outcomes shown in Tables 3 and 8, there was no significant change (*p* < 0.05) observed in plants with or without amino acid treatment. It was observed that all individual amino acid treatments, except for one L-methionine concentration, led to no significant (*p* < 0.05) impact on vitamin C content. The special case was L-methionine at 2200 mg L<sup>−</sup>1, which prompted a decrease in vitamin C content, essentially contrasting with the control plants (*p* < 0.05).

#### **4. Discussion**

Amino acid application is a common practice for horticultural crops worldwide, with the majority of treatments making use of biostimulants with a mixture of amino acids [53]. In our study, we checked the activity of a single amino acid that regulates the nutrient contents involved in growth variables.

Previous investigations have demonstrated that plant developmental cues respond distinctively to the provision of amino acids [54–56]. It is likely that the effects of amino acids on plants rely on the kind of amino acids supplied [56] and the plant cultivars [57]

From our results, we can presume that amino acids (L-methionine, L-tryptophan, and L-glycine) not only make nutrients available to plants but also act as signal transducing molecules [31], as small doses are sufficient for plant development response, while these molecules can act as signals of several beneficial plant physiological processes. Studies demonstrate that amino acids in the form of a foliar spray on plants is a promising technique [38]. In this manner, L-methionine induces more prominent absorption of sulfur and nitrogen in plants, which also depends on the amount applied [26,58–60].

Plants utilize amino acids according to their nutritional needs and genetic background, as well as environmental and developmental cues [61,62]. This might be the reason why amino acid reactions were not consistent in both experiments. Therefore, we can assume that the decreasing effect of L-tryptophan on yield might be due to the inhibitory impact of auxin on vegetative growth. In this association, the reduction of lettuce yield per plant caused by the inhibition effect might be due to the detrimental effect of auxin accumulation stress on growth, the aggravation of mineral nutrient uptake, and the improvement of plant respiration [41]. The distinctive response was reported by Abbas et al. [54], in an investigation on L-tryptophan applied to chickpea at rates of 10−<sup>1</sup> M, 10−<sup>2</sup> M, and 10−<sup>3</sup> M. They found random results with different parameters: root length was increased only in the control compared to the three treatments, the number of nodules increased only with 10−<sup>1</sup> M, and nodule fresh and dry weight decreased with 10−<sup>3</sup> M treatment and increased with the other two compared to control, while control remained nonsignificant. The most pods and highest plant weight were shown with 10−<sup>2</sup> M, but pod weight per plant was significantly affected by all treatments due to the production of phytochromes suitable to chickpea. This experiment may provide evidence for the substantiating inconsistency of lettuce observed in our previous experiment.

However, our results contrast those described in [63], in which numbers of strawberry leaves per plant were significantly higher with the application of L-tryptophan than control.

A few reasons can clarify the positive effects of L-methionine. First, it has a role in maintaining the structure of proteins required for cell division, cell differentiation, and growth. Second, it provides sufficient sulfur and nitrogen according to plant needs. Third, the ability of L-methionine to be converted into polyamines and enlarge by entering the hormone structures [64] allows nitrogen movement between cells and organs [65]. It also functions as a buffer and behaves as a source of carbon and energy [66], and as a precursor of spermidine and gibberellin biosynthesis [43,67], growth regulators, and many secondary metabolites [43]. L-methionine also acts as a growth regulator of cytokinin, brassinosteroids, and auxin, increasing the initiation of roots; helps with the absorption of more nutrients by the plant [39,67], which may stimulate endogenous hormone homeostasis [68,69]; and is required for the development of hairy roots [67] at optimum levels.

Increased L-methionine levels influence phytohormones, which ultimately increases the chlorophyll content and chloroplast development or cytokinins [70,71]. An expected requirement for the prompting of L-methionine application might be the proximity of phytohormones (e.g., auxins and cytokinin). The phytohormones and signaling compounds may improve the photosynthetic activity, leading to better yield. Another possible mechanism involved with the amino acid effect could be related to the stimulation of root growth of treated plants, which may improve water and nutrient uptake capability, leading to yield productivity [68,69], as well as enhanced cell formation and increased fresh and dry matter [72], with increased growth behavior [69,70,73].

Our results demonstrate that high L-methionine concentration reduced plant growth due to damage to the photosynthetic apparatus [61] and blocking of nutrient uptake. Higher levels of this nutrient cause blockage of photosynthesis in stressed environments [65]. Padgett (1996) applied L-methionine to the root zones of chrysanthemum plants, producing a physiological disorder called methionosis, with the typical pattern of a metabolite–antimetabolite relationship [47]. It is thought that in this case, L-methionine, especially because of the large amounts applied, may function as an antimetabolite that interferes with normal amino acid metabolism. In other words, amino acids should be meticulously applied, as they could reduce the percentage of dry weight because they cause swollen, water-filled tissues due to depressed vegetative growth [65,74].

Thus, we speculate that application of L-methionine with 0.22 mg/L in the nutrient solution was sufficient, but other concentrations were too high and might have been a source of stress. Our results are consistent with previous studies [42,75] proposing that the improvement of novel "bio-sound items" ought to continue based on a foundational approach established in chemical synthesis, natural chemistry or biochemistry, and biotechnology connected to genuine plant physiological, agrarian, and environmental constraints. It was proposed that these items should work at low dosages, be biologically and ecologically friendly, and have reproducible advantages in horticultural plant development. The high amount of L-methionine also reduced vitamin C content. Therefore, we conclude that plant metabolism is affected by external N and thus can reflect the changes in N absorption, transport, and metabolism [76]. Similar findings have been reported in Chinese cabbage

and lettuce [77]. The optimal required concentration is essential for optimal growth. It was confirmed from the previous study that when amino acids are added alone, care must be taken, as they can inhibit cell growth [63]. In general, the use of amino acids on plants can improve their capacity of transporting mineral components [66].

In view of the synthetic pathway of vitamin C and the synthesis of ascorbic acid requiring plant climatic changes and the conditions of plant sustenance, it may be hypothesized that whatever factor builds the sugar (or glucose) content in plant tissues can thus increase the vitamin C content [78]. It has been reported that amino acids [43] and nitrogen fertilizers do not impact the vitamin C content in broccoli. Conversely, in cauliflower, when nitrogen fertilizers are extended from 80 kg to 120 kg per ha, the ascorbic acid content is reduced by 7% [79].

Optimizing the amino acid content can bring about different morphogenetic responses; higher concentrations generally inhibit growth in *Cicer arietinum* [80]. The available information from various studies suggests that optimal levels of various amino acids may be species- or genotype-dependent, which needs to be determined before recommending their use [81]. Increasingly, plant ecologists working at all levels have become interested in the role of amino acid nutrition in the lives of plants and determining the proper amount of amino acids suitable for plant growth [82].

The depressive effect of L-tryptophan and high amounts of L-methionine on yield may be attributed to the inhibitory effect of auxin accumulation on vegetative growth, the disturbance in mineral uptake, and/or the enhancement of plant respiration [83].

#### **5. Conclusions**

Taking into consideration the discussion above, it can be inferred that L-methionine increases the chlorophyll content of plants and contributes to the saving of energy, thus boosting the plant yield. L-methionine led to significant increases in observed physiological factors in lettuce leaves at lower concentrations because at high concentrations it affects auxin uptake, which can kill plants. In brief, L-methionine at a concentration of 0.2 mg/L showed the best effect on the growth of lettuce plants. Therefore, we can say that L-methionine can contribute as a suitable substitute for fertilizers to increase crop yield. Future research should concentrate on assessing the mechanisms of how amino acids can influence the genetic transcription of various parameters, including supplement transporters, hormone production, and antioxidant metabolism. Along these lines, it will be possible to acquire the best understanding of the role of amino acids as biostimulants in lettuce plants.

**Author Contributions:** Conceptualization, W.J.; methodology, Q.L. and Y.G.; software, S.K.; validation, S.K.; formal analysis, S.K. and B.N.S.; investigation, S.K.; data curation, S.K.; writing—original draft preparation, S.K.; writing—review and editing, H.W., P.L., H.Y., and Q.L.; visualization, H.Y.; S.K.; supervision, W.J.; project administration, Y.G.; funding acquisition, W.J.

**Funding:** Demonstration of innovative and resource-efficient urban agriculture for multiple benefits in China (National key R&D program of China, (2017YFE0118500).

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Review* **Biostimulants Application in Horticultural Crops under Abiotic Stress Conditions**

#### **Roberta Bulgari, Giulia Franzoni \* and Antonio Ferrante**

Department of Agricultural and Environmental Sciences, Università degli Studi di Milano, via Celoria 2, 20133 Milano, Italy; roberta.bulgari@unimi.it (R.B.); antonio.ferrante@unimi.it (A.F.)

**\*** Correspondence: giulia.franzoni@unimi.it; Tel.: +39-02-503-16593

Received: 10 April 2019; Accepted: 10 June 2019; Published: 12 June 2019

**Abstract:** Abiotic stresses strongly affect plant growth, development, and quality of production; final crop yield can be really compromised if stress occurs in plants' most sensitive phenological phases. Additionally, the increase of crop stress tolerance through genetic improvements requires long breeding programmes and different cultivation environments for crop performance validation. Biostimulants have been proposed as agronomic tools to counteract abiotic stress. Indeed, these products containing bioactive molecules have a beneficial effect on plants and improve their capability to face adverse environmental conditions, acting on primary or secondary metabolism. Many companies are investing in new biostimulant products development and in the identification of the most effective bioactive molecules contained in different kinds of extracts, able to elicit specific plant responses against abiotic stresses. Most of these compounds are unknown and their characterization in term of composition is almost impossible; therefore, they could be classified on the basis of their role in plants. Biostimulants have been generally applied to high-value crops like fruits and vegetables; thus, in this review, we examine and summarise literature on their use on vegetable crops, focusing on their application to counteract the most common environmental stresses.

**Keywords:** plant biostimulant; environmental stress; vegetables

#### **1. Abiotic Stresses**

Plants are continuously subjected to a multitude of stressful events, from seed germination through to the whole life cycle. These stresses are commonly divided into two categories—biotic and abiotic stresses—depending on the nature of the trigger factor. The first are caused by other living organisms, including insects, bacteria, fungi, and weeds that affect plant development and productivity. The second are generally linked with the climatic, edaphic, and physiographic components of the environment, when they are limiting factors of plant growth and survival. The most important abiotic stresses limiting agricultural productivity, almost all over the world, are drought, salinity, non-optimal temperatures, and low soil fertility. Among these, drought, and nutrient deficiencies are major problems, mostly in developing countries where the incomes of rural people depend on agriculture [1]. Actually, in "The State of Food and Agriculture 2007", FAO reported that only 3.5% of the global land area is not affected by some environmental constraints. In 1982, Boyer estimated that yield losses caused by unfavourable environments were as much as 70% [2,3]. Farooq et al. [4] reported that drought induced a reduction of yield between 13% and 94% in several crops, depending on the intensity and duration of the stress. Afterwards, Cramer et al. [5] estimated the impacts of different abiotic stresses on crop production in terms of the percentage of global land area affected, considering the 2000 and 2007 FAO reports. They also referred to the increasing number of publications focused on this topic between 2001 and 2011. The exact impact of these changes on agricultural systems is extremely difficult to predict and it depends on numerous parameters that are all not always

included in predictive models. Even if some projections show that positive and negative outcomes on crop production could be balanced in the medium term, several studies agree that in the long term, the negative ones will prevail [6,7]. Based on future scenarios, adaptation and mitigation are essential to increase the resilience capacity of agricultural systems and to ensure crops yield and quality. Since environmental conditions cannot be controlled, several strategies on different levels are required, such as agronomical techniques or breeding of more tolerant cultivars [8].

In 2010, at the society's annual conference, Vegetable Breeding and Stress Physiology working groups of the American Society for Horticultural Sciences focused particularly on the "Improvement of Horticultural Crops for Abiotic Stress Tolerance" considering the effects of climate change [9]. Up to now, most studies on climate change impacts focus on major crops, and only few papers pay attention to fruit and vegetable in terms of production, quality, and supply chain [10,11]. An important aspect to take into consideration is the effect of the combination of different stressful factors. Most of the time, crops are subjected to several abiotic stresses that occur simultaneously in the field. In these situations, studying the stresses separately is not enough because plant response is unique and cannot be predicted by the reply obtained when each factor is applied individually [12–14]. Moreover, biotic and abiotic components typically interact in an ecosystem. For instance, environmental conditions affect plant-pest interaction in different ways, by decreasing plant tolerance or increasing the risk of pathogen infection [15,16].

Focusing on horticultural species, the tolerance to abiotic stresses is an important trait because their cash value is usually higher than field crops, they require more resources for farming and because they provide a source of many nutrients, fibre, minerals, and carbohydrates, which are essential in a healthy diet [17]. Food and Agriculture Organization (FAO) reports that about 90% of essential vitamin C and 60% of vitamin A for human comes from vegetables. Indeed, low fruit and vegetable intake is a major contributing risk factor to several widespread and debilitating nutritional diseases. According to the Global Burden of Disease Study, 3.4 million deaths can be attributed to low consumption of fruit and 1.8 million to low vegetables diets worldwide [18]. Therefore, growing high-quality vegetables becomes one of the most important goals of current agriculture, in order to meet the needs of the population and the increasing demand for fruit and vegetables. Abiotic stresses do not only affect the yield but also the quality of these products, triggering morphological, physiological and biochemical changes that can alter the visual appearance and/or the nutraceutical value in a way that the product could become unmarketable [19]. Bisbis et al. [11] investigated the double effects of elevated temperature and increased CO2 on the physiology of different vegetables. They observed several responses according to plant species and severity of the stress, taking into consideration the possible adaptation strategies that could be implemented in order to mitigate the effects of climate change. Nonetheless, these mechanisms are still under-researched and should be studied in depth, because not only different species but different cultivars also could respond differently to the same environmental stress. For example, cultivars with low levels of antioxidants are particularly vulnerable to oxidative stress compared to those with high antioxidant activity [20–23]. This aspect has a particular importance as selection criterion in the choice of appropriate cultivars for a specific situation. Oxidative stress is a common phenomenon caused by several adverse conditions; it generally occurs when the balance between the production of reactive oxygen species (ROS) and the quenching activity is upset by a stressful event [24]. Low levels of ROS are normally produced by different reactions during physiological metabolisms like photosynthesis or respiration, and they play an important signaling role in plant growth and development. Their amount dramatically increases under abiotic stress conditions and, if not controlled could result in cellular damage and death. Besides their toxicity to proteins, lipids or nucleic acids, the increased production of ROS under stressful conditions plays a key role in the complex signaling network of plants stress responses. Their concentration is maintained at non-toxic levels by the activity of the antioxidant system: a wide range of enzymatic or non-enzymatic antioxidant molecules are accumulated in plant tissues to quench ROS induced by stress [25–28]. Moreover, the maintenance of this equilibrium is also dependent on numerous factors, such as the timing of stress application,

its intensity and duration. Indeed, moderate or controlled stress conditions could have a positive effect on quality traits of several crops [29]. For example, water deprivation might be a useful crop management strategy to improve the quality of lettuce and fleshy fruits in terms of nutritive and health-promoting value and taste, by stimulating the secondary metabolism and concentration of different phytochemicals such as α-tocopherol, β-carotene, flavonoid and so on [30,31]. Besides the production of ROS scavenging compounds, plants also increase the biosynthesis and accumulation of compatible solutes with an osmoprotective role, like sugars and proline.

Plants generally reply to non-optimal environmental conditions both with short- and long-term adaptation strategies, by the activation and regulation of the expression of specific stress associated genes [32,33].

Since plants are sessile organisms and they have to cope with adverse external conditions; all these mechanisms are essential for their survival. These strategies are effective if they are activated in time, in order to set a defense response and anticipate the environmental changes that might affect plant growth irreversibly. The trade-off between growth and acclimation metabolisms results in a sort of fitness cost for plants, since energy and nutrients normally destined to growth and production are intended for stress responsive mechanisms [34].

Agronomic management conducted in order to enhance plant tolerance towards abiotic stresses evolved over the centuries due to the technologic progress, climate change, scientific knowledge, and farmers' experiences. The choice of the correct cultivar, the best growing period, the sowing density, and the amount of water or fertilizers are some of the most common strategies applied to mitigate the negative effects of abiotic stresses [8]. Protected cultivation is a cropping technique adopted to preserve plants from unfavourable outdoor conditions. It is mainly suited to vegetables and floriculture production in a non-optimal environment, through the control of temperatures, radiation or atmospheric composition. Another agronomical strategy, especially applied in vegetable crops, is soilless cultivation. This approach allows controlling of water and nutrients, avoiding the use of soil for cultivation and all the problems related to it, like poor quality or contamination.

Grafting is an additional tool adopted to counteract environmental stresses and increase tolerance in vegetable crops. This technique is applied especially to high-yielding fruits and vegetables such as cucurbits and solanaceous to enhance tolerance against saline soil, nutrient or water deficiency, heavy metals or pollutants toxicity [35–37].

Agronomical strategies are essential in mitigating the negative effect of several abiotic stresses, but sometimes their application is not enough. Moreover, current experiments aim to transfer one or more genes involved in signaling or regulatory pathways, or genes encoding to molecules, such as osmolytes and antioxidants, conferring tolerance to a specific abiotic stress [38]. Several functional and regulatory genes involved in abiotic stress tolerance have been identified and studied. Results of these studies can be exploited for genetic improvement aiming to introduce tolerance traits in cultivated crops. Since different physiological traits related to stress tolerance are under multigenic control, the manipulation of a single gene generally is not enough. Hence, scientists have paid more attention to regulatory genes, including transcription factors, due to their ability to regulate a vast array of downstream stress-responsive genes at a time [39–41].

However, the huge existing genetic variability among vegetable species, the lack of knowledge about minor cultivars genome, the complex responses triggered by abiotic stress conditions and the limited strategies currently available make genetic improvement really difficult and often inefficient. Moreover, besides the wide diversity of germplasms available, plant tolerance to stress depends both on stress features such as duration, severity, and frequency, as well as the affected tissues and development stages of crops [24,42–44].

Additionally, the increase of crop tolerance through genetic improvements requires many years of work and different cultivation environments that cannot be always taken into consideration. As a result, several new cultivars that can be used by the growers are released each year.

Another technique widely used for developing stress tolerance in plants is in vitro selection. This culture-based tool allows better understanding of several plants' physiological and biochemical responses to adverse environmental conditions. It has been applied specially to obtain salt/ and drought/tolerant lines in a wide range of plant species, including vegetables [45]. In vitro selection is based on the induction of a genetic variation among cells, tissues or organs, their exposure to a stressor, and the subsequent regeneration of the whole organism starting from the surviving cells [46]. Even if in vitro selection is a less expensive and time-saving approach compared with classic molecular engineering, some limitations, mostly concerning the stability of the selected traits and epigenetic adaptation, still exist.

In addition to these strategies, it has been observed that stress tolerance can also be induced by biostimulants or specific bioactive compounds, if they are applied on vegetable crops when they really need to be protected [47–49]. Biostimulant application on horticultural crops under environmental stress conditions will be discussed in detail below.

#### **2. Biostimulants**

Biostimulant products have been considered innovative agronomic tools as demonstrated by the increase of scientific publications and by the constant expansion of their market [50]. France, Italy, and Spain are the leading EU countries in the production of biostimulants [51]. According to a new report by Grand View Research, Inc., the biostimulant market size is expected to reach USD 4.14 billion by 2025 [52]. The complex nature of the composition of these products and the wide range of molecules contained makes it complicated to understand and define which compounds are the most active. The isolation and study of a single component is almost impossible and the efficacy of a biostimulant is not due to a single compound but is the consequence of the synergistic action of different bioactive molecules. Moreover, the application rules and time are not always clear. For all these reasons, the European Commission developed a proposal for a new regulatory framework and a draft for a new fertilizer regulation was prepared in 2016. The amendments to the proposal of the European Commission were adopted by the European Parliament in October 2017, while the legislative resolution on the proposal was approved on 27 March 2019 [53–55].

Plant biostimulants are defined as products obtained from different organic or inorganic substances and/or microorganisms, that are able to improve plant growth, productivity and alleviate the negative effects of abiotic stresses [56,57]. Mineral elements, vitamins, amino acids, and polyand oligosaccharides, trace of natural plant hormones are the most known components. However, it is important to underline that the biostimulant activity must not depend on the product's nutrients or natural plant hormones content. The mechanisms activated by biostimulants are often difficult to identify and are still under investigation [58]. High-throughput phenotyping and omic technologies seem to be useful approaches to understand biostimulants activity and hypothesize a mode of action [59–61]. They can act directly on plant physiology and metabolism by improving soil conditions [62,63]. They are able to modify some molecular processes that allow to improve water and nutrient use efficiency of crops, stimulate plant development, and counteract abiotic stresses [47] by enhancing primary and secondary metabolism [55,61,63].

One of the key points of the discussion is about the application of these products in stressful conditions and their role as nutrients, not with a curative function. In particular, if a product has a direct effect against biotic stresses, it should not be included in the biostimulant category but should be registered as plant protection products.

#### *2.1. Classification of Biostimulants in Categories*

During the years, different authors have proposed several categorizations of biostimulant products on the basis of their main component or mode of action. In many countries outside the European Union, both kinds of information must be reported on the label in order to register these products [55]. The current classification is based on source of raw material, even if this choice does not always

provide the correct information about the biological activity of the product [56]. Thus, biostimulants are classified as these major groups:

*Humic substances*(*HSs*): they include humic acids, fulvic acids and humins. *HSs* are natural constituents of soil organic matter, resulting from the decomposition processes of plants, animals, and microbial residues, but also from the metabolic activity of soil microbes [57]. It has been observed that treatments with humic substances stimulate plants root growth and development [64,65]. This is reflected in a better uptake of nutrients and water, and enhanced tolerance to environmental stresses, [66,67]. How the *HSs* affect plant physiology is not fully understood. This is due to the molecular complexity of these substances and to the abundance and diversity of plants responses altered by their application. Moreover, a strong relationship between medium properties and *HSs* bioactivity has been reported [68]. The positive effects exerted by these complex aggregates could be ascribed both to the hormone-like activity of some of their component and also to IAA-independent mechanisms [69]. For example, like auxins, *HSs* are able to promote plant growth and induce H+ATPase activity in plasma membrane [70–72].

*Seaweed extracts*: seaweeds are a vast group ofmacroscopic, multicellularmarine algae that can be brown, red, and green. They are an important source of organic matter and fertilizer nutrients. Seaweed extracts have been used in agriculture as soil conditioners or plant stimulators. They are applied as foliar spray and are able to enhance plant growth, abiotic stresses tolerance, photosynthetic activity, and resistance to fungi, bacteria and virus, improving yield and productivity of several crops [73–75]. Seaweeds used for biostimulant production contain cytokinins and auxins or other hormone-like substances [76]. They also contain many active mineral and organic compounds, including complex polysaccharides such as laminarin, fucoidan, alginates and plant hormones that contribute to plant growth [77]. Recently the potential application of micro-algae as plant biostimulants has been considered [78–80].

*Hydrolysed proteins and amino acids containing products*: hydrolysed proteins are a mixture of amino acids, peptides, polypeptides and denatured proteins that can be obtained by chemical, enzymatic and thermal hydrolysis of proteins (or by combining these different hydrolysis types) from both plant and animal sources [67,81]. Studies reported that the applications of some commercial protein hydrolysate products from animal origin were phytotoxic, having negative effects on plant growth when compared to a commercial protein hydrolysate of plant origin [82,83]. In another study, Botta et al. [84] observed that lettuce plants treated with an animal-based protein hydrolysed had a higher fresh and dry weight compared with the control. Generally, they can induce plant defense responses and increase plant tolerance to many abiotic stresses, as reported by several authors [85–88].

*Microorganisms*: this group includes bacteria, yeast, filamentous fungi, and micro-algae. They are isolated from soil, plants, water, and composted manures or other organic materials. They are applied to soil to increase crop productivity through metabolic activities. They enhance the uptake of nutrients through nitrogen fixation and the solubilization of nutrients, they modify a hormonal status by inducing plant hormones biosynthesis such as auxins, cytokinins, etc.; they also enhance tolerance to abiotic stresses and produce volatile organic compounds (VOCs), which may also have a direct effect on plants. Plant growth-promoting rhizobacteria (PGPR) are able to ameliorate plant responses to abiotic stresses stimulating physical, chemical and biological activities [89,90]. Positive effects are given by microorganisms that form a protective biofilm on root surface enhancing nutrient and water uptake.

Another category of biostimulants includes those derived from extracts of food waste or industrial waste streams, composts and compost extracts, manures, vermicompost, aquaculture residues and waste streams, and sewage treatments among others [91]. Biostimulants derived from agro-industrial by-products were reported to be effective in improving plant productivity, increasing the synthesis of secondary compounds involved in several plant physiological responses, and enhancing the activity of the enzyme phenylalanine ammonia lyase (PAL E.C. 4.3.1.5) [92]. The effect of biostimulant application on PAL activity and on the expression of genes encoding for this enzyme was observed by several authors [56,88,89] and references therein, even if at present it is not possible to define if this is a direct or indirect effect. Because of the diversity of source materials and extraction technologies, the mode of action of these products is not easily determined [55]. The use of by-products as raw material that can be transformed into fertilizing

products is the idea underlying the new fertiliser regulation and the Circular Economy Action Plan, which is focused on reaching a sustainable agriculture. The guidelines for fertiliser regulation, the need to produce in a more environmentally friendly cultivation system maintaining good crop yield and quality, the increase in price of synthetic fertilizer, the withdrawn of several agrochemicals and the multifaceted effects on plants or soil of biostimulants are favouring the expansion of this market.

A new category of biostimulant products, including nanoparticles and nanomaterials, has been recently proposed by Juárez-Maldonado et al. [93]. Nanoparticles and nanomaterials are usually defined as particles with dimensions between about 1 nm and 100 nm that show properties that are not found in their bulk form. They are able to modify the quality of the production and the tolerance to abiotic stresses when applied in small quantities as foliar spray or in nutrient solution, also in vegetable crops [94–97]. Their biostimulant properties seems to be associated with the structure and nature of the materials. The interaction between plant and nanoparticles and nanomaterials surfaces can positively affect ions and metabolites transport and receptors activity by modifying the surrounding environment in terms of energy and charges. This activity is not dependent on chemical composition. Moreover, nanoparticles and nanomaterials release chemical elements like iron or carbon that could be useful for plant when are metabolised.

A study showed that application of zinc oxide nanoparticles on tomato as soil amendment or by foliar spray increased plant height, chlorophyll and total soluble protein content [98].

#### *2.2. E*ff*ect of Biostimulants on Chlorophyll Content, Photosynthesis and Growth in Vegetables*

Biostimulants can be used in vegetable cultivation to improve productivity and yield, and to enhance plant health and tolerance to stress factors. Indeed, they have positive effects on plant metabolism, both in optimal and sub-optimal environmental conditions.

Many authors have observed that plant based biostimulants and seaweed extracts often increase the colour of leaves by stimulating chlorophyll biosynthesis or reducing its degradation [99,100]. Leaf colour is an important quality parameter in vegetable crops because it contributes to the visual appearance of the product, especially in leafy vegetables for which the greenness influences the consumer's appeal. In addition, a higher chlorophyll content also allows for a greater photosynthetic activity of leaves. High concentration of leaf pigments (chlorophyll and carotenoids) has been observed after biostimulant treatments in rocket [101,102], in lettuce, and endive by Bulgari et al. [103]. Amino acids or seaweed extract application had positive effects on photosynthetic pigments, P and K content, fresh and dry weight of celeriac leaves [104]. Similar results have been observed after root inoculation with several plant growth promoting bacteria (PGPR) in broccoli (*Brassica oleracea* 'italica') using *Bacillus cereus*, *Brevibacillus reuszeri*, and *Rhizobium rubi* [105], and tomato under non-stressful conditions treated with PGPRs belonging to the genera *Bacillus*, *Pseudomonas* and *Azotobacter* [106], in strawberry (*Fragaria ananassa*) with five PGPRs (*Bacillus subtilis, Bacillus atrophaeus, Bacillus spharicus* subgroup, *Staphylococcus kloosii*, and *Kocuria erythromyxa*) [107] and also in lettuce grown under salt stress after inoculation with *Serratia* sp., *Rhizobium* sp., and *Azospirillum* [108,109]. Brown seaweeds are widely used as a biostimulant products to improve plant growth, and recently a phenolic compound isolated from *Ecklonia maxima* showed stimulatory effects in cabbage plants, improving photosynthetic pigments concentration, phytochemicals and myrosinase activity [110].

Abdalla [111] reported that moringa leaf extracts increased vegetative growth, chlorophyll content, total sugars, phenols, ascorbic acid, and photosynthetic rate of rocket salad. Similar effects have been observed in fennel [112,113] and squash under water stress condition (plants under a deficit irrigation of 80% or 60% ETc) [114]. In tomato plants it led to a greater fruit weight, volume and firmness, and enhanced titratableacidity, chlorophyll and ascorbic acid content [115].

Luziatelli et al. [116] recently found that different vegetal-derived bioactive compounds significantly increased the chlorophyll content and fresh weight of lettuce. Kulkarni et al. [117] investigated the promoting effect of bioactive molecules derived from smoke and seaweed in spinach and they observed that morphological, physiological and biochemical parameters including growth, chlorophyll and carotenoids content were positively improved.

Broccoli plants were significantly affected by two different products: Goemar BM86 and Seasol. The content of micro- and macro-nutrients increased, and also the leaf area, stem diameter and biomass, as reported by Gajc-Wolska et al. [74] and Mattner et al. [118].

Paradikovi´c et al. [119] studied the effect of four different commercial biostimulants (Radifarm, Megafol, Viva, and Benefit), containing amino acid, polysaccharides and organic acids as active compounds on pepper plants and observed an increase in both yield and fruit quality. Radifarm and Viva treatments also affected tomato plants, stimulating the root apparatus in optimal and drought condition, respectively [120,121].

Recently, a sago bagasse hydrolysate was tested on tomato plants. The product showed a growth promoting ability as observed by the higher seed germination and protein and sugar content compared to the control. Moreover, the expression of the genes related to carbon and nitrogen metabolisms increased [122].

#### *2.3. Biostimulants and Crop Tolerance to Abiotic Stresses*

Table 1 is a summary of biostimulant products or bioactive molecules from different origins that have been evaluated for amelioration of abiotic stresses in several vegetables species. The biostimulants effectiveness to counteract the stressful condition depends on several factors, such as timing of application and their mode of action. The application of biostimulants can be carried out with different timings: before the stress affects the cultivation, during the stress, or even after. They could be applied on seeds, when plants are in early stages of growth, or when crops are fully developed, depending on the desired results [123]. As general consideration, biostimulants that contain anti-stress compounds, such as proline or glutamic acid, can be applied when the stress occurs or during stress conditions. On the contrary, those that are involved in the activation of bioactive compounds biosynthesis must be applied before the stress occurs. Proper timing of application during crop development differs from species to species and it also depends on the most critical phases for crop productivity. Thus, the identification of the right time of biostimulant application is as important as the determination of the exact dose, in order to avoid waste of product, high production costs, and unexpected results. Biostimulants can be applied as foliar spray or to the roots, at sowing for protecting the seedling in the early development stages, in a floating system nutrient solution or during blooming or fruit setting. There is no general recipe that works for a crop species and in each stress situation.

The protective role of biostimulants on plants has been increasingly studied. These products are able to counteract environmental stress such as water deficit, soil salinization, and exposure to sub-optimal growth temperatures in several ways [47,56,124,125]: They improve plant performance, enhance plant growth and productivity, interact with several processes involved in plant responses to stress, and increase the accumulation of antioxidant compounds that allow decrease in plant stress sensitivity.

More recent results of interest on vegetable crops tolerance have been obtained after the application of different exogenous treatments. Cao et al. [126] reported that a lower red to far-red ration improved tomato seedling tolerance to salt stress, acting on phytochrome activity. Mertinez et al. [127] showed positive results obtained after the application of exogenous melatonin in tomato plants grown under a combination of salinity and heat. Another interesting approach to induce tolerance to abiotic stresses is soaking plant seeds with different compounds, synthetic or natural. This strategy is generally called seed priming and has been deeply reviewed by Asharaf et al. [128].

#### 2.3.1. Biostimulants and Cold or Chilling Stress

Low temperatures reduce plant metabolism and delay physiological responses. A reduced metabolism, consequent to cold stress, leads to an inhibition of the activity of photosystem II, called photoinhibition. Cold induces damages to cell membranes with destabilization of the phospholipid layers.

In tomato, cold tolerance has been enhanced by the application of psychrotolerant soil bacteria. Several strains have been isolated from soil during winter conditions and used as a cold protectant. Tomato treated with these psychrotolerant bacteria showed higher seeds germination, reduced membrane damage, and antioxidant systems activation when exposed to chilling temperatures [129,130]. These soil bacteria can be considered as putative biostimulants for protecting plants against cold stress. Since low temperature causes stress to plant, especially during transplant, Marfà et al. [131] studied the effect of an enzymatic hydrolysates obtained from animal haemoglobin on strawberry plants in the firsts growing stages. They observed an increase in roots biomass and in the early production of fruit. The same product was also tested on lettuce plants subjected to cold stress and an increase in fresh weight, dry weight, specific leaf area, and relative growth rate was observed [132].

External applications of an amino acid biostimulant (Terra-Sorb® Foliar) on lettuce plants grown in different cold situations led to an increase in fresh weight and to an higher stomatal conductance [84]. A typical plants response to stress is the accumulation of compatible osmolytes, such as amino acids, which confer tolerance. The exogenous application of amino acids has the benefit of avoiding protein breakdown and saving energy resources in plants, even if the exact mechanism of action is not fully understood. Pepper (*Capsicum annuum*) seedlings were treated with 5-aminolevulinic acid in order to improve chilling tolerance through three different methods—soaking the seeds, spraying the leaves or drenching the soil. All the applications showed good effects in terms of stress tolerance. Fresh biomass, proline, sucrose, and water content were significantly higher while membrane permeability was reduced [133].

Positive effects on coriander plant grown in cold vegetative chambers have been observed in response to Asahi SL or Goemar Gateo (Arysta Life Science) treatments [124]. Results obtained by the study of stress indicators such as antioxidant activity, photosynthetic pigment concentration and activity, hydrogen peroxide and malondialdehyde amount showed that biostimulant application affected different metabolic pathways in a positive way, leading stressed plants to a phase of acclimation to low temperature. The biostimulant action against cold stress usually increases the accumulation of osmotic molecules by stimulating the biosynthetic pathways that lead to the cold protectant substances. These biostimulants also increase membrane thermostability, reducing the chilling injury.

#### 2.3.2. Biostimulants and Heat Stress

Global warming and the projection of a rising temperature have a negative impact on agriculture [134,135]. High temperatures could induce several damages to plant cells, disturbing proteins synthesis and activity, inactivating enzymes and damaging membranes. The range between 30 ◦C and 45 ◦C is the optimal temperature for structural integrity and enzymal activity, which are irreversibly denatured when temperature increases above 60 ◦C. As a consequence, physiological activities like photosynthesis or respiration are affected. An overproduction of toxic compounds, like reactive oxygen species, causing oxidative stress, is one of the most frequent throwbacks [136]. As response, plants start synthesizing compatibles solutes in order to maintain cell homeostasis and turgor, organize proteins, and cellular structures. Moreover, they generally close stomata and increase the number of trachomatous, in order to prevent water loss. Also, at the molecular level there is a variation of the expression of genes involved in the synthesis or activity of antioxidant enzymes related to ROS scavenging, osmolytes or transporters. Temperature above optimum inhibits seeds germination and retards plant growth. Heat stress could negatively affect the yield by interfering with the reproductive phase, decreasing pollen vitality and germination, inhibiting flower differentiation and development and reducing fruit set, which ultimately reduces growth and yield.

Tomato is considered one of the most sensitive species to non-optimal temperatures, and heat stress often results in long style lengths and in a decreased fruit set [137]. There is little information in the literature about treatments specifically applied to vegetable crops exclusively against high temperature since, most of the time, heat stress is combined with drought or salinity. The application of brassinosteroids on tomato [138] and snap bean [139] has resulted in a higher biomass accumulation and net photosynthesis rate, increased growth and quality of snap bean pod in terms of NPK content and the total free amino acids levels in leaves. This might be due to the protective role of brassinosteroids on the photosynthetic apparatus from oxidative stress, increasing the ability to regenerate RuBP and carboxylation efficiency.

Nahar et al. [140] investigated the effect of exogenous application of glutathione against heat stress. Mung bean seedlings treated before their exposition to high temperature, showed a reduced oxidative stress and methylglyoxal content, a reactive compound that damages cells. This results in a more efficient antioxidant defense system. Pre-treatment with glutathione enhanced tolerance to short-term heat stress, improving plant physiological adaptation. For example, leaf relative water content and turgidity, which usually decreases under high temperature, were protected. Positive effect on mung bean has been observed in response to the application of nitric oxide [141] and ascorbic acid [142]. Nitric oxide treatment resulted in a promotion of photosynthetic activity, increasing the quantum maximum efficiency of PS2. It also affected electrolyte leakage, leading to a better cell membrane integrity. Oxidative stress, lipid peroxidation, and H2O2 content were decreased and antioxidant enzyme activity was restored. Similar results have been obtained after the application of proline and abscisic acid on chickpea [143,144]. Chickpea is sensitive to high temperature that generally leads to yield and quality losses. After treatments, membrane damage, measured as electrolyte leakage, MDA and H2O2 levels was decreased, while leaf water content was increased. These effects might be related with the osmoprotectant role of proline and with the accumulation of osmolytes after ABA treatments. Treated plants also showed a high chlorophyll content and this result, which has already been seen in other experiment with exogenous proline, could be related to membrane stability. The activity of oxidative metabolism was enhanced in treated plants, as expected also by the less oxidative damage of cells.

As discussed above, melatonin treatment exerts a positive effect to counteract chilling stress in coriander plants; otherwise, Martinetz et al. [127] found that melatonin treatments also have a protective role against the combination of heat and salt stress in tomato plants. Biostimulant treatments used against heat stress protect cell membranes by increasing their stability and reduce or avoid the accumulation of ROS.

#### 2.3.3. Biostimulants and Salinity Stress

Among abiotic stresses, salinity is one of the main damaging factors affecting plant growth and metabolism as an effect of osmotic stress caused by salt. Sodium chloride (NaCl) is the more abundant salt presents in saline environments and is toxic in higher concentrations [145]. It happens especially near the coasts, where crops are frequently irrigated with saline water [85,146]. In many Mediterranean areas, the problem of seawater intrusion may cause a reduction of 50% of yield in lettuce cultivation, as reported by Miceli et al. [147]. A significant reduction of both fresh weight and chlorophyll content is a typical effect of salinity condition on plants and was observed also in spinach [148], in bean [149] and other crops [150]. Besides, chlorophyll content is a central parameter of the product quality particularly in green leafy vegetable, not only in terms of plant physiology status but also from a market point of view. This is a huge problem for vegetable crops where the edible parts are leaves, sprouts or flower buds. Consumers choices, in fact, are guided mostly by the visual appearance of products, hence a less green leafy vegetable or a malformed fruit are generally not accepted.

Salt stress causes a nutrient imbalance due to the limited uptake of the nutrients from the soil, threatening the nutritional quality of horticultural crops. Nutrient availability is compromised by salinity that causes several disorders such as competitive uptake with other ions like Ca2+, P and K, mobility problems within the plant and a reduced water potential [151–155]. The solubility of micronutrients such as Cu, Fe, Mn, Mo and Zn is also affected by the pH of the soil solution, and in saline condition their availability is very low. Bano et al. [156] reported an important reduction of total phenolics, total soluble proteins and a suppressed activity of catalase, superoxide dismutase and peroxidase in carrot under saline condition. Salt stress could also alter several metabolic processes in plants, such as photosynthesis [157,158], respiration [159], phytohormone regulation, protein biosynthesis, nitrogen assimilation [160], and can also generate secondary oxidative stress [146,161]. It generally leads to a decrease of production and to a lower quality of the final product, due to an

inhibition of leaves and roots growth and a change in leaf colour [17]. To verify the effects deriving from the applications of biostimulants, several trials on lettuce plants under salt stress were performed, since this crop is considered moderately sensitive to salinity.

Lucini et al. [85] showed that a plant-derived protein hydrolysate improved tolerance to salinity in lettuce plants, increasing yield and dry weight. Treated plants also have a higher performance and an increased maximum quantum efficiency of PS2 compared to the control. Similar results have been recently observed in lettuce plants in response to the application of an organic commercial biostimulant named Retrosal® [162].

Several experiments have been carried out using different PGPR that are able to enhance abiotic stress tolerance. Inoculation with *Azospirillum brasilense* showed positive results on lettuce [163,164], sweet pepper [165], chickpea and faba beans [166] grown under salty environment. Lettuce fresh weight, dry weight, ascorbic acid content, and germination percentage were increased; also, the visual appearance of the final product was better because of higher chlorophyll levels. In chickpeas and faba beans, the inoculation relieved the stress caused by salinity, increasing the root and shoot growth compared with the non-inoculated plants. Sweet pepper is a salt-sensitive crop and inoculation showed positive effect mitigating deleterious effects of NaCl. Dry weight, indeed, was higher than non-inoculated plants under several salt concentrations. Moreover, the inoculation also increased the CO2 assimilation rate. A similar result has been obtained by Cordovilla et al. [167] applying two different *Rhizobium* strain on faba bean and pea plants. Pea plants inoculated with tolerant strain showed no reduction by salt stress condition in shoot and roots dry weight. The same strain was, however, not effective on faba beans. These results highlight the variation existing inter and intra species, and the difficulty in improving tolerance through selection and breeding. A comparable experiment has been carried out by Mayak et al. [168] on tomato seedling. They tested several strains of *rhizobacterium* and found that plants inoculated with *Achromobacter piechaudii* and irrigated with saline water had a higher fresh and dry weights and an increased water use efficiency. Yildirim et al. [169] obtained similar results in squash with the application of several biological products based on the *Bacillus* and *Trichoderma* species.

It is known that humic acids have a lot of beneficial effect stimulating shoot and root growth and improving environmental stress tolerance even if the exact mechanism of action is not completely clear. These activities were confirmed in several vegetable crops like sweet pepper [170], beans [171] and cucumber [172] grown under different salt stress conditions.

Bioactive compounds present in seaweed extracts are able to improve plant tolerance against abiotic stresses too. Two seaweed-based plant biostimulants containing *Ascophyllum nodosum* named Super Fifty® and Acadian were applied respectively on lettuce [173] and strawberry [174] and were associated with a significant increase in yield and root dry weight, despite the adverse salinity condition.

Sulphated exopolysaccharides extracted from the microalgae *Dunaliella salina* were applied on tomato plants to investigate their potential effect alleviating salt stress damages. Results obtained showed that treatment enhance plant growth, antioxidant enzymes activities and several metabolic mechanisms related to jasmonic acid pathway [175].

The application of seaweed extracts from *Sargassum muticum* and *Jania rubens* significantly alleviated the negative effects of salt through regulation of amino acids metabolism, ionic content balanced and improved antioxidant defence in chickpeas plants. Amino acids such as serine, threonine, proline and aspartic acid were identified in roots as responsible for salt stress amelioration [176].

Besides lettuce and pepper, bean is also considered a salt sensitive plant but in most developing countries it is cultivated in saline conditions. Several plant extracts based on licorice root, *Moringa oleifera* or maize grain have been tested on common bean by Egyptian researchers [177–181]. They observed that soaking seeds in propolis or maize grain extract improves seed germination percentage, stability of cell membrane and relative water potential under saline conditions. Antioxidant system activity was increased while lipid peroxidation and electrolyte leakage were reduced compared with the control plants. *Moringa oleifera* leaf extract, used alone or in combination with salicylic acid, and administered

as foliar spray or as seed soaking, improved several physiochemical parameters as chlorophyll and carotenoids concentration, total soluble sugars and ascorbic acid content. A very similar trial has been carried out with licorice root extract and best results have been recorded integrating seed soaking and foliar spray applications.

A recent study highlighted the ability of a bee-honey based biostimulant to improve the tolerance of onion plants to salinity stress. Indeed, treated plants showed higher biomass, bulb yield, and photosynthetic pigments. Moreover, the osmoprotectans content as proline, soluble sugars and total free amino acids, the membrane stability index and the enzymatic and non-enzymatic antioxidant activity were enhanced [182]. Hence, biostimulants applied in case of salinity stress induce the accumulation of osmolytes, in order to enhance the cell osmotic potential and the level of protective molecules against oxidative stress.

#### 2.3.4. Biostimulants and Drought Stress

Abiotic stresses are closely connected with the problem of resources availability and farmers are frequently forced to work in suboptimal conditions. A more sustainable use of resources also concerns water availability, a critical growing factor. The increasing use of aquifer-based irrigation by farmers worldwide poses a serious threat to the long-term sustainability of the agricultural system. Over-utilization of this dwindling water supply is leading to an ever-enlarging area in which productive farming itself has ceased or is threatened. Moreover, the increase of irrigation leads to a higher risk of soil salinization. Scientists generally agree with the perspective that several regions could become arid due to the negative impacts of global climate change on water resources [183]. Since one of the main effects of biostimulants is to improve water use efficiency, their application could be a possible strategy to reduce the amount of water added to crops [184]. Drought stress strongly influences plant gas exchange changing photosynthetic and transpiration rates, which are directly linked to yield. Application of *Ascophyllum nodosum* on broccoli [185] and spinach [186] enhanced gas exchange through the reduction of stomatal closure, resulting in increased plant resistance to water stress. Leaf yellowing is another common symptom of drought stress due to chlorophyll degradation during leaf senescence and is used as reliable indicator of metabolic and energetic imbalance in plants under stress. Biostimulant treatments with *A. nodosum* increased total chlorophyll content in tomato leaves [187]. A reduction of water loss, wilting damages and 3-carbon dialdehyde MDA after biostimulant applications were observed. Similar results have been obtained by Petrozza et al. [188] in responses to Megafol treatments in tomato plants. The results revealed that treated plants were healthier than non-treated ones in terms of biomass and chlorophyll fluorescence. Moreover, plants treated with the biostimulant product were able to recover more quickly when they had access to water. The expression of two drought stress marker genes was analysed and the results obtained showed that treated plants were experiencing a low level of water stress.

Sometimes, water stress in plants is caused by bacterial infection clogging xylem vessels and preventing water flow. Romero et al. [189] demonstrated that treatments with *Azospirillum brasilense*, a strain isolated in arid environments, delayed wilting of tomato plants. Treated plants, indeed, showed a high xylem vessels area, resulting in a more efficient water transport from the soil to the leaves. On the other hand, there are several strains of bacteria populating soil promoting plant growth through its metabolic activities and plant interactions. They produce exopolysaccharides, phytohormones, 1-aminocyclopropane-1-carboxylate (ACC) deaminase, volatile compounds, inducing several metabolic plant responses as accumulation of osmolytes and antioxidants, or up or down regulation of stress responsive genes and alteration in root morphology leading to a tolerance of water stress [190,191]. Some examples are reported below. Tomato seedlings treated with *Achromobacter piechaudii* were stimulated to accumulate biomass during the stress period and, the amount of ethylene that usually has negative effects on membrane status was lower than control [168].

Arshad et al. [192] investigated the growth of two plants promoting rhizobacteria on pea (*Pisum sativum*) crop grown under drought stress condition in different phenological phases. They observed that PGPR containing ACC-deaminase, a precursor of ethylene, significantly decreased the stress effects on growth and yield too. Positive results in terms of antioxidant and photosynthetic pigments activity have been collected in basil plants treated with *Pseudomonas sp*. under water stress conditions [193].

Seaweed extracts are already largely used for cultivated plant treatments and most of them contain plant growth hormones, auxins, abscisic acid, cytokinins, gibberellins, polyamines, oligosaccharides, betaines and brassinosteroids. A micro-algae-based biostimulant with known composition was tested on water stressed tomato plants. Results revealed that biostimulant application reduced the damaging effects of stress, increased plant height, root length, and enhanced the number and area of the leaves [78]. Biostimulants are capable of reducing drought injures, are able to enhance the biosynthesis of osmolytes and antioxidants against ROS, such as observed for salinity stress, and of plant hormones, like abscisic acid, regulating transpiration and avoiding excessive water losses.

#### 2.3.5. Biostimulants and Nutrient Deficiency

One of the roles ascribed to biostimulant products is the ability to increase nutrient uptake [53] through different strategies. For instance, they are able to change soil structure or nutrient solubility, modify roots morphology directly or ameliorate nutrient transport in plants [194]. Their application might be really useful in poor soil conditions and in low input horticultural cultivation systems [195]. Indeed, soil nutrient imbalance is an increasing problem for farmers that spend a lot of money every year on fertilizers to resume soil fertility. All these mechanisms result in better nutrient use efficiency for both micro- and micro-nutrients.

Several experiments have been performed to investigate if the application of biostimulants allows a reduction of fertilizers without affecting crop yield and quality.

Koleška et al. [196] showed that the application of a biostimulant product named Viva® on tomato plants, growing under reduced NPK nutrition, help counteract the negative effects of nutrient deficiency. For example, lycopene and chlorophyll content that is usually affected by the availability of macronutrients was preserved in treated plants grown with NPK reduction. Moreover, biostimulant application helped maintain cell homeostasis and prevent oxidative stress. A similar experiment was performed by Anjum et al. [197] on garlic plants grown with half of the recommended dose of nutrients. Garlic growth and yield were positively affected by the biostimulant application in combination with a low dose of macronutrients.

A seaweed-based product (Kelpak®) has been tested on okra seedlings grown with different nutrient deficiencies [198]. Treatments were applied three times a week and were compared with a polyamine solution treatment. Plants treated with the biostimulant showed an increase in growth parameters, such as shoot length, stem thickness, leaves and roots numbers, and fresh weight under phosphorous and potassium deficiency. Kelpak® efficacy might be due to the combination of auxins, cytokinins and polyamines contained in the product.

Spinelli et al. [199] measured the effects of another commercial seaweed extract, named Actiwave® on the vegetative and productive performance of strawberry plants grown on an iron deficient substrate. They found that vegetative growth, chlorophyll content, stomatal density and photosynthetic rate were enhanced after biostimulant treatment. Fruit production and weight were also increased. Nutrient uptake might have been positively influenced by the more developed root system of treated plants. Treatment also contrasted the negative effects of iron chlorosis and this could be linked to betaine contained in this product.

The positive effects of seaweed extracts are usually ascribed to their polysaccharide content that helps the soil structure; nevertheless, Vernieri et al. [102] obtained good results by applying Actiwave in a hydroponic system with different concentrations of nutrient solutions. Yield and leaf area were higher in rocket plants grown with the lowest nutrient concentration, indicating a better nutrient use efficiency.

Most of the biostimulant contains a mixture of different amino acids and short peptides that are usually called protein hydrolysates. They have a positive effect on plant growth and protection against several stresses. The Cerdán et al. [200] study showed that amino acids origin might influence the efficacy of the product. Tomato plants grown under iron deficiency conditions and treated with two products containing amino acids from plant and animal origin showed different responses. Plant-derived amino acids promoted growth and chlorophyll content both in controlled and iron deficiency conditions. This effect might be ascribed to glutamic acid content. Indeed, this amino acid plays an important role in nitrogen metabolism [201] and chlorophyll biosynthesis [202].

Nutrient imbalance might be the cause of several disorders during plant growth and development. Blossom-end rot in pepper is usually caused by a local calcium deficiency in young fruits. Parađikovi´c et al. [203] tested four different biostimulant products for their effects on yield and BER incidence on pepper. They also evaluated the application as foliar spray or in a nutrient solution of the same products. The results obtained revealed that biostimulants applications helped to reduce the occurrence of BER and increase yield. Moreover, nutrient accumulation in fruits and leaves was promoted by the treatments.

These experiments revealed that biostimulant products cannot totally replace fertilizers but could be really useful to reduce the amount of mineral nutrition or help in nutrient deficiency and imbalanced situations. For example, in the floating system cultivation of baby leaf such as rocket, the nutrient solution can be reduced by 75% of Hoagland's solution [101].

The biostimulants that help reduce nutrient deficiencies usually improve crops nutrient uptake by increasing root biomass, nutrient transport/translocation, and enzyme activities involved in nutrient assimilation.

#### **3. Conclusions and Future Prospects**

This review reports the progress on the recent development of biostimulant products with special emphasis on their effects, improving tolerance to abiotic stresses in vegetable crops. During their life cycle, crops are often exposed to abiotic stresses, acting individually or in combination, which could dramatically reduce the yield and quality of products. Biostimulants could represent an effective and sustainable tool to enhance plant growth and productiveness, improving tolerance against abiotic stresses. In fact, biostimulants have been successfully applied for:


It is important to consider that the complex and variable nature of raw materials used for their production and the heterogeneous mixture of components of the final product can make it difficult to attribute a specific mode of action to each biostimulant. The situation is further complicated by the high number of plants, bacteria and in general, substances included into the category of plant biostimulants. For example, two products obtained by two different plants would fall in the same category, but their effects and their mode of action might be completely different. Moreover, the opposite situation may occur; the same product may produce different effects when applied on different plants. This could be related to the genetic variability among species, variety or cultivars. In addition, the biostimulant activity of a product may also depend on the nature and severity of the abiotic stress.

It must also be considered that trying to link a specific mode of action only to the main component of a product might be a mistake because it would be like excluding the effect of the molecules that are presents in small quantities or in traces, but it is known that the efficacy of biostimulant products is the result of a synergistic or antagonistic effect of many components. Furthermore, our understanding of the mode of action also depends on the amount of information provided by scientific papers, on the numbers of analyses performed, and on their investigation level. The availability of innovative research tools will surely improve the knowledge of biostimulant composition, but this information will not be exhaustive. Therefore, the biostimulant mode of action can be understood through plant responses at the physiological, biochemical, and molecular levels.





↓protein oxidation

**Table 1.** *Cont.*

158




**Table 1.** *Cont.*


#### **Table 1.** *Cont.*


**Table 1.** *Cont.* POX peroxidase; GR glutathione reductase; HI harvest index; ABA abscisic acid; ETR electron transport rate. The symbol ↑ means an increase or ↓ a decrease of the parameter measured.

The symbol ×

represents how many times the treatment was applied.

**Author Contributions:** Conceptualization, R.B., G.F., A.F.; writing—original draft preparation, R.B. G.F.; writing—review and editing, A.F.

**Funding:** This research received no external funding.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Review*

### **Plant Biostimulants: Importance of the Quality and Yield of Horticultural Crops and the Improvement of Plant Tolerance to Abiotic Stress—A Review**

#### **Magdalena Drobek, Magdalena Fr ˛ac and Justyna Cybulska \***

Institute of Agrophysics, Polish Academy of Sciences, Do´swiadczalna 4, 20-290 Lublin, Poland; m.drobek@ipan.lublin.pl (M.D.); m.frac@ipan.lublin.pl (M.F.)

**\*** Correspondence: j.cybulska@ipan.lublin.pl; Tel.: +48-81-744-5061

Received: 26 April 2019; Accepted: 20 June 2019; Published: 24 June 2019

**Abstract:** Biostimulants are among the natural preparations that improve the general health, vitality, and growth of plants and protect them against infections. They can be successfully used in both agri- and horticultural crops. The main active substances used in such preparations are humic and fulvic acids, protein hydrolysates, compounds containing nitrogen, seaweed extracts, beneficial fungi, and bacteria. Biostimulant formulations may be single- or multi-component, but the synergic action of several different components has been observed. Many groups of biostimulants have been distinguished through their method of application (soil, foliar), the material from which they were produced (plant, animal), or the process by which they were created (hydrolysis, fermentation, extraction). Natural soil stimulants can induce the development of beneficial soil organisms that provide substrates for plant growth. The use of natural preparations that are not harmful to the environment is particularly important in connection with the progressive processes of soil degradation and atmospheric pollution. This review gives an overview of the importance and influence of different natural plant biostimulants on both the yield and quality of crops.

**Keywords:** fruit quality; plants biostimulants; yielding

#### **1. Introduction**

The quality and quantity of crops are influenced by both biotic and abiotic factors. Quality may be defined as a set of agronomic (e.g., fruit size, yield, resistance to bacteria and fungi) and organoleptic (e.g., colour, shape, firmness) properties as well as nutrient and vitamin content [1]. The abiotic factors include soil composition, extreme salinity, acidity, high and low temperatures, drought, pollution, humidity, rain, wind, or ultraviolet radiation. Stress caused by unfavourable stimuli can significantly reduce harvest yields because plants respond by using their energy reserves to fight stress instead of concentrating on yielding. Biotic factors include various bacteria, fungi, or viruses that are the cause of numerous plant diseases. Fungal and bacterial infections may not only reduce yield but may also lead to the loss of the entire harvest. To prevent this, various types of plant protection products are used. In accordance with the recommendations of the European Union [2], chemical and mineral plant protection agents are intended to be slowly replaced by natural preparations. The reason for this is the adverse influence of chemical and mineral plant protection agents on the natural environment, as well as on the health benefits of plant crops. Moreover, artificial fertilizers are responsible for the eutrophication of many bodies of water. This results in the formation of dead zones devoid of living organisms. The Baltic Sea alone is distinguished by having oxygen-free zones making up around 60,000 km<sup>2</sup> of area caused by water pollution due to fertilizers. This area constitutes, on average, 3.5% of the catchment area of the Baltic Sea [3]. The effects of fertilizers have an unfavourable effect on algae, plants, animals, and people. Due to the fact that man is a higher-order consumer, people are particularly severely exposed to the harmful effects of fertilizer compounds accumulated at the lower levels of the food chain. Harmful compounds from fertilizers may weaken enzymes or interfere with protein production or vitamin absorption in the human body [3]. Natural preparations called biostimulants increase the efficiency of nutrient utilization and tolerance to abiotic stress and improve the quality of crops [4]. Biostimulants include organic and non-organic substances and/or microorganisms [5]. Farmers who manage organic farms are also eager to use natural stimulants to improve crop quality [6]. Increasing consumer awareness concerning healthy food favours the enhancement of the significance of organic farming [7].

The effects of the stimulators may be multifaceted. The effects of their activities vary depending on the type of biostimulant used and the plant variety. However, it should be noted that most of them have a beneficial effect on crops [8].

#### **2. Definitions and Classification of Biostimulants**

Biostimulants can be treated as an additive to fertilizers and support the uptake of nutrients, promote plant growth, and increase tolerance to abiotic stress [9]. The definition of biostimulants is wide and not sufficiently precise. However, there are two main features that distinguish biostimulants from other growth and plant-protection agents. A biostimulant may be any substance or mixture of substances of natural origin or microorganism which improves the condition of crops without causing adverse side effects [10].

Enzymes, proteins, amino acids, micronutrients, and other compounds may be used as biostimulants. Natural stimulants are often included under the term biostimulants, including phenols, salicylic acid, humic and fulvic acids, or protein hydrolases [10,11]. An important group of plant biostimulants are organisms including fungi and bacteria that change the species composition of organisms found in the soil or plants. Their presence may accelerate the rate of degradation processes or limit the number of specific fungal and bacterial groups [12,13]. Popular fungi used as biostimulants include *Glomus intraradices* [14], *Trichoderma atroviride* [14], *Trichoderma reesei*, and *Heteroconium chaetospira* [10,15–19]. Useful bacteria include *Arthrobacter* spp., *Enterobacter* spp., *Acinetobacter* spp., *Pseudomonas* spp., *Ochrobactrum* spp., *Bacillus* spp., and *Rhodococcus* spp. [18,19].

Biostimulants cannot be defined as fertilizers because they do not provide nutrients directly to plants. Biostimulants may facilitate the acquisition of nutrients by supporting metabolic processes in the soil and plants. An example of such an activity is the facilitation of the development of arbuscular mycorrhizal fungi that transport nutrients to the host plant [20].

#### **3. Sources of Biostimulants**

Biostimulants are preparations made from natural raw materials. Some of them are plant extracts such as rosemary, which stimulates the growth of tomato plants with a concentration of 1000 ppm. Rosemary oil contributes to improved nutrient uptake and increases the fresh mass of roots [21]. Plant and animal biostimulants are formed, for example, as a result of chemical or enzymatic hydrolysis. The products of hydrolysis are mixtures of peptides and amino acids (protein hydrolysates). Chemical acid or alkaline hydrolysis is used to produce biostimulants of animal origin, from raw materials such as hen feathers, bone meal, casein, collagen from skins, animal tissue or fish waste (Table 1). Biostimulants of plant origin are produced using enzymatic hydrolysis. In the production of plant biostimulants, for example, alfalfa hay, pulses, and vegetable or fruit waste may be used [13,22]. Protein hydrolysates contain amino acids, peptides, and non-protein compounds. Protein hydrolysates stimulate plant growth, reduce the use of inert fertilizers, and are environmentally friendly [22]. The solution, which at the same time allows for a reduction in the amount of organic waste and the creation of biostimulating preparations, is actually a fermentation process. Biostimulants may also be the products of anaerobic digestion (Table 1). Dissolved organic matter is formed in fermentation chambers and has stimulating properties. The source of dissolved organic matter is usually plant, animal, and lignin biomass [23]. Biopreparations from marine algae may contain low-molecular polypeptides and amino

acids, vitamins, enzymes, phytohormones, sugars, and antioxidants. These compounds activate the processes of rhizogenesis and lead to positive morphological and anatomical changes in plants (Table 1). Two applications of a biopreparation derived from algae enhanced the development of *Cornus alba* "Aurea" roots by 80% when compared to the control. Such biopreparations can be used for rooting young seedlings or improving the process of adult rhizogenesis [24].

Literature data indicate the positive effect of seaweed extracts as plant biostimulants. The extracts from *Ascophyllum nodosum* are listed as the most frequently used [25,26]. Equally popular are *Solanum lycopersicum* L. [27], *Ecklonia maxima*, *Sargassum* spp. [12], *Laminaria* spp., *Durvillaea potatumum*, *Ulva lactuca*, *Caulerpa sertularioides*, *Padina gymnospora*, *Sargassum liebmannii*, and *Sargassum johnstonii* [25,28].

The group of biostimulants also includes consortia of beneficial fungi or bacteria. Among the fungi used in the cultivation of plants, the following species are noted: *Glomus intraradices*, *Trichoderma atroviride*, *Trichoderma reesei*, and *Heteroconium chaetospira* (Table 1) [10,14–17]. Symbiotic arbuscular mycorrhizal fungi have a positive effect on crop quality. Arbuscular mycorrhizal fungi, along with *Rhizoglomus irregular*, promote the growth of *Stevia rebaudiana* Bertoni [20]. Plant growth bacteria include *Arthrobacter* spp., *Enterobacter* spp., *Acinetobacter* spp., *Pseudomonas* spp., *Ochrobactrum* spp., *Bacillus* spp., and *Rhodococcus* spp. (Table 1) [18,19]. The largest group of beneficial bacteria includes *Rhizobium* spp. and plant growth-promoting rhizobacteria [10]. The plant growth-promoting rhizobacteria group includes *Streptomyces* spp., *Pseudomonas* spp., and *Bacillus* spp. The literature results indicate that *Streptomyces* spp. protect tomato plants against putrefactive bacteria *Pectobacterium carotovorum* subsp. brasiliensis (Pcb). In addition, the volatiles produced by six *Streptomyces* spp. isolates stimulate the growth of tomato roots. The largest fresh root mass obtained due to the volatile substances was 138.2 ± 16.1 mg in comparison to a control mass of 111.5 ± 10.3 mg. The volatile substances also positively affect the dry matter content and root volume as well as the dry and fresh mass of the shoot [29]. Zhao et al. [30] managed to isolate 276 endophytic bacteria from soybean root nodules that protected soybean roots against fungal infections of *Phytophthora sojae*. The antagonistic bacteria included *Enterobacter* spp., *Acinetobacter* spp., *Pseudomonas* spp., *Ochrobactrum* spp., *and Bacillus* spp. Also, lactic acid bacteria, e.g., *Lactobacillus plantarum*, and *Lactobacillus paracasei* are used to promote plant growth and indirectly control diseases [31].


**Table 1.** Activity of different types of biostimulants.

#### **4. Application Method of Biostimulants**

Biostimulants may be used in the form of soil preparations (powders, granules, or solutions added to the soil) or as liquid foliar application products [41]. Biostimulants containing humic substances and nitrogen compounds are often applied directly onto the soil, whereas various types of extracts from plants and seaweed are used in the form of foliar applications. Biostimulants can be introduced into the irrigation system and taken up by plants along with water. One example is the Kelpak SL (*Ecklonia maxima* extract) biostimulant which was sprayed in an aqueous solution of *Phaseolus vulgaris* L. [41]. Biostimulants are used regularly during the whole vegetative period or proactively, i.e., once during the decline of vital forces of the plant. In this case, the biostimulants were administered once during the occurrence of a strong stress factor, e.g., frost [42–44]. The results show that the soil application of the biostimulant was not as effective as foliar application. The foliar application of a biostimulant obtained from sewage sludge increased the level of macro- and micronutrients in the leaves of maize. The nitrogen content in maize leaves increased by 26% (dose 3.6 L/ha) and 46% (dose 7.2 L/ha) [45]. Biostimulants can also be used in the form of biomass or meal from seaweed, however, this method has some limitations. Biomass and meal may be used in areas located close to the source of seaweed acquisition due to transport problems. Biomass or meal is applied directly to the soil long before planting in order to enrich the substrate with nutrients. Agro-technical measures such as ploughing are used to mix biomass or meal with the topsoil [12].

Common forms in which biostimulants occur are ready-to-use extracts or powder to make an aqueous solution. Soil biostimulants often affect the structure of the root, increasing, among other factors, its ability to absorb nutrients. Foliar extracts protect the plant against biotic and abiotic stresses. The circadian rhythm of plants should be taken into consideration. Biostimulants should be applied in the morning when the stomata are open and the assimilation rate is at its peak [12,38]. Biostimulants are also applied directly onto harvested fruits. It was noted that biostimulants containing a combination of extracts from *Sargassum* spp., *Laminaria* spp., and *A. nodosum* (Table 1) significantly extended the shelf-life and the storage life of oranges. After using the biostimulant, the fruits became more resistant to mechanical damage and putrefaction which allowed for an extension of the storage time and suitability for consumption. The extract used produced a better effect than, for example, the calcium chloride normally used in the industry to protect fruit against putrefactive bacteria [12,38].

#### **5. E**ff**ect of Biostimulants on Yielding**

The popularity of biostimulants in agriculture is associated with the possibility of obtaining higher yields without the need to discontinue the production of ecological crops. According to numerous scientific studies, biostimulants have a positive effect on yielding plants [36]. The yield is usually determined as the amount of fruit obtained from one plant or plot. The yield depends on the type of biostimulant used, the dose, the method of application, and the plant variety. Increased yield is often associated with improving the quality of vegetables or fruit. This is particularly important in organic farming, where artificial fertilizers cannot be used [37,45]. The quality of fruits and vegetables is shaped from the moment of plant growth to the time of harvesting of fruits and vegetables and it consists of the taste and the content of nutrients. The quality is influenced by genetic and agro-environmental factors [46].

The positive influence of biostimulants based on humic, fulvic, and carboxylic acids on the yielding of apricot fruits has been proven [8]. Control trees showed a yield of 12 kg fruit/tree and after the application of humic and fulvic acids together and carboxylic acids in a separate experiment, the yield of the trees increased to 21 kg of fruit/tree and 19 kg of fruit/tree, respectively (Table 2). However, this relationship was observed only in the second year of using the biostimulant. During the first growing season, the yield of the control trees was higher than that of the trees that were treated with biostimulants containing humic and fulvic acids. The biostimulant based on polysaccharides turned out to be ineffective with this variety of apricot trees, which showed a yield comparable to the control in both growing seasons [8]. Preparations containing all amino acids allowed for an increase in mango

yield. With a dose of 3 L/ha, the yield increased by 18% compared to the control. At the same time, the biostimulants caused a 15% decrease in fruit weight. The authors explained this phenomenon through the competition of fruit for nutrients [32]. Biostimulants containing phenolic compounds such as sodium para-nitrophenolan, sodium orto-nitrophenolan, and sodium 5-nitroguajakolan proved to be good preparations for raspberry bushes. As a result of the foliar application of the phenolic compounds, a 20% (Table 2) increase in raspberry yield was obtained [47]. The highest yield was achieved when a biostimulant containing phenol compounds was applied to the "Polka" raspberry variety (yield = 23.03 kg/plot), while the yield in the control was 18.28 kg/plot. Already in the first year after the use of biostimulants, the amount of fruit collected from one bush increased. The best results were produced by 6-benzyladenine in a dose of 100 mg/L, which caused an increase in yield in the first year by about 0.5 kg. In the second year of using the biostimulant 6-benzyladenine at a dose of 100 mg/L and α-naphthaleneacetic acid at a dose of 20 mg/L, the yield increased by more than 1 kg from a single shrub [37]. Strawberry yielding significantly increased after using biostimulants containing herbal and marine plant extracts. In this case, the study proved that soil biostimulants are a source of nitrogen compounds. Moreover, foliar biostimulant application did not produce results as good as biostimulants added to the soil. Biostimulants added to the soil caused a significant increase in the amount of fruit and also improved the condition of the plants. Plants were more resistant to weather conditions and pathogens [42]. Extracts of "Moscatel" vine-shoots improved the yielding of the grapevine variety "Airén". The literature results indicate that two foliar biostimulant variants were prepared, non-toasted and toasted vine-shoots. A significantly higher yield was achieved in the case of two preparations, which were the non-toasted vine-shoots extract (3.09 ± 0.05 kg/plant) and the toasted vine-shoots extract (3.57 ± 0.05 kg/plant) in comparison to the control (2.54 ± 0.03 kg/plant) (Table 2) [48].

A mixture of four biostimulants containing amino acids, polysaccharides, vitamins, humic acids, organic carbon, and enzymatic proteins caused a comparable increase in the yield of two varieties of yellow pepper [49]. The Blondy F1 variety produced a crop at a level of 5.98a 0.23 kg/plot (yield = 5.24 ± 0.30 kg/plot). The Century F1 variety produced a yield of 5.76 ± 0.20 kg/plot (yield in the control = 5.06 ± 0.28 kg/plot) (Table 2). The peptides and amino acids contained in these formulations demonstrated a protective action against excessively high temperatures in the summer season and induced root growth and development, while vitamins and humic acids resulted in fruit growth [49]. Horseradish extract increased pumpkin yield by 12.5% [50] and the beneficial effect of fungal species *Glomus intraradices* and *Trichoderma atroviride* positively influenced the yielding of zucchini, resulting in a yield increase of 0.39 kg per plant on average, probably by increasing the effectiveness of nutrients uptake [14]. A 3% *Moringa oleifera* extract in combination with 0.6% ZnSO4 and 0.25% K2SO4 increased the yield of "Kinnow" mandarin plants by 65% (Table 2) compared to the control [51].

The results of the study underline the positive effect of humic acids on the yielding of fruit trees [8]. The use of phenolic compounds [47] resulted in the increased yielding of fruit bushes. A high yield of vegetables may be obtained by using a mixture of amino acids, polysaccharides, vitamins, humic acids, and other compounds. Each of the substrates of the mixture affects another biochemical process occurring in the soil and plant, which allows for the achievement of the desired effect [49].


Effect of selected biostimulants on quality of fruit and vegetables.

> **Table 2.**


**Table 2.** *Cont.*

#### **6. E**ff**ect of Biostimulants on the Growth and Size of Plants**

The way in which biostimulants work may be defined as multifaceted. The literature describes the positive effect of biostimulants on the growth of fruits and vegetables. At the same time, there are studies in which no effect of biostimulants on fruit size was found. The lack of biostimulant effects is explained by the use of a biostimulant unsuitable for the tested cultivar [37,53].

Fruit producers are interested in biopreparations that allow for the attainment of the largest and healthiest-looking fruit that draw consumers' attention [52]. An increase in the average length and diameter of cucumbers was attained after the use of humic acids and a mixture of nitrogen, amino acids, and auxins. Humic acids in a concentration of 3g/L increased the average length of the fruit in the first and second season by 9.9 cm and 12.2 cm, respectively. The same concentration of humic acids increased the diameter of the cucumbers in the first and second season by an average of 1.23 cm and 1.55 cm, respectively [52]. Three tested biostimulators caused an elongation and increase in the diameter of the vegetables. One biostimulant (containing nitrogen, amino acids, and auxins) led to the elongation of the cucumbers by 3.85 cm in the first and 3.49 cm in the second growing season. The diameter of the fruit increased by 1.12 cm and 1.56 cm, respectively, in the first and second growing season under the influence of this biostimulant. The application of humic acids and biostimulants containing auxins in particular makes it possible to obtain elongated and thickened cucumbers [52]. The use of biostimulants containing humic and fulvic acids as well as carboxylic acids led to a tenfold enlargement of the apricot fruit. The greatest influence on the size of the fruit was carboxylic acids, which contributed to the widening of fruit by 2.6 mm on average in the second growing season [8].

Studies show that the perfect biostimulants that cause the growth of fruits and vegetables are consortia of microorganisms. Examples are arbuscular mycorrhizal fungi and plant growth-promoting bacteria, the use of which resulted in increased tomato weight. A positive effect was obtained through the combination of arbuscular mycorrhizal fungi containing fungi of the following species: *Rhizophagus* spp., *Rhizophagus aggregatus*, *Septoglycus viscosum*, *Claroideoglomus etunicatum*, *Claroideoglomus claroideum* and various types of plant growth-promoting bacteria. All of the biostimulants based on microorganisms (arbuscular mycorrhizal fungi + *Pseudomonas* sp. Strain 19Fv1T, arbuscular mycorrhizal fungi + *Pseudomonas fluorescens* C7, arbuscular mycorrhizal fungi + *Pseudomonas* sp. 19 Fv1T and *Pseudomonas fluorescens* C7) caused an increase in tomato fruit mass, but the most effective result was demonstrated by biostimulants including arbuscular mycorrhizal fungi and *P. fluorescent* C7, which caused an increase in tomato mass to 71.3 ± 0.6 g (weight of control tomatoes = 64.4 ± 0.9 g). The microorganisms used caused a slight elongation of tomatoes as well. The length of the fruit in the controls ranged from 5.49 ± 0.03 cm to 5.81 ± 0.03 cm, while in combination with the biostimulants used, it rose to 5.88 ± 0.03 cm to 6.05 ± 0.02 cm. There was also a slight increase in fruit diameter from 4.24 ± 0.02 cm to 4.62 ± 0.03 cm to 4.64 ± 0.03 cm to 4.78 ± 0.02 cm [53].

The application of 6-benzyladenine as a biostimulant at a dose of 100 mg/L resulted in an increase in the weight of blueberry fruit by about 32.4% (first season) and 33.6% (second season) for the blueberry cultivar Duke and 43.5% (first season) and 33.1% (second season) for the blueberry cultivar Bluecrop compared to the control. The literature results showed that α-naphthaleneacetic acid at a dose of 20mg/L was also an effective biostimulant, which increased the weight of the blueberry cultivar Duke fruit by 41.9% (first season) and 20.0% (second season) and the blueberry cultivar Bluecrop by 55.0% (first season) and 25.4% (second season). As demonstrated in the study, one inefficient biostimulant was gibberellic acid at a dose of 200 mg/L. Gibberellic acid increased the weight of blueberry cultivar Duke fruit by 4.7% (first season) and 14.3% (second season) as well as Bluecrop blueberry cultivar by 0.8% (first season) and 11.5% (second season) compared to the control. For this reason, gibberellic acid is not recommended as a biostimulant for soft fruits [37].

The titanium compounds with which raspberries were treated caused an increase in fruit weight from 4.44 g (control) to an average of 5.4 g, but only at the beginning of the harvest season. At the end of the harvest, a 57% decrease in raspberry weight was observed [47]. The same relationship was observed for phenolic compounds. At the beginning of the harvest, the mass of raspberries treated with phenolic compounds was 5.04 g on average, while at the end of the harvest there was a 44.4% loss in fruit weight. The fact is that the raspberry collection is characterized by the loss of fruit mass at the end of the harvest, but if these three biostimulants were used, the resulting losses were greater than in the control. The control fruits were characterized by a 42.3% weight loss, while the fruits treated with biostimulants showed a mass loss at the end of the harvest in the range from 44.4% to 57.0% [47]. The increase in fruit weight after the use of biostimulants compounds was also observed in the case of cherries. The use of salicylic acid with the addition of calcium resulted in a 15% increase in sour cherries "Sweetheart" (2015) and "Skeena" (2016) [57].

Regarding the multifaceted effect of biostimulants, it should be emphasized that these formulations may affect many of the characteristics of the plant, e.g., fruit size, plant height, and root length [52]. The use of humic acids in a concentration of 3g/L and other biostimulants caused an increase in cucumber plant height, the number of leaves, and the number of stems in both growing seasons [52]. In the first growing season, the height of the control plants was 78.13 cm. Plants treated with 3 g/L of humic acids were 14.25 cm taller on average. In the second growing season plants treated with humic acids were taller by 13.25 cm. Similar dependencies may be observed after using other biostimulators. In the first and second growing season, this biostimulant (including, among others, nitrogen, amino acids, and auxins) was the most effective, causing an increase in plant height of 14.5 cm (in the first growing season) and 19.75 cm (in the second growing season) in comparison with the control plants. The biostimulant including, among others, naphthyl acetic acid was the least effective, and resulted in an increase in the growth of plants by 4.38 cm and 7.92 cm, respectively to the first and second growing seasons. Similar dependencies were noted in the number of leaves and new stems [52].

#### **7. Impact of Biostimulants on Physical Characteristics**

Biostimulants also have an influence on mechanical properties, i.e., the firmness of fruits or vegetables. Depending on the type, biostimulants may cause the stiffening of cell walls, thereby reducing their extensibility [8]. Biostimulants that increase the flexibility of cell walls at the same time extend the shelf-life of fruits and vegetables for consumption and facilitate their storage. Biostimulants based on carboxylic, humic, and fulvic acids and also the biopolymers of polysaccharides increased the mechanical strength of apricot fruits during two years of biostimulant use [8]. In turn, biostimulants containing phenolic compounds or chitosan resulted in the loss of fruit firmness of the three raspberry varieties studied. The use of biostimulants based on titanium compounds did not alter the fruit firmness, which was comparable to the firmness in the control test [47]. The use of spic cytozyme containing essential plant nutrients and growth biostimulants in the amount of 4 mL/L significantly reduced the cracking of pomegranate fruit [58]. In addition to improving mechanical properties, biostimulants change the shape and colour of fruits and vegetables. Fruit with larger length and diameter, as well as the right colours, are preferred by consumers [8]. However, consumer preferences are subject to dynamic changes.

An important visual feature that proves the quality of fruit is colour. The colour of the fruit is substantially influenced by the content of anthocyanins. Weber et al. [59] examined the content of anthocyanins in strawberries treated with *Ascophyllum nodosum* extract with silicon. Fruits treated with a biostimulator were characterized by a higher content of anthocyanins in the initial fruiting period, therefore, they were more red than the control fruits [59]. "Sweetheart" cherries treated with glycine and betaine were characterized by a darker skin than the control fruits. Although the mechanism of action of betaine and glycine on the formation of anthocyanins is not fully understood, it is known that the darker colour of the fruit was caused by a higher content of antioxidants [57]. Tarantino et al. (2018) in the second year of using biostimulants obtained apricot fruits with a lighter skin compared to the first year. This could be due to the higher concentration of biostimulants used in the first growing season. There were significant differences in the colour of the fruit. In the second year of the experiment the colour of the apricots was redder than in the first year of fruiting. There were no significant differences

in the colour of the fruit produced by the three biostimulants used (1—biopolymers of polysaccharides; 2—humic and fulvic acids; 3—carboxylic acids) [8].

It should be emphasized that the increased mechanical strength or fruit colour change results from the good condition of fruit plants, which in turn is a result of their proper nutrition [50]. Biostimulants are supplied externally, which indirectly, e.g., induced by the photosynthesis process, plays an important role in the nutrition of plants. Indirect induction, for example, consists in increasing the leaf area. Leaves are the main organs in which photosynthesis takes place, therefore increasing the leaf area leads to an increase in photosynthesis. Intensively photosynthetic plants are better nourished. Increasing the leaf area also leads to an increase in the transpiration surface. This phenomenon has the especially important function of protecting the plant from overheating. *Moringa oleifera* leaf extract increased the surface area of the *Cucurbita pepo* L. by 9.7% and simultaneously led to a 34.6% increase in the chlorophyll content of the leaves compared to the control [50]. The positive effect of the *A. nodosum* extract in a period of drought on the growth of spinach has been proven. Seaweed extract increased the relative water content of the leaves from 76% to 82%. The surface area of the leaf was also increased by 16% (foliar spray), 21% (biostimulant in the irrigation system), and 38% (biostimulant in the irrigation system and in a spray). Increasing the area and turgor of the leaf led to an increase in the intensity of photosynthesis and improved the conditions for growing spinach under stress conditions [25].

#### **8. E**ff**ect of Biostimulants on Chemical Composition**

Biostimulants can affect a number of the chemical properties of fruits and vegetables, including dry mass, acidity or vitamin content. The chemical composition of the fruit directly affects their palatability. It is assumed that fruits with a content of dissolved solids (SSC) above 12◦Brix are characterized by an excellent taste [47]. In the first year of using biostimulants containing the biopolymers of polysaccharides, humic and fulvic acids as well as carboxylic acids, the average value of SSC in apricots stood at 10.7◦Brix. In the second year of using these biostimulants, fruit taste values improved significantly, as evidenced by the increase in the SSC level to an average of 14.1◦Brix [8]. Biostimulants containing phenol compounds or chitosan reduced the dissolved solids content in the fruits of the three raspberry varieties (Pokusa, Polka, and Poranna Rosa). The opposite effect was produced by biostimulants based on titanium compounds, the use of which resulted in an increase in the content of dissolved solids in the raspberry fruit [47]. The quality of the fruit is also demonstrated by the ratio content of dissolved solids to their titratable acidity. Fruit quality is defined as good if the ratio content of dissolved solids to titratable acidity is within the range of 10 to 15. The treatment of fruit trees with biostimulants containing biopolymers of polysaccharides (16.7) and humic and fulvic acids (16.1) leads to an increase in the ratio content of dissolved solids to titratable acidity in relation to the control (14.0) and thus negatively influenced the sensory quality of fruit [8].

It is important to grow fruit that has an appropriate level of acidity but it is difficult to say, however, whether changes in acidity at the level of several percent have a significant impact on the fruit taste, because it is based on the subjective impression of the consumer. Although the literature data present studies on the effect of biostimulants on fruit acidity, there is no explicit interpretation of the results. It is not clear whether the changes in fruit acidity should be understood in terms of the positive or negative effects of the biostimulants used. The use of phenolic compounds and titanium compounds as a biostimulant in the cultivation of raspberries led to an increase in fruit acidity to 2.26% and 2.18%, respectively (control, 2.08%) [47]. A decrease in apricot acidity was noted after the use of biostimulants containing polysaccharides, humic and fulvic acids, and carboxylic acids [8]. In the second year after using these biostimulants, fruit acidity was reduced from an average of 3.45 (control pH) to an average of 3.7–3.8 (pH after using biostimulants) [8].

An important health-related feature of fruits and vegetables is the content of vitamin C and nitrogen compounds. The role of nitrogen in plants results from its influence on growth and development. It is a component of nucleic acids, it participates in the process of photosynthesis, and it builds amino acids that form a part of plant proteins [60]. The content of vitamin C and nitrogen compounds in the fruit depends mainly on the plant variety [47]. The use of selected biostimulants (phenolic compounds, chitosan, and titanium compounds) increased both the levels of ascorbic acid and nitrates. It turned out that the three biostimulators tested positively influenced the level of nitrates in raspberries. It was found that phenolic compounds contained in one of the biostimulants increased the level of vitamin C most effectively [47]. Assuming that phenolic compounds increased the content of vitamin C in fruit, it was necessary to determine how different biostimulators affect the content of phenolic compounds. Zarzecka et al. [61] studied the effect of herbicides and biostimulators on the polyphenol content of potato tubers. The experiment was conducted over a period of two years. Three potato varieties were treated with different substances: Harrier herbicide 295 ZC, Harrier 295 ZC + Kelpak SL growth regulator, and Sencor 70 WG herbicide, 5 Sencor 70 WG + Asahi growth regulator. In the case of Asahi, the active substances were phenolic compounds, while for Kelpak SL, auxins and cytokinins were the active substances. The applied biostimulants and herbicides caused an increase in the polyphenol content in tubers of all potato varieties (on average, 159.8–161.3 mg/kg) compared to the control (average of 156.0 mg/kg). The use of biostimulants and herbicides increased the content of polyphenols in the leaves of the potato to an average of 289.2–291.2 mg/kg compared to the control (287.8 mg/kg). The content of polyphenols in tubers is of particular importance for humans. Polyphenols reduce the risk of numerous diseases, e.g., blocking carcinogenic compounds [61]. It was observed that after using a biostimulant containing seaweed *A. nodosum* and silicon, the content of phenolic compounds in strawberries was slightly lower. Phenols are also defined as compounds produced by plants under stressful conditions, hence the conclusion about the positive effect of the tested biostimulant on strawberries [59]. The foliar spraying of the "Airén" grapevines by non-toasted and toasted biostimulants increased the content of phenolic compounds. In this case, the biostimulants were extracts from the "Moscatel" vine shoots. An important group of phenolic grape buds are hydroxycinnamic acids (trans-caffeic and trans-p-coumaric), which affect the taste of wine. Both non-toasted biostimulants (14.10 ± 0.13) and toasted biostimulants (11.26 ± 0.27) led to a trans-p-coumaric acid increase relative to the control (8.60 ± 0.03). The non-toasted biostimulant (1.14 ± 0.01) and toasted biostimulant (0.95 ± 0.03) also led to a trans-caffeic acid growth compared to the control (0.92 ± 0.03). The effect of the higher content of, among other compounds, hydroxycinnamic acid is a better quality wine [48]. A biostimulant containing *A. nodosum* seaweed extract increased the phenol content of "Sangiovese" grapes. A 1.5 kg/ha dose of biostimulant increased the phenolic content to 1.063 mg/cm2, while the biostimulant in a dose of 3 kg/ha increased the phenolic content to 0.951 mg/cm2. The phenol content of the control was 0.753 mg/cm2. The results were statistically significant [55].

The positive effect of biostimulants is also based on increasing the content of chlorophyll in leaves and thus increasing the efficiency of the process of photosynthesis. Salicylic acid-chitosan nanoparticles used as a biostimulant led to an increase in the content of chlorophyll in leek corn [54]. While the chlorophyll content in the control was 10.72 mg/g, the chlorophyll content in the maize leaves treated with the biostimulant (concentration 0.01%–0.16%) was in the range of 16.43 to 25.88 mg/g on average. In plants treated only with chitosan and salicylic acid, a decrease in the chlorophyll content to an average of 9.24 mg/g and 9.79 mg/g was observed [54]. After the foliar application of the *Moringa oleifera* leaf extract, a 34.6% increase in the chlorophyll content of *Cucurbita pepo* L. leaves was recorded compared to the control (plants sprayed only with water) [50]. The increase in the chlorophyll content combined with the increased intensity of the photosynthesis process was noted during the cultivation of Hibiscus treated with biostimulants formed in the process of the hydrolysis of waste. A 15% increase in the chlorophyll content of leaves resulted in a 24% increase in the photosynthesis rate compared to the control [62].

During the tests to determine the chemical composition of fruits, the content of glucose, fructose, sucrose, ascorbate, proteins, and macro- and micro-elements is often determined. Plants treated with one of the tested biostimulators (arbuscular mycorrhizal fungi + *Pseudomonas* sp. 19 Fv1T and *P. fluorescens* C7) showed an increase in the concentration of glucose in tomatoes at 11.83 g/kg, while in controls the content of glucose was 10.45–11.0 g/kg. After using this biostimulant, the fructose

content also increased to about 12.86 g/kg, while in the controls it was 10.77–11.14 g/kg. After using a biostimulator based on *A. nodosum* seaweed extract and a silicon extract, a slight increase in the level of sugars in the strawberry fruit was observed. The most common sugars were glucose and fructose. Sucrose accounted for 11% of total sugars [59]. The use of an extract of *M. oleifera* leaves increased the total soluble sugar content in pumpkin by about 80.6% [50]. An interesting relationship was observed in the case of ascorbate. The use of biostimulant containing arbuscular mycorrhizal fungi, *Pseudomonas* sp. 19 Fv1T and *P. fluorescens* C7 led to an increase in the ascorbate content (10.75 mg/100 g), whereas the use of a biostimulant containing arbuscular mycorrhizal fungi and *P. fluorescens* C7 reduced the ascorbate content in tomatoes (4.30 mg/100 g). In the controls, tomatoes contained about 5.47–7.12 mg/100 g ascorbate. In plants treated with the biostimulant containing arbuscular mycorrhizal fungi and *P. fluorescens* C7, an increase in β-carotene in tomatoes was observed (controls: 2.117–2.224 μg/100 g fresh weight; β-carotene content in the biostimulator study: 2.829 μg/100 g fresh weight). β-carotene may be converted into vitamin A and can protect against the adverse effects of free radicals [53]. The protein content is particularly important in the case of grain plants. The foliarly used biostimulator containing sewage sludge caused an increase in protein content in maize grains by about 30% in both growing seasons [45]. The biostimulator formed as a by-product of the two-stage process of pressing olive oil led to an increase in the protein content of maize grains by 19% [63].

Combining the biostimulant (chicken feathers) with a fertilizer gave a better quality of maize yield than using only the fertilizer. Three combinations of the agent used for spraying the corn were used. In the first variant, only fertilizer was used (300 kg N/ha + 120 kg K/ha). For the second variant, fertilizer was used (300 kg N/ha + 120 kg K/ha) in combination with a biostimulant (3.6 L/ha). In the third variant, fertilizer (300 kg N/ha + 120 kg K/ha) was used in combination with a biostimulant (7.2 L/ha). The treatment was applied during two seasons. The highest level of nitrogen was obtained after the application of a biostimulant containing fertilizer in combination with the highest dose of the biostimulant (7.2 L/ha). The nitrogen content of corn leaves increased by 14.4% (fertilizer with the biostimulant of 3.6 L/ha) and 39.1% (fertilizer with the biostimulant of 7.2 L/ha) in the first vegetative season in comparison to the nitrogen content of maize leaves treated only with fertilizer. In the second growing season, the nitrogen content increased by 15% (fertilizer with the biostimulant of 3.6 L/ha) and 33.3% (fertilizer with the biostimulant of 7.2 L/ha) respectively. The use of a biostimulant in combination with the fertilizer also resulted in an increase in phosphorus content in the leaves of maize. The treatment increased *p* levels by 32.8% (fertilizer with a biostimulant of 3.6 L/ha) and 52.2% (fertilizer with a biostimulant 7.2 L/ha) in the first season and by 43.5% (fertilizer with a biostimulant of 3.6 L/ha) and 51.1% (fertilizer with a biostimulant of 7.2 L/ha) in the second season compared to the control [33]. An effective biostimulant for the "Kinnow" mandarin trees proved to be *Moringa oleifera* extract. The 3% *Moringa oleifera* extract foliar application with 0.6% ZnSO4 and 0.25% K2SO4 resulted in a 1.35-fold (first season) and 1.42-fold (second season) increase in nitrogen content compared to the control. Trees sprayed only with 3% *Moringa oleifera* extract showed a 1.09 times (first season) and 1.07 times (second season) higher phosphorus content compared to control trees [51].

The improvement in the chemical properties of fruits may increase not only their pro-health values, but also lead to an improvement in their sensory values. One example may be guaiacol, which was applied in a foliar way to improve the quality of wine. Guaiacol was shown to increase the amount of glycosylated aromatic compounds in "Microvine" grapes. These compounds have influenced the improvement of wine quality in the final step of wine formation. The guaiacol-treated fruits were characterized by a higher aglycone content (534.25 μg/g) compared to the control (157.52 μg/g). Treatment with guaiacol also increased the content of monomethyl alcohols from 2.94 μg/g in control fruits to 170.30 μg/g in guaiacol-treated fruits [64].

Biostimulants are becoming a viable option for solving the problem of the ineffective uptake of nutrients from fertilizers by plants. The fact is that a large proportion of fertilizer nutrients are not taken up by plants. Reducing the amount of mineral fertilizers introduced into the soil limits environmental degradation. It is thought that the development of certain biostimulants has the potential to increase the amount of nutrients taken up by plants [65]. The increase in the amount of nutrients taken up by plants may be achieved through the use of fertilizers and biostimulants in combination [33]. An increased nitrogen content was obtained thanks to the use of a biostimulant formed in the process of the hydrolysis of chicken feathers in combination with nitrogen fertilizer, while the increase in phosphorus content was the result of using a biostimulant formed in the process of the hydrolysis of chicken feathers with phosphate fertilizer [33] and an extract of *Moringa oleifera* as a biostimulant [51].

#### **9. E**ff**ect of Biostimulants on Antioxidant Properties**

Antioxidant activity is an often-studied property of fruits and vegetables. Antioxidants are listed as compounds that inhibit tumour cell proliferation and protect against oxidative stress caused by excess free radicals. The result of oxidative stress may be, among other factors, damage to DNA, cell membranes, or enzymes [66]. It was shown that the use of biostimulants in plant breeding can change the activity of enzymes and affect the antioxidant properties. Lycopene, ascorbic acid, phenolic compounds and others have antioxidant properties. Reactive oxygen molecules, e.g., OH, O2<sup>−</sup>, and H2O2, are detoxified by antioxidant compounds (e.g., phenols, ascorbic acid) and enzymes (e.g., catalase, peroxidase, superoxide dismutase) [67].

Protein hydrolysate applied as a biostimulant to tomatoes had no effect on the level of phenolic compounds, while its effect on the content of ascorbic acid and lycopene was noted. After using biostimulant doses of 5.0 and 2.5 mL/L, the content of lycopene increased by 34.9% and 18.0%, respectively, compared to the control. The dose of 2.5 mL/L biostimulant increased the content of ascorbic acid by 27.3% [68]. The use of biostimulants on apricot fruit trees increased the antioxidant capacity of fruits. In the first season (average 76.8 mg/100 g), after using the stimulants, the antioxidant capacity of fruit was higher than in the second season (average 66.5 mg/100 g). The observed differences in antioxidative abilities between the two seasons were explained by changes in climatic conditions [8].

Also, a biostimulant based on salicylic acid and chitosan nanoparticles (SA-CS NPs) had an effect on the enzyme and antioxidant activity in maize leaves. The enzyme activity in leaves treated with chitosan, salicylic acid, and a control was comparable. After two days of treating the plants with the biostimulant, the activity of superoxide dismutase increased by two times compared to plants treated with salicylic acid. After three days of treating plants with a biostimulant, superoxide dismutase activity was 3.2 times higher than for plants treated with only salicylic acid. Peroxidase activity in plants treated with a biostimulant was 7.7 (after two days) and 5.2 (after three days) times higher than for plants treated with only salicylic acid. Catalase activity, phenylalanine ammonia lyase, and polyphenol oxidase increased by 2.9, 2.3, and 1.5-fold, respectively, after the second day of treatment with nanoparticles compared to salicylic acid treatment. It should be emphasized that the enzyme activity occurring in the leaves of plants treated with a biostimulant increased during the first three days. After the fourth day of treatment, the enzyme activity decreased in all variants of the experiment. The content of hydrogen peroxide in leaves treated with SA-CS NPs was 1.7 (first day of treatment), 3.6 (second day of treatment), and 1.7 (third day of treatment) times higher than in plants treated with salicylic acid [54].

The use of *M. oleifera* extract as a biostimulant resulted in a decrease in the activity of the antioxidant enzymes (catalase, peroxidase, and superoxide dismutase) in rocket plants (*Eruca vesicaria* subsp. *Sativa*). At the same time, the content of phenol and ascorbic acid was higher with increasing concentrations of the biostimulant [67]. Aqueous garlic extract improved tomato oxidation properties. Superoxide dismutase activity increased in proportion to the aqueous garlic extract concentration. The highest activity of this enzyme was observed with the foliar application of the biostimulant in a volume of 200 μg/L; also, the peroxidase activity was highest after using the biostimulant at this concentration. A lower aqueous garlic extract concentration (50 μg/L) did not affect the activity of these enzymes [69]. Soaking sunflower seed *Helianthus annuus* L. in a 3% corn seed extract and spraying 1 mM Mg plants stimulated the sunflower s antioxidant system. The enzymatic activity of superoxide dismutase, catalase, and peroxidase increased by 65.5%, 77.8%, and 84.6%, respectively, as compared

to the controls. The increased level of antioxidant enzymes was related to the foliar application of Mg ions, the use of which also increased the intensity of the photosynthesis process [56].

Biostimulants increased the phenylalanine ammonia lyase enzyme activity. While the phenylalanine ammonia lyase level in the control was 7.9 ± 0.22 IU/mL × min (0.4% *E. maxima extract*), after using biostimulants it increased to 9.0 ± 0.01 and 9.7 ± 0.01 IU/mL × min (10−6 M *E. maxima* extract). Phenylalanine ammonia lyase is an enzyme catalyzing the first step in the synthesis of phenyl compounds. An increased production of phenolic compounds is observed during plant stress. From this, it may be concluded that biostimulants can induce plant stress to increase the production of secondary metabolites [70]. Biostimulatory properties also reveal many components of compost. Depending on the raw materials and methods used for the composting process, the compost may contain, among others, polysaccharides, amino acids, and organic nitrogen. Compost can be used to replace peat in greenhouse cultures. In addition, it may be produced from organic waste, such as wood, plant residue, or other residues. Compost which is considered by the European Union to be ecological must consist solely of natural raw materials, characterized by a limited content of heavy metals and hazardous elements (Se, Mo, S). The product of the composting process must be free of pathogenic agents (*Salmonella* sp. and *Escherichia coli*) [71]. Agroindustrial compost proved to be an alternative to peat in the cultivation of red lettuce. The compost increased the content of antioxidant compounds in lettuce leaves. In the autumn season, lettuce leaves cultivated in the compost showed 1.5 times more antioxidant activity than lettuce grown in compost in the summer season and it was also higher that of lettuce grown in peat in the autumn season [72].

#### **10. Conclusions**

Biostimulants are preparations of natural origin that support the pro-ecological cultivation of vegetables and fruits. Although for several years a positive effect of biostimulants has been widely reported, they are rarely introduced into standard cultivation technologies. This is connected with the insufficient knowledge of farmers on functions and usage of biostimulants what results in a fear of an increase in the cost of cultivation and a reduction in the quality and quantity of plants, which would affect the profitability of crops. The problem is also the multitude of preparations and the need to select a proper biostimulant for a specific plant variety in order to obtain the highest and the best quality yields. The market requires the development of preparations with a broad spectrum of functionality, which is easy to apply and has the possibility of combination with other agents.

The use of biostimulants on a commercial scale would limit the amount of mineral fertilizers introduced into the environment, thus reducing the pollution of soils, water, and air. This is especially important in the case of global warming. Global agriculture accounts for an average of 21% [73] of the global greenhouse effect, of which around 13% [74] is concerned with the effect of artificial fertilizers. The newly developed technologies of biopreparations may constitute a significant contribution to environmental protection, but primarily they are closely linked with sustainable agricultural and horticultural production with the aim of obtaining cheap, easily available, and high quality food. The effect of biostimulants depends on many factors, from the raw material and the process as a result of which they arose to the plant varieties, application method, and climate. The positive effect of consortia of microorganisms and plant hydrolysates on growth and yield of crops plants should be particularly emphasized. It is also important to increase the antioxidant potential of plants treated with biostimulants containing algae. A positive impact on crop quality and performance, no negative or harmful impact on people, animals, or the environment, increased biodiversity of beneficial microorganisms, and improvement of soil properties are the main advantages of biostimulants. However, the nature of their positive influence is not fully understood, therefore their mechanisms of action are, in some cases, still a challenge and need to be recognized. For this reason, biostimulants are among the hot topics in agriculture and still require detailed research.

**Author Contributions:** M.D. collected the data, interpreted the results and wrote the manuscript. M.F. provided ideas and performed reviews and corrections of the manuscript. J.C. designed the study, helped draft the manuscript, interpreted results and corrected the text. All authors read and approved the final manuscript.

**Funding:** The National Centre for Research and Development, Poland, project BIOSTRATEG, contract number BIOSTRATEG3/344433/16/NCBR/2018.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Paramylon Treatment Improves Quality Profile and Drought Resistance in** *Solanum lycopersicum* **L. cv. Micro-Tom**

#### **Laura Barsanti 1, Primo Coltelli <sup>2</sup> and Paolo Gualtieri 1,\***


**\*** Correspondence: paolo.gualtieri@pi.ibf.cnr.it; Tel.: +39-050-621-3026

#### Received: 15 May 2019; Accepted: 14 July 2019; Published: 17 July 2019

**Abstract:** Tomatoes, the most cultivated vegetables worldwide, require large amounts of water and are adversely affected by water stress. *Solanum lycopersicum* L., cv. Micro-Tom was used to assess the effects of β-(1,3)-glucan (paramylon) purified from the microalga *Euglena gracilis* on drought resistance and fruit quality profile. Plants were grown in an aeroponic system under three cultivation conditions: optimal water regimen, water scarcity regimen, and water scarcity regimen coupled with a root treatment with paramylon. Eco–physiological, physicochemical and quality parameters were monitored and compared throughout the lifecycle of the plants. Drought stress caused only a transient effect on the eco–physiological parameters of paramylon-treated plants, whereas physicochemical and biochemical parameters underwent significant variations. In particular, the fruits of paramylon-treated plants reached the first ripening stage two weeks before untreated plants grown under the optimal water regime, while the fruits of stressed untreated plants did not ripe beyond category II. Moreover, antioxidant compounds (carotenoids, phenolic acid, and vitamins) of fruits from treated plants underwent a two-fold increase with respect to untreated plants, as well as soluble carbohydrates (glucose, fructose, and sucrose). These results show that paramylon increases plant resistance to drought and highly improves the quality profile of the fruits with respect to untreated plants grown under drought stress.

**Keywords:** Biostimulants; *Euglena gracilis*; algal polysaccharide; β-glucan; water stress; tomato; aeroponics

#### **1. Introduction**

Paramylon is the storage product of the unicellular alga *Euglena gracilis.* This polysaccharide is a β-(1,3)-glucan endogenously synthetized as 1–2 μm granules consisting of 100% glucose [1]. The granules are composed of concentric segments, which possibly indicates the successive deposition of unbranched linear β-(1,3)-glucan chains on a central nucleus [2–4]. Wild type (WT) photosynthetic cells can accumulate paramylon up to 60% of cell dry weight (DW) [2], while the WZSL mutant of *E. gracilis* (spontaneous, non-chloroplastic, osmotrophic mutant; W describes the white color of the cells; Z means *E. gracilis* Klebs, Z strain; S means spontaneous mutant, and L means light grown parent culture) [5] can accumulate large amounts of it (up to 95% DW) when grown in the dark with an adequate carbon source [2].

B-glucans are PAMPs (pathogen-associated molecular patterns) recognized by specific membrane receptors (pattern recognition receptors, PRRs) which trigger the activation of the innate immune system [6,7]. Both the molecular structure and degree of polymerization affect the strength and efficacy of β-glucans recognition by PRRs, as well as their successive reactions [2,8–10].

Linear β-(1,3)-glucans bind preferentially to Dectin-1, a C-type lectin receptor expressed on most cells of the innate immune system [11,12]. A minimum of 10 units of glucose is necessary to trigger an immune response [8,13,14].

Evaluations of the effective potential of linear β-(1,3)-glucans have been often made by testing preparations from plant/algae/fungal sources that are always contaminated by pigments, proteins, and membranes resulting in non-specific immunoresponses. β-(1,3)-glucan purified from paramylon synthetized by the WZSL mutant lacks any kind of contaminations from cellular components, which are always present in the paramylon extracted from WT cells. β-1-3-glucan purified from the WZSL mutant is further processed to produce linear nanofibers suited for binding to Dectin-1 receptors of target cell membranes. The effect of these nanofibers has been already investigated in our laboratory on tomato plants, animals, and humans [4,15–17]. In tomatoes, paramylon nanofibers modulate conductance to carbon dioxide (CO2) diffusion from air to the carboxylation sites by regulating hormone levels and water-use efficiency, leading to an increase of plant defense capacity against drought [16].

Tomatoes are the most cultivated vegetable worldwide, being one of the most nutritionally and economically important crops. They require large amounts of water and are adversely affected by drought, which limits photosynthesis and, consequently, plant growth and yield worldwide [18]. Hence, we investigated the role of β-1,3-glucan nanofibers as elicitors of tomato plants response to drought to understand their physiological and photosynthetic responses to this stress.

In this study, tomato cv. Micro-Tom was chosen because of its small size (10–20 cm in height) and short life cycle of about 3 months. Plants were grown in an aeroponic system under three cultivation conditions: optimal water regimen, water scarcity regimen (drought), and water scarcity regimen (drought) coupled with a root treatment with paramylon to monitor and compare eco–physiological (leaf water potential, CO2 assimilation rate, stomatal conductance, internal CO2 concentration, photosystem II (PSII) photochemical efficiency, actual photon yield of PSII, and photochemical quenching of PSII), physicochemical (dry biomass, ashes, dry matter, moisture, microelements, weight, and size), and quality parameters (antioxidant compositions and activities, as well as soluble carbohydrates) throughout the lifecycle of the plants.

#### **2. Materials and Methods**

#### *2.1. Aeroponic Culture System*

The aeroponic cultivation system used was the Nutriculture Twin Amazon 16 (Nutriculture DGS, UK), which consists of a 100 L reservoir tank (160 × 75 × 46 cm), a root chamber housing the delivery system and a molded plastic lid holding sixteen 75 mm mesh pots (Figure 1). The Maxi Jet 1000 pump (14 W, flow rate 1000 Lh−1, max head height 142 cm, NEWA Tecno Industria SRL, Italy) supplies a powerful spray creating a miniature rainstorm inside the chamber through eight 360 degree sprinklers that shower the whole root area, leaving no blind spots. Since the aeroponic container can be assimilated to a closed system, the condensation of growth medium eventually balances its evaporation, reducing the amount of evapotranspiration.

**Figure 1.** The aeroponic cultivation system.

#### *2.2. Paramylon Nanofibers Preparation*

Paramylon granules were extracted and purified from two-days-old cultures of *Euglena gracilis* WZSL mutant according to Barsanti et al. [2]. Nanofibers were obtained by alkaline degradation of the granules [2]. Assuming a 20% loss during the procedure, the final concentration of β-1,3-glucan nanofibers was about 0.8% w/v.

#### *2.3. Plant Material, Growth Conditions and Paramylon Treatment*

Seeds of *Solanum lycopersicum* L., cv. Micro-Tom [19] were surface sterilized with a bleach solution (commercial bleach 30% v/v, Triton X-100 0.02% v/v) for 15 min, washed 3 times with sterile water, and placed overnight in the dark at 4 ◦C. Seeds were then sown on filter paper covering the bottom of 150 mm diameter Petri dishes (about 20 seeds per plate); the paper was wetted with sterile demineralized water, the dishes were wrapped with Parafilm, and they were placed in the dark for about 2 days to aid germination. Upon germination, the cover was removed to allow the development of the roots (2–3 days). Seedlings were then transferred to rock wool (Grodan® Pro Plug) plugs, one seed per plug, to grow under controlled climate conditions (16/8 h light/dark; 400 μmolm−<sup>2</sup> s−<sup>1</sup> PAR, Photosynthetic Active Radiation; 22 ◦C) for approximately 2 weeks in a plastic tray containing half strength nutrient solution [Ca(NO3)2 4H2O, 0.944 gL−1; KNO3, 0.808 gL−1; MgSO4 2H2O, 0.492 gL−1; NH4H2PO4, 0.152 gL−1; microelements, 0.050 gL−1; pH 5.5] according to the protocol by Motohashi et al. [20] in a non-circulating hydroponic method. Upon appearance of the first true leaf, seedlings selected for uniform development were transferred to 3 aeroponic cultivation systems, 16 plants per system, in a glasshouse with a mean temperature of 24 ◦C, a 14/10 h light/dark photoperiod, and 800 μmolm−<sup>2</sup> s−<sup>1</sup> PAR irradiance. Plants were transferred into the greenhouse on May 14, 2018, 39 days after germination (dag). Each aeroponic tank was filled with 100 L of half strength nutrient solution, average pH (±SE) 5.50 ± 0.02.

Three cultivation conditions were tested: optimal water regimen (WW\_P−), water scarcity regimen (drought, WS\_P−), and water scarcity regimen (drought) coupled with root treatment with paramylon (WS\_P+).

Paramylon was added to the nutrient solution of one of the 3 tanks to a final concentration of 500 mg L<sup>−</sup>1. The paramylon concentration was chosen according to the results of previous experiments [16]. The atomization spray time and interval time were 3-s on/5 min off in the control system (optimal water regimen), and 3-s on/120 min off in the other two systems (water scarcity regimen with and without paramylon). As a consequence, the irrigation supply was 8.64 Lplant−<sup>1</sup> d−<sup>1</sup> in WW\_P<sup>−</sup> and 0.36 Lplant−1d−<sup>1</sup> in both WS\_P<sup>+</sup> and WS\_P−. The parameters of the aeroponics system used in the experiment are shown in Table 1.

The optimal water regimen was chosen according to Johnstone et al. [21], who used an aeroponic system for nutritional studies on *Lycopersicon esculentum* Mill. cv Cannery Row and established a misting regimen to plant roots of about 10 Ld−<sup>1</sup> plant<sup>−</sup>1. The water scarcity regimen (0.36 Ld−<sup>1</sup> plant<sup>−</sup>1) was assessed previously by a watering threshold experiment and by checking the representative features of the plants for yellowing and wilting symptoms in 100 tomato plants 30 dag in early spring. We want to stress that the irrigation supply did not correspond to the water needed by the plants.

The levels of nutrient solution and pH were monitored daily; a fresh solution was added in order to maintain a volume of 100 L in each tank. Controlled conditions were maintained throughout the experiment (14/10 h light/dark; 800 <sup>μ</sup>molm−<sup>2</sup> <sup>s</sup>−<sup>1</sup> PAR; 24 ◦C; pH (±SE) 5.5 <sup>±</sup> 0.022).


**Table 1.** Parameters of the aeroponic system used in the experiment.

#### *2.4. Scanning Electron Microscopy (SEM) Preparations*

Root samples were fixed in 100% methanol for 20 min and then transferred in 100% dry ethanol for 30 min with a further change into fresh 100% ethanol overnight. After dehydration, samples were dried in a critical-point dryer apparatus, coated with gold and viewed using a Philips-SEM 505 microscope (Eindhoven, The Netherlands).

#### *2.5. Water Potential, Gas Exchanges and Chlorophyll a Fluorescence*

Preliminary measurements of water potential, gas exchanges and chlorophyll *a* fluorescence were done to assess whether there were variations of these parameters in the different leaves of each plant. Since no variation was detected, measurement was performed on a single leaf per plant, using the same leaf for all the measurement of the experiment.

The first measure of all the eco–physiological parameters was performed upon the transfer of the plants to the aeroponic system, i.e., 39 dag. The predawn leaf water potential (Ψw) was measured on one fully expanded mature leaf per plant (*n* = 16), using a Scholander-type pressure chamber (model 600, PMS Instrument, Albany, OR, USA) and N2 for the application of pressure, following the precautions suggested by Turner and Long [22].

Leaf gas exchanges and chlorophyll *a* fluorescence measurements were determined between 10:00 AM and 1:00 PM (solar time) on one fully-expanded mature leaf plant (*n* = 16). Instantaneous measurements of steady state photosynthetic carbon dioxide (CO2) assimilation rate (A), stomatal conductance (gs), and internal CO2 concentration (Ci) were performed using an LI-6400 portable photosynthesis system (Li-Cor, Lincoln, NE, USA), according to Scartazza et al. [16]. The modulated chlorophyll *a* fluorescence and the status of the electron transport of photosystem II (PSII) were measured with a PAM-2000 fluorometer (Walz, Effeltrich, Germany) on the same leaves used for gas exchange after 40 min of dark-adaptation. Fluorescence in light-adapted leaves was induced according to Scartazza et al. [16] and Schreiber et al. [23]. The maximum efficiency of PSII photochemistry was calculated as Fv/Fm = (Fm−F0)/Fm; the actual photon yield of PSII photochemistry (ΦPSII) was calculated as (F'm−F')/F'm; the photochemical quenching qP was calculated as (F'm−Ft)/(F'm−F'0). The used variables in dark-adapted state were as follows: Fv was the variable fluorescence, Fm was the maximal fluorescence, and F0 was the minimum fluorescence. The used variables in light-adapted state were: F was the fluorescence at the actual state of PSII reaction centers, F'm was the maximal fluorescence, F'0 was the minimal fluorescence, and Ft was the transient fluorescence.

#### *2.6. Physicochemical Parameters and Mineral Content*

At the end of the experiment, all plants were sampled and separated in fruits, leaves, stems, and roots. Fruits were harvested at the ripening category intermediate between VI and VII [24]. All the fruits produced by each plant were gathered and samples (16 fruits) for the analysis were taken from this pool. Several quality attributes were determined on the plants: fresh and dry weights, dry matter content, size (using Vernier calipers), number of fruits, and total ash. The roots, stems, leaves, and fruits of all the plants were dried at 40 ◦C for 96 h in a ventilated oven. One gram of dry matter was ashed at 550 ◦C for 6 h. Total micronutrient content (Ca, Cu, Fe, K, Mg, Mn, Na and Zn) was determined in a sub-sample after digestion with HNO3 and HClO4 [25]. The digests were analyzed by means of a Varian AA240 FS (Varian, USA) hydride generation atomic absorption spectrophotometer equipped with flow vapor generation accessory VGA 77 (Agilent Technologies, USA). Eight independent replicates were used for chemical analysis, and data are presented as mean ± standard deviation (*n* = 8).

#### *2.7. Antioxidant Compounds, Total Antioxidant Capacity and Carbohydrates*

Retinol content was obtained using standard conversion formula (1 μg retinol = 1 retinol equivalent (RE) method used; 1 μg β-carotene = 0.167 μg RE), according to Aremu and Nweze [26]. Lycopene and β-carotene contents were measured spectrophotometrically according to Georgé et al. [27]. Dried tomatoes (500 mg) were homogenized in a mortar with 100 mL of hexane/acetone/ethanol (50/25/25, v/v/v) in the dark, ultrasonically disrupted, and centrifuged at 12,000× *g* for 20 min at 4 ◦C. The supernatants were filtered through 0.2 μm Minisart SRT 15 filters and transferred into a separating funnel. The organic phase was washed three times with 20 mL of distilled water (in order to remove acetone and ethanol). The aqueous phase was discarded, and the remaining water in the organic phase was removed by adding anhydrous sodium sulphate. The final volume was made up to 50 mL with hexane. The reaction mixture absorbance was measured at 436 and 450 for β-carotene determination and 503 nm for lycopene determination.

Reduced (ascorbic acid, AsA) and oxidized (dehydroascorbate, DHA) ascorbate contents were measured spectrophotometrically according to Kampfenkel et al. [28]. Fresh fruit samples (250 mg) were homogenized in a mortar with 0.5 mL of 2% (w/v) phosphoric acid and centrifuged at 12,000× *g* for 15 min at 4 ◦C. The AsA assay mixture contained 50 μL of sample extract, 150 μl of K/P 200 mM (pH 7.4), 50 μl of trichloroacetic acid (TCA) 6% (w/v) and 50 μL of water. The total ascorbate (AsA + DHA) assay mixture contained 50 μl of sample extract, 150 μl of K/P 200 mM (pH 7.4), 50 μl of TCA 6% (w/v) and 50 μL of dithiothreitol (DTT) 10 mM. The reaction mixture was left at room temperature for 15 min; 50 μL of N-ethylmaleimide 0.5% (w/v) were added after the reduction of DHA to AsA. The color was developed in both assays by adding the reagents in the following sequence: 250 μL of TCA 10% (w/v), 200 μL of ortho-phosphoric acid 42% (v/v), 200 μL of 2,2-dipyridyl 4.0% (w/v) in ethanol 70% (v/v) and 100 μL of FeCl3 3% (v/v) to a final volume of 1 mL. Controls were also run, and the solution was allowed to stand at 40 ◦C for an Fe2+-bathophenanthroline complex to develop. The DHA levels were estimated on the basis of the difference between total ascorbate and AsA values. A standard calibration curve covering 0–10 nM of AsA or DHA range was used.

Tocopherols were determined by HPLC according to Döring et al. [29]. Fresh fruit samples (250 mg) were homogenized in a mortar with 0.4 mL of 100% HPLC-grade methanol and incubated overnight at 4 ◦C in the dark. The supernatant was filtered through 0.2 μm Minisart SRT 15 filters and immediately analyzed at room temperature with a reverse-phase Dionex column (Acclaim 120, C18, 5 μm particle size, 4.6 mm internal diameter × 150 mm length). Tocopherols were eluted at a flow rate of 1 mL min−<sup>1</sup> using 100% solvent A (acetonitrile/methanol, 75/25, v/v) for the first 14 min, followed by a 3 min linear gradient to 100% solvent B (methanol/ethylacetate, 68/32, v/v) and 15 min with 100% solvent B. Tocopherols were detected at 280 nm. Authentic standards (Sigma-Aldrich, Italy) were used to quantify the tocopherols content of each sample.

The content of carbohydrates in fruits was determined spectrophotometrically according to Aguiar et al. [30] and quantified using a K-SUFRG commercial kit (Megazyme, Wicklow, Ireland), following the manufacturer's protocol.

The antioxidant properties of the fruits were assessed spectrofluorimetrically by the oxygen radical absorption capacity (ORAC) and hydroxyl radical antioxidant capacity (HORAC) assays [31,32]. Fresh fruit samples (10 mg) were added to 0.75 mL of 100% ethanol/methanol/water/formic acid (35:35:28:2, v/v/v/v) and centrifuged at 12,000× *g* for 10 min at 4 ◦C. The supernatant was collected, and 10 μL were mixed with 170 μL of 48 nM fluorescein (FL). The reagents were transferred into the main reagent wells (OptiPlate 96 F plates, Perkin Elmer, Waltham, MA, USA) and incubated at 37 ◦C for 20 min before recording the initial fluorescence (excitation/emission = 485/527 nm). After incubation, 20 μL of the 2,2 -azobis(2-methylpropionamidine) dihydrochloride reagent (51.5 mM final concentration) were added, and fluorescence readings were taken every minute for 60 min. A phosphate buffer (75 mM, pH 7.4) was used as a blank, and a Trolox solution (0.78–25 μM) was used as a standard. The final ORAC values were calculated by using a regression equation between the Trolox concentration and the net area under the FL decay curve and expressed as μmol Trolox equivalents (TE) per gram of fresh weight (FW). In the HORAC assay, 10 μL of supernatant were mixed with 170 μL of 48 nM fluorescein (605 mM final concentration) and incubated at 37 ◦C for 10 min, before recording the initial fluorescence (excitation/emission = 485/520 nm). After incubation, 10 μL of H2O2 (27.5 mM final concentration) and 10 μL of Co(II) (230 μM final concentration) solutions were added, and fluorescence readings were taken every minute for 60 min. A phosphate buffer was used as a blank, and a gallic acid solution (100–600 μM) was used as a standard. The final HORAC values were calculated using a regression equation between the standard antioxidant concentration and the net area under the curve. One HORAC unit was assigned to the net protection area provided by 1 μM gallic acid, and the activity of the sample was expressed as μmol gallic acid equivalent (GAE) per gram of fresh weight (FW).

The content of total phenolic compounds was determined spectrophotometrically according to Waterhouse [33]. Fresh fruit samples (100 mg) were homogenized in a mortar with 5 mL of methanol acidified with 1 % HCl (v/v) for 20 h in the dark at 4 ◦C. Extracts were centrifuged for 15 min at 12,000× *g* at 4 ◦C, and the supernatants were filtered through 0.2 μm Minisart SRT 15 filters and stored in test tubes at −20 ◦C. Fifty μL of a 4-times diluted extract was mixed with 2.45 mL of distilled water and 250 μL of Folin–Ciocalteu's phenol reagent. After incubation at room temperature for 6 min, 750 μL sodium carbonate 7.5% (w/v) and 500 μL of deionized water were mixed. After 120 min incubation at room temperature, the reaction mixture absorbance was measured at 760 nm. A calibration curve was prepared using a standard solution of gallic acid (range 0–1 mg mL<sup>−</sup>1). Eight independent replicates were used for chemical analysis, and data are presented as mean ± standard deviation (*n* = 8).

#### *2.8. Statistical Analysis*

The statistical analysis was performed using JMP 12 (SAS institute, Cary, NC, USA). The normality of the data was preliminary tested by the Shapiro–Wilk W test. If measurements were carried out for more than two time points, data were analyzed using one-way repeated measures ANOVA, and comparison among means was determined by Tukey's HSD (honestly significant difference) multiple comparison test (*p* < 0.05). All the other data were analyzed by Student's *t* test.

#### **3. Results and Discussion**

Aeroponics is a soil-less cultivation system considered a specialized version of hydroponics [34]. It is an air-water system in which the roots of the plant extend and grow inside a closed container in the dark, are exposed to air, and directly sprayed with a nutrient-water mix through atomizers (Figure 1). The aerial portions of the plant (leaves, stem and crown) extend above the wet zone separated from the root. Aeroponics systems are very useful for plant root studies under controlled conditions; we chose it to monitor the response of tomato plants to drought and feasibility of paramylon nanofibers, directly applied to the root system, in modulating the response of the whole plant to this stressor.

The representative features of the plants (aerial parts and root system) 60 dag under the three cultivation conditions tested are shown in Figures 2 and 3. Under the optimal water regimen (Figure 2A), plants showed a normal compact growth habit with short internodes, fully expanded leaves, and regular fruit size typical of the determinate growth of Micro-Tom. The root system was fully developed

with extremely long roots and plenty lateral roots, indicating a superior growth with respect to the plants under the other cultivation conditions (Figure 3A). Drought stressed plants (Figure 2B) showed wilting symptoms, with yellowing and rolling of the lower leaves, as well as a reduced fruit size. The root system appeared reduced in density and length (Figure 3B). Stressed plants with paramylon root treatment (Figure 2C) did not show any wilting; the growth habit was quite compact, the internode length was greater with respect to plants grown under optimal water regimen, and the fruit sizes were comparable. The root system showed a dramatic reduction of both density and length, coupled with an increase of the lateral rootlets (Figure 3C).

**Figure 2.** The representative features of the tomato plants (aerial parts) under the three cultivation conditions tested 60 days after germination (dag): (**A**) Optimal water regimen; (**B**) water scarcity regimen; (**C**) water scarcity regimen coupled with paramylon root treatment.

**Figure 3.** The root system of the tomato plants under the three cultivation conditions tested 60 dag. (**A**) Optimal water regimen, root length about 100 cm; (**B**) water scarcity regimen, root length about 80 cm; (**C**) water scarcity regimen coupled with paramylon root treatment, root length about 10 cm.

Figures 4 and 5 show the variations of the main eco–physiological parameters, monitored to highlight the effect of the paramylon root treatment during the life cycle of the plants.

Figure 4 shows that the leaf water potential (Ψw) (Figure 4A), CO2 assimilation rate (A) (Figure 4B), stomatal conductance (gs) (Figure 4C), and internal CO2 concentration (Ci) (Figure 4D) had time series with a similar trend in each growth condition tested. In WW\_P− plants, the values of these parameters were almost constant and comparable with the values present in the literature [35] (Figure 4A–D). These data confirm that the chosen water regimen (8.64 Ld−<sup>1</sup> plant−1) was optimal for Micro-Tom growth [21]. In WS\_P− plants, the four parameters underwent a steep decrease after a week of treatment (46 dag), which reached saturation before the end of experiment (Figure 4A–D). The reduction of the stomatal conductance as a reaction to water stress was the cause of this decreasing trend [36,37]. Additionally, in WS\_P<sup>+</sup> plants, the four parameters underwent a steep decrease after a week of treatment (46 dag), but the values recovered to those of control plants (WW\_P−) after one or two weeks (Figure 4A–D). This delayed effect of paramylon nanofibers was mainly due to the time necessary to colonize the root system (Figure 6).

**Figure 4.** (**A**) Leaf water potential (Ψw), (**B**) CO2 assimilation rate (A), (**C**) stomatal conductance (gs), and (**D**) internal CO2 concentration (Ci) in leaves of *Solanum lycopersicum* cv. Micro-Tom grown under well-watered (WW) and water-stressed (WS) conditions coupled (or not) with root treatment with paramylon (P<sup>+</sup> and P−). Data are shown as mean <sup>±</sup> standard deviation (*n* = 16), and measurements were made starting from 39 dag. In each graph, different letters indicate significant differences among treatments (*p* < 0.05, Tukey's HSD post hoc test).

**Figure 5.** (**A**) Photosystem II (PSII) photochemical efficiency (Fv/Fm), (**B**) actual photon yield of PSII photochemistry (ΦPSII), and (**C**) photochemical quenching state of PSII (qP) in leaves of *Solanum lycopersicum* cv. Micro-Tom grown under well-watered (WW) and water-stressed (WS) conditions coupled (or not) with root treatment with paramylon (P<sup>+</sup> and P−). Data are shown as mean ± standard deviation (*n* = 16), and measurements were made starting from 39 dag. In each graph, different letters indicate significant differences among treatments (*p* < 0.05, Tukey's HSD post hoc test).

**Figure 6.** Optical microscopy and scanning electron microscopy images of tomato root hairs 50 dag. (**A** and **B**) Hair surface of non-treated plants. (**C** and **D**) Hair surface of paramylon-treated plants.

Figure 5 shows that the PSII photochemical efficiency (Fv/Fm) (Figure 5A), photon yield of PSII photochemistry (ΦPSII) (Figure 5B), and photochemical quenching of PSII (qP) (Figure 5C) also had time series with a similar trend for each growth condition tested.

In WW\_P− plants, the values of the three parameters were almost constant (Figure 5A–C). In WS\_P− plants, they underwent a steep decrease after a week of treatment (46 dag), which reached saturation before the end of experiment (Figure 5A–C). Additionally, in WS\_P<sup>+</sup> plants, the three parameters underwent a steep decrease after a week of treatment (46 dag), but their values recovered to those of control plants (WW\_P−) after several weeks (Figure 5A–C).

We can say that paramylon nanofibers induced dehydration tolerance and improved intrinsic water use efficiency by influencing stomatal behaviour. Scartazza et al. [16] monitored the effects of water removal on the Ψ<sup>w</sup> and ΦPSII of paramylon-treated tomato plants. They suggested that the paramylon caused an increase of CO2 diffusional constraints but also promoted the ability of tomato plants to reduce water losses and counteract the reduction of ΦPSII caused by the drought. According to these authors [16], β-glucan nanofibers play a potential role in reducing the sensitivity of PSII to potential dehydration damages thanks to a strong stomatal control associated with the transient modified profile of the three major plant hormones content (i.e., abscissic acid, jasmonic acid, salicylic acid) in the xylem sap. Furthermore, they stated that paramylon induced a consistent increase of gm/gs ratio (mesophyll conductance/stomatal conductance) and carbon gained per unit water used, which represents a relevant adaptive trait under water-limited conditions [16,36].

All our eco–physiological data confirmed the potentiality of paramylon nanofibers in the buffering water stress effect on both the photosynthetic rate and the PSII photochemical efficiency. The rate of linear electron transport (ΦPSII) returned to the value of the control (WW\_P<sup>−</sup> plants) with the consequent increase of the proportion of PSII reaction centres that were open (qP). Therefore, PSII photoinhibition and photodamage were counteracted (e.g., a complete recovery of Fv/Fm and ΦPSII values), reducing the sensitivity of PSII to potential dehydration damages.

The values of the root, stem, leaf and fruit dry biomass measured 95 dag on the plants under the three different cultivation conditions are shown in Figure 7. WS\_P− plants showed a significant reduction only in leaf DW (−23% compared with WW\_P−) data relative to the fruits are not shown because the fruit did not ripe beyond the II category (mature green) [24]. Drought stress conditions were so severe that all the eco–physiological parameters were deeply altered (Figure 2, Figure 5, and Figure 6) and inhibited the progress of ripening.

**Figure 7.** Dry biomass expressed as g dry weight (DW) of leaf, stem, root, and red ripe fruits of *Solanum lycopersicum* cv. Micro-Tom grown under well-watered (WW) and water-stressed (WS) conditions coupled (or not) with root treatment with paramylon (P<sup>+</sup> and P−). Data are shown as mean ± standard deviation (*n* = 8). Measurements were made 95 days after germination (dag). For each parameter, different letters indicate significant differences among treatments (*p* < 0.05, Tukey's HSD post hoc test).

A significant reduction of root, stem and leaf DW was observed in WS\_P<sup>+</sup> plants (−61%, <sup>−</sup>33% and −48% compared with WW\_P−). This reduction of growth could be explained as an adaptation strategy to reduce resource spending and allow the plants to divert energy in the fruits. No significant differences (*p* > 0.05) were observed between red ripe fruits DW of WW\_P<sup>−</sup> and WS\_P<sup>+</sup> plants. It is clear that WS induced a negative effect on leaf biomass and fruit ripening, as confirmed by the reduction of leaf DW and the alteration of fruit-ripening processes. Our results are not consistent with those by other authors [38–40] but are in accordance with Khan et al. [41]. The differences observed in the WS sensitivity may be due to the severity of drought.

The other physicochemical parameters (ashes, microelements, weight, size and yield) and the quality parameters (antioxidant compositions and activities, soluble carbohydrates) were measured on fruits assigned to a ripening category intermediate between the VI (red ripe) and the VII (red overripe) categories (Figure 8) [24].

**Figure 8.** Ripening categories table according to Leide et al. [24]: Fruits used for the measurement of physicochemical and quality parameters belong to a ripening category intermediate between the VI (red ripe) and the VII (red overripe) categories.

No significant differences were observed between the number, weight and dimensions of the fruits of WW\_P<sup>−</sup> and WS\_P<sup>+</sup> (data not shown), though paramylon-treated plants showed precocious fruiting and ripening (Figure 9). As already stated, data relative to fruits of the drought stressed plants without paramylon (WW\_P−) are not shown because the fruits did not ripe beyond the II category (mature green).

The effects of cultivation conditions on red ripe fruits were monitored by the quantification of biometric parameters (ashes, dry matter, and moisture; Figure 10), microelement content, and bioactive compounds (antioxidant compositions and activities, soluble carbohydrates; Figures 11–13). In fruits harvested from WS\_P<sup>+</sup> plants, a significant increase of ashes and dry matter was observed compared with WW\_P− plants, while moisture showed an opposite trend (Figure 10). These data confirmed that paramylon treatment could improve fruit quality.

**Figure 9.** Precocious fruiting and ripening of the fruits of WS\_P<sup>+</sup> plants (**A**) with respect to WW\_P<sup>−</sup> plants (**B**) 74 dag.

**Figure 10.** Biometric parameters (dry ash, dry matter and moisture) in red ripe fruits of *Solanum lycopersicum* cv. Micro-Tom grown under well-watered (WW) and water-stressed (WS) conditions coupled (or not) with root treatment with paramylon (P<sup>+</sup> and P−). Data are shown as mean ± standard deviation (*n* = 8). Measurements were made 95 dag. For each parameter, the data were analyzed by Student's t test. The significant differences are for: \*\*\* = *p* < 0.001 and \* = *p* > 0.05.

**Figure 11.** Antioxidant compounds in red ripe fruits of *Solanum lycopersicum* cv. Micro-Tom grown under well-watered (WW) and water-stressed (WS) conditions coupled (or not) with root treatment with paramylon (P<sup>+</sup> and P−). Data are shown as mean <sup>±</sup> standard deviation (*<sup>n</sup>* <sup>=</sup> 8). Measurements were made 95 dag. For each parameter, the data were analyzed by Student's t test. The significant differences are for: \*\*\* = *p* < 0.001. Abbreviations: DW, dry weight; Total AsA, total ascorbate (reduced and oxidized forms).

**Figure 12.** Antioxidant capacity expressed as oxygen radical absorbance capacity (ORAC) and hydroxyl radical antioxidant capacity (HORAC) in red ripe fruits of *Solanum lycopersicum* cv. Micro-Tom grown under well-watered (WW) and water-stressed (WS) conditions coupled (or not) with root treatment with paramylon (P<sup>+</sup> and P−). Data are shown as mean <sup>±</sup> standard deviation (*<sup>n</sup>* <sup>=</sup> 8). Measurements were made 95 dag. For each parameter, the data were analyzed by Student's t test. The significant differences are for: \*\*\* = *p* < 0.001. Abbreviations: DW, dry weight; GAE, gallic acid equivalent; TE, trolox equivalent.

**Figure 13.** Carbohydrates in red ripe fruits of *Solanum lycopersicum* cv. Micro-Tom grown under well-watered (WW) and water-stressed (WS) conditions coupled (or not) with root treatment with paramylon (P<sup>+</sup> and P−). Data are shown as mean <sup>±</sup> standard deviation (*n* = 8). Measurements were made 95 dag. For each parameter, the data were analyzed by Student's t test. The significant differences are for: \*\*\* = *p* < 0.001. Abbreviations: DW, dry weight.

No significant differences were observed between WW\_P<sup>−</sup> and WS\_P<sup>+</sup> fruits regarding microelements content (data not shown).

The cultivation conditions significantly affected the amount of antioxidant compounds of the fruits: all biochemical parameters (retinol, i.e., Vitamin A, tot AsA, i.e., Vitamin C, tocopherols, i.e., Vitamin E, lycopene, β-carotene and phenols) deeply increased in fruits harvested from WS\_P<sup>+</sup> plants (about two-fold higher than those recorded in WW\_P− plants; Figure 11). A high AsA content is an important trait in tomato fruits, as it prevents oxidative stress (especially during fruit ripening) and thus enhances shelf life [42]. Similarly, an increase of lycopene (a highly characteristic phytonutrient of tomatoes) could affect the final nutritional quality and commercial value of tomato fruit [43]. In fact, lycopene greatly enhances fruit quality thanks to its intense antioxidant activity that suppresses cell proliferation and interferes with the growth of cancer cells [44,45]. In addition, a marked increase of the antioxidant activity expressed as the ORAC and HORAC values was observed in fruits harvested from WS\_P<sup>+</sup> plants (+38% and +15%, compared with WW\_P−; Figure 12). This result confirms the involvement of lycopene, β-carotene, and retinol in the antioxidant response of WS\_P<sup>+</sup> fruits. Tocopherols are non-enzymatic lipid-soluble antioxidants that protect the pigments, proteins, and polyunsaturated fatty acids of the photosynthetic apparatus against reactive oxygen species [46]. It has been reported that the tocopherols content in tomato fruits depends on many factors such as the level of irrigation, light, and NaCl [47]. We found that plants grown under WS\_P<sup>+</sup> conditions produced fruits with a high tocopherol content, thus indicating an induction of defense mechanisms. Phenolic compounds and

AsA represent the main water-soluble antioxidants in tomatoes [48]. In WS\_P<sup>+</sup> fruits, a significant increase of total phenols was observed, suggesting that they contribute positively to the antioxidant activity of the tomato water-soluble fraction by reducing the levels of free radicals due to WS (as confirmed by the increase of ORAC and HORAC levels). This response can be considered an adaptive mechanism to water stress that promotes the *de novo* synthesis of these metabolites [40]. At the end of the experiment, the content of carbohydrates showed a similar trend. Glucose, fructose, and sucrose concentrations, as well as total carbohydrates, in the fruits of WS\_P<sup>+</sup> plants were nearly three-fold higher than in fruits harvested from WW\_P− plants (Figure 13). Carbohydrates are essential for plant growth and survival, as well as maintenance and repair processes, and they are also major sources of cellular energy [49]. These compounds play a key role in regulating overall cellular metabolism, maintaining osmotic equilibrium, and preventing turgor loss in tissue [50,51]. They also can act as scavengers of reactive oxygen species (ROS) and contribute to the protection of membranes and macromolecules [52,53]. Micro-Tom fruits contain the reducing sugars fructose and glucose with trace amounts of sucrose, typical of tomatoes [54]. The carbohydrates composition found here was in agreement with that reported in the literature [55].

#### **4. Conclusions**

Drought is by far the most important environmental stressor in agriculture worldwide, and it is expected to contribute to the severe salinization of more than 50% of world arable land by 2050. The research efforts to improve crop productivity under water limiting conditions, focused mainly on natural selection and the breeding activity of tolerant genotypes. Root treatment could be another method to cope with the drought stress. In this paper, we showed the results of the direct application of paramylon on the root system of Micro-Tom tomatoes. Paramylon extracted from the *E. gracilis* WZSL mutant was processed to linear nanofibers that interacted with the Dectin-1 receptors present on the target cell membranes of tomato roots, enhancing the plant defense capacity against drought. Drought tolerance was achieved by influencing stomatal behavior and inducing an effective improvement of water use efficiency, obtained by modulating the conductance to CO2 diffusion from air to the carboxylation sites through the modulation of hormone levels [16]. We observed that the paramylon treatment allowed the optimal water regimen of about 8.64 L plant−<sup>1</sup> day−<sup>1</sup> to be lowered to 0.36 Lplant−<sup>1</sup> day−<sup>1</sup> without a detrimental effect on the yield and eco–physiological parameters. The great increase of antioxidant compounds (Vitamin A/C/E, lycopene, β-carotene and phenols) together with the increase of carbohydrates (glucose, fructose and sucrose) in the fruits of paramylon-treated plants improved their nutritional value and sensory quality.

These results confirm the biostimulant activity of paramylon in increasing plant adaptation capacity for abiotic stress.

**Author Contributions:** Conceptualization, methodology, investigation, writing—L.B. and P.G.; resources, software, and data curation—P.C.

**Funding:** This research received no external funding.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

*Article*

### **Metabolite-Targeted Analysis and Physiological Traits of** *Zea mays* **L. in Response to Application of a Leonardite-Humate and Lignosulfonate-Based Products for Their Evaluation as Potential Biostimulants**

### **Andrea Ertani , Serenella Nardi 1,\*, Ornella Francioso 2, Diego Pizzeghello 1, Anna Tinti <sup>2</sup> and Michela Schiavon <sup>1</sup>**


Received: 24 June 2019; Accepted: 9 August 2019; Published: 12 August 2019

**Abstract:** The main aim of this study is to identify and investigate specific humates (Hs) as potential biostimulants. Five specialty lignosulfonates (LS1-5), one commercial leonardite-humate (PH), and one commercial lignosulfonate (LH), were analyzed for their carbon, nitrogen, and sulfur contents, and the distribution of functional groups using Fourier transform infrared (FTIR) and Raman spectroscopies. Hs were further supplied for two days to *Zea mays* L. in hydroponics to test their capacity to trigger changes in physiological target-responses. LS1, LS2, LS3, and LS5 determined the most pronounced effects on plant growth and accumulation of proteins and phenolics, perhaps because of their chemical and spectroscopic features. Root growth was more increased (+51–140%) than leaf growth (+5–35%). This effect was ascribed to higher stimulation of N metabolism in roots according to the increased activity of N-assimilation enzymes (GS and GOGAT) and high consumption of sugars for energy-dependent processes. Increased values of RuBisCO, SPAD (Soil Plant Analysis Development values), and leaf sugar accumulation refer to enhanced photosynthesis attributed to Hs. We conclude that Hs tested in this study functioned as biostimulants, but the specialty lignosulfonates were more efficient in this role, possibly because of the type of starting material and process used for their production, which may have influenced their chemical properties.

**Keywords:** *Zea mays* L; lignohumate; lignosulfonate; biological activity; nitrogen metabolism; carbon metabolism; proteins; phenolics; sugars

#### **1. Introduction**

Increasing food production for a developing world population and the protection of environmental resources represents a great challenge in the field of agricultural sciences. Traditional agronomical practices especially have negatively impacted a number of environmental aspects and have been in part responsible for soil and water pollution [1]. In addition, the quality of most agricultural soils has long been injured by the thorough application of mineral fertilizers in order to achieve high crop yield requirements [2].

The decline of soil chemical and physical properties is generally accompanied by the decrease of soil fertility, a reduced content of soil organic carbon, and the impoverishment of microbial communities' biodiversity [3]. Therefore, new advances in support of environmentally friendly crop productions are required. Among them, one potential strategy could be the application of biostimulant products during crop cultivation [4]. Biostimulants are "formulated products of biological origin, either with or without plant growth promoting microorganisms (PGPMs), able to stimulate plant productivity at very low dosages by virtue of synergic effects of the different bioactive constituents" [5]. Biostimulants promote plant nutrition and tolerance to environmental stresses [6,7] and, based on their origin and the starting source for their manufacturing, they are divided in different groups, as follows: Humic substances (HS), seaweed extracts, protein hydrolysates, and microbial inoculants, such as mycorrhizal fungi and rhizobacteria, and beneficial elements [8]

Humic substances or humates are regarded as a major category of biostimulants, with a big market share [9]. They represent the most stable and recalcitrant component of soil organic matter and derive from the chemical and microbial degradation of vegetal and animal residues [10,11]. They are useful for improving the quality of soils, as well as the plant metabolism and root morphological traits, via their interaction with a plurality of biochemical mechanisms and physiological processes occurring at the plant-soil interface [10,11]. Specifically, humic substances stimulate plant growth via hormone-like effects and increased photosynthesis efficiency, enhance the respiration rate, and improve root nutrient uptake through an effect, either direct or indirect, on the expression of genes encoding H+-ATPase isoforms and membrane transporters [7,10–12].

Over the last decades, commercial humic products designed lignohumates have found various applications in environmental technologies and agriculture [13,14] and are commonly used for several industrial purposes [14]. They share similar properties with humic substances in terms of chelation, buffering, and cation exchange capacity because of the great number of carboxylic and phenol groups bonded to the aromatic ring [10,15]. Lignohumates are water soluble anionic polymers containing high and low molecular weight molecules, as well as a large number of charged groups, and are by-products generated from the sulphite process of wood, in which fibers of cellulose are separated from lignin by the action of bisulphite [16,17]. The lignin fraction in wood is sulfonated, degraded and solubilized in water during this procedure [18]. In this way, the production of humates from materials that do not primarily contain them becomes a very fast process, which otherwise would naturally take many years. Researches have only clarified the primary structure of these polymers in part, so far, and only a few studies have investigated their effects on plant growth and metabolism [19–21].

The production of humates derived from different salts of humic acids, such as ammonium humates and potassium (K) humates is increasingly growing. Potassium humates, in particular, are used as biostimulants to ameliorate soil chemical, physical, and biological properties, such as the content of organic matter, water retention capacity, structure, deactivation of toxic metals, and microbiome. In addition, they can increase the efficiency of inorganic fertilizers by prompting plant growth, yield and quality, enhancing nutrient uptake and assimilation, and promoting plant resistance to stress conditions [22–27].

Interestingly, the chemistry and physiological functioning of humates can vary depending on the starting material (e.g., leonardite, wood) from which they originate, extraction processes (KOH extraction for leonardite, wood bisulphite extraction for lignosulfonates) and modification technologies used to obtain the products. Indeed, humates derived from the same source and obtained by the same company can widely differ in composition [28]. On this account and in view of the plant diversity from which humates can be produced, it appears relevant to characterize the marketed products to test their effectivity in agriculture as biostimulants.

In light of such considerations, seven humates were investigated in this study to evaluate their biostimulant potential. The humates included a commercial lignosulfonate-based product (LH, LignoHumate®, produced using a patented oxidation process) consisting of a highly concentrated plant and soil amendment, a commercial humate extracted from leonardite (PH), produced and marketed by Borregaard. The remaining humates (LS1, LS2, LS3, LS4, LS5) were specialty lignosulfonates developed by Borregaard and applying proprietary technology (different from the one used to obtain LH) to

modify the starting material. We first assayed differences in their content of main elements (C, N, and S), and in the occurrence and distribution of principal functional groups using two complementary spectroscopic techniques (FT-IR and FT-Raman). Then, we applied these products to *Zea mays* L. plants in order to evaluate differences in their capacity to trigger positive changes in physiological and biochemical traits associated with plant productivity. We chose to test the products on *Zea mays* L. because it is a relevant staple crop for many populations worldwide. One of the novelties of the study is that most of products tested in this study were specialty lignosulfonates developed by Borregaard's company using proprietary technology and, thus, they were supposed to be very different in chemical features from standard lignosulfonates.

#### **2. Results**

#### *2.1. Chemical and Spectroscopic Features of Hs*

The elemental composition in percent content (w/w) of Hs is reported in Table 1. Carbon (C) content was strongly correlated (*R*<sup>2</sup> = 0.83) with nitrogen (N) content for all Hs and varied from 33.04% (w/w) in LH to 54.56% (w/w) in LS1. Nitrogen content was also maximum in LS1 (2.18% w/w), but minimum in LS4 (1.58% w/w). Sulfur (S) content was low only in PH (1.30%), while it was higher in lignosulfonates, varying from 5.13% (w/w) in LS5 to 7.83% (w/w) in LS4.

**Table 1.** Elemental analysis of carbon (C), nitrogen (N), and sulfur (S) in the different humates.


FTIR and Raman analyses were performed to evaluate the main chemical attributes of Hs. The attributions of the main peaks for different functional groups identified in the FTIR and Raman spectra were mainly obtained by references [29]. With respect to FTIR spectra, we decided to display only the peak fitting results obtained in the region from 1800 to 1370 cm<sup>−</sup>1, because the main differences in variation were observed in this region. The region between 1200 cm−<sup>1</sup> and 1000 cm−<sup>1</sup> was heavily dominated by strong bands, probably originated by the SO3H group vibrations (Figure 1) [30].

**Figure 1.** FTIR histograms of humates (Hs) peak areas processes by using curve fitting (from 1800 to 1370 cm<sup>−</sup>1).

In LS2 and LH, due to C=O bonds of acetyl ester from residual hemicelluloses, a band between 1735 and 1725 cm−<sup>1</sup> was evident. A very weak band at 1705 cm−<sup>1</sup> was observed only in LS5. This band, associated with those at 1258 and 1418 cm<sup>−</sup>1, may be attributed to the C=O stretching of COOH groups, while the other two bands may be due to C (=O)\O stretching vibration and OH in-plane deformation vibrations, respectively (spectra not shown). The appearance of carboxyl acid groups could be related to the removal of hemicellulose in this sample [31]. The bands at 1644 in LS2, LS3, LH, and PH, and at 1632 cm−<sup>1</sup> in LS1 and LH, were likely associated with H2O and C=O stretching in conjugated *p*-substituted aryl ketones [32]. In addition, the peak at 1655 cm−<sup>1</sup> recorded in LS4 could be assigned to C=O in alkyl groups of the lignin side chains, conjugated with the aromatic rings [33]. These bands were completely absent in LS5. Other bands identified between 1600 and 1573 cm−<sup>1</sup> corresponded to vibration of aromatic rings. The intensity of these bands depends on the number of C-O bonds to the aromatic ring [34]. Intermolecular aromatic C=C bonds may also have contributed to the intensity of these bands. The peaks from 1512 to 1498 cm−<sup>1</sup> are typical of the skeletal and stretching vibration of aromatic moieties in lignin. Such peaks were present in all products. The bands at around

1460 and 1414 cm−<sup>1</sup> were attributed to the bending vibration of the methoxyl on benzene rings and methylene groups, respectively. The peak at 1370 cm<sup>−</sup>1, observed only in PH, may be due to aromatic CH generated by cleavage of ether bonds within the lignin (spectra not shown).

The relative area percentage gave an estimation of the functional group distribution in the Hs (Figure 1). The band at around 1640 cm−<sup>1</sup> showed a variable distribution among products. For instance, it was dominant in LS1 (24%), LH (18%), LS2 (13%), and totally absent in LS5. The aromatic structure was diversified into different bands at around 1580, 1559, and 1500 cm<sup>−</sup>1. The first band was dominant in LS4 (21%), LS3 (9.4%), and LH (9.0%), and absent in PH. In the other products, this band ranged from 8% to 2.4%. The second band at 1559 cm−<sup>1</sup> accounted for 24% in PH and 9% in LS1. The last band at around 1500 cm−<sup>1</sup> was prevalent in PH (7.5%), LS1 (7%), and LS3 (5%). In other lignosulfonates, it varied from 4% in LS4 and LS5, 2.5% in LS2, and 1% in LH. Finally, the band at 1371 cm−<sup>1</sup> accounted for 23% in the commercial humate PH.

The Raman spectra of LS2 and LS5 are reported in Figure 2, while the complete attributions of the two lignosulfonates are shown in Table 2. Both spectra display bands at 3490 and 3250 cm−1, attributable to OH stretching free or H-bonded, respectively, and both aliphatic (at 2940 and 2846 cm<sup>−</sup>1) and aromatic (at 3070 cm<sup>−</sup>1) CH stretching in the higher wavenumber region. Moreover, the shoulder at about 1670 cm−<sup>1</sup> could be ascribed to conjugated C=O stretching [35], the bands at 1630, 1604, and about 1500 cm<sup>−</sup>1, together with that one at 1190 cm−1, were all attributable to phenolic rings, the last one specifically to lignin [35,36]. The peaks at 1460, 1370, and 1330 cm−<sup>1</sup> corresponded to bending vibrations of O-CH3, CH, and aliphatic OH in lignin and cellulose, respectively [35]. The peaks at 1284 and 1082 cm<sup>−</sup>1, together with that recorded at 815 cm−<sup>1</sup> indicated the presence of sulfated groups [37,38]. Other bands observed in the Raman spectra were less indicative to identify the functional groups present in LS2 and LS5. The relative intensity of the over reported bands is different in the two examined spectra. In particular, for LS5 the bands attributable to aromatic groups (at 3070, 1633, 1604, and 1190 cm<sup>−</sup>1) displayed a higher intensity compared to LS2, indicating that the aromatic component was higher in LS5. On the contrary, the bands at 1330 and 898 cm<sup>−</sup>1, both attributable to cellulose, were more intense in LS2, indicating a higher content of this component in LS2 compared to LS5.

**Figure 2.** FT-RAMAN spectra of lignosulfonates LS2 and LS5.


**Table 2.** Main bands observed in the Raman spectra of humates LS2 and LS5. S = strong; m = medium; w = weak; v = very; sh = shoulder.

#### *2.2. E*ff*ect of Hs on Maize Plant Growth*

The effect of Hs application on maize plant growth is reported in Figure 3. Results indicated that LS5 was the most effective in promoting the leaf (Figure 3A) and root (Figure 3B) dry weight (+140% and +35%, respectively), compared to the untreated plants. The remaining Hs did not substantially improve the leaf biomass produced by plants. However, they all stimulated the root growth appreciably. Specifically, LS3, LS4, and LH increased the root biomass of plants by 51%, 57% and 52%, respectively, while LS2 and PH were by about 85%, and LS1 was by 111%.

**Figure 3.** Effect of individual humates (Hs) on leaf (**A**) and root (**B**) dry weight of *Z. mays* L. plants. Twelve-day-old plants were supplied for two days with Hs at 1 mg C L<sup>−</sup>1. Different letters above bars indicate significant differences at *p* < 0.05, according to the Student–Newman–Keuls test. Data represent the means of three measurements with ten plants in each (±SD). C = control; LH = commercial lignosulfonate-based product; PH = commercial humate extracted from leonardite; LS1 − LS5 = specialty lignosulfonates.

#### *2.3. E*ff*ects of Hs on SPAD, RuBisCO activity, and N-compounds (Proteins and Phenolics)*

The effect of Hs on maize plants was additionally evaluated in terms of photosynthetic efficiency by measuring the SPAD index (Figure 4A) and the activity of the RuBisCO enzyme (Figure 4B). In general, Hs prompted the increase of the SPAD index values of plants to a similar extent (Figure 4A). Analogously, RuBisCO activity was increased by all Hs, but differences in the percent stimulation

caused by individual Hs were observed in this case (Figure 4B). LS2, in particular, was the most effective in enhancing the activity of this enzyme (by about 70%), followed by LS1, LS3, LS5, and PH (+30–50%). The other Hs stimulated the RuBisCO activity to a lower extent.

**Figure 4.** Effect of humates (Hs) on SPAD index (**A**), RuBisCO activity (**B**), protein content (**C**), and total phenolic compounds (**D**) in leaves of *Z. mays* L. plants. Twelve-day-old plants were supplied with Hs at 1 mg C L−<sup>1</sup> for two days. Different letters above bars indicate significant differences at *p* < 0.05, according to Student–Newman–Keuls test. Data represent the means of three measurements with three plants in each (±SD). C = control; LH = commercial lignosulfonate-based product; PH = commercial humate extracted from leonardite; LS1-LS5= specialty lignosulfonates.

As the SPAD index is associated to the amount of N compounds in plants, the quantification of proteins, total phenols, and individual phenolic acids was performed. It is noteworthy that the content of total N was also measured in the plants (data not shown), but no significant differences were recorded, likely because of the limited duration of the experiment. Protein accumulation was enhanced in leaves of maize plants supplied with Hs (Figure 4C). LS2, LS3, and PH, in particular, induced the most pronounced increases (+74%, +98%, and +104%, respectively). The synthesis of phenol compounds (Figure 4D) was stimulated in leaves of maize plants treated with Hs as well. In this case, however, LS1, LH, LS4, and LS5 were responsible for the greatest increments (by about 80%).

Differential accumulation of individual phenolic acids was also observed between maize plants supplied with Hs and the controls, as well as among plants treated with distinct Hs (Table 3). There were three derivatives of cinnamic acids (caffeic, *p*-coumaric, and ferulic acids), one ester of caffeic acid and (−)-quinic acid (chlorogenic acid), and one derivative of benzoic acid (*p*-hydroxybenzoic acid). In most cases, Hs induced significantly higher accumulation of chlorogenic, caffeic, p-coumaric, ferulic, and p-hydroxybenzoic acids in leaves of maize plants compared to the controls. LS1, LS2, LS3, LS4, and LS5 especially, accounted for the most appreciable effects in this respect. Specifically, very high values of leaf phenolic acid accumulation were measured for chlorogenic and caffeic acids in plants treated with LS2 (+168% and 184%, respectively) and LS4 (+651% and 262%, respectively), for ferulic acid in plants provided with LS1 (+472%), LS2 (328%), LS3 (+222%), and LS4 (+413%), and for *p*-hydroxybenzoic acid in plants given with LS1 (+193%), LS2 (+187%), and LS4 (+202%).

**Table 3.** Profile of phenolic compounds in leaves and roots of*Z. mays* L. Plants were grown for 12 days in a nutrient solution and supplied with individual humates at 1 mg C L−<sup>1</sup> for two days. n. d. =not detectable. Values along the same column following by different letters are statistically different at *p* < 0.05 (*n* = 3, ± SD) according to Student–Newman–Keuls test. C= control; LH = commercial lignosulfonate-based product; PH = commercial humate extracted from leonardite; LS1−LS5= specialty lignosulfonates.


In roots, only chlorogenic and ferulic acids were more accumulated in plants treated with Hs than the controls. The highest values of chlorogenic acid content were observed in roots after plant treatment with LS2 (+71%), LS4 (+60%), LS5 (+115%), PH (+113%). With respect to ferulic acid, maximum accumulation was measured in roots of plants supplied with LS1 (+436%), LS2 (+396%), and PH (+361%).

#### *2.4. E*ff*ects of Hs on GS and GOGAT Activities*

Further effects of Hs on maize plant metabolism were investigated by measuring the activities of two enzymes (GS and GOGAT) that catalyze key steps in N assimilation (Figure 5). Overall, a greater activity of such enzymes was determined in plants supplied with Hs. The activity of GS in leaves in particular, was increased by LH (+44%), LS4 (+24%), and LS5 (+18%) (Figure 5A), while the activity of GOGAT was stimulated by all Hs applied to plants (Figure 5B). LS3 accounted for the maximum leaf activity of GOGAT (+98%). In roots, the activity of both GS and GOGAT enzymes was enhanced by all Hs (Figure 5C,D). In the case of GS, the highest activity was detected in roots of plants treated with LS2 (Figure 5C), while maximum GOGAT activity was measured in plants supplied with LS1 and LS5 (Figure 5D).

**Figure 5.** Effect of humates (Hs) on glutamine synthetase (GS) and glutamate synthase (GOGAT) activity in leaves (**A**, **B**, respectively) and roots (**C**, **D**, respectively) of *Z. mays* L. plants. Twelve-day-old plants were supplied with Hs at 1 mg C L−<sup>1</sup> for two days. Different letters above bars indicate significant differences at *p* < 0.05, according to the Student–Newman–Keuls test. Data represent the means of three measurements with three plants in each (±SD). C = control; LH = commercial lignosulfonate-based product; PH = commercial humate extracted from leonardite; LS1 − LS5 = specialty lignosulfonates.

#### *2.5. E*ff*ects of Hs on Reducing Sugar Accumulation*

The content of soluble reducing sugars (glucose and fructose) was increased in leaves of plants treated with Hs (Figure 6). Precisely, improved glucose accumulation was observed in leaves of maize plants after treatment with LS1, LS2, and LH (+39%, +58%, +41%, respectively, Figure 6A). With respect to fructose, all Hs stimulate its accumulation, with maximum values determined by LS2 and LS3 (+92% and +111%, respectively, Figure 6A). In roots, an opposite trend was evident, as the content of both sugars decreased when plants were treated with Hs, with few exceptions (Figure 6B).

**Figure 6.** Effect of individual humates (Hs) on glucose and fructose accumulation in leaves (**A**) and roots (**B**) of *Z. mays* L. plants. Twelve-day-old plants were supplied with Hs at 1 mg C L−<sup>1</sup> for two days. Different letters above bars (un-bolded for glucose and bolded for fructose) indicate significant differences at *p* < 0.05, according to the Student–Newman–Keuls test. Data represent the means of three measurements with three plants in each (±SD). C = control; LH = commercial lignosulfonate-based product; PH = commercial humate extracted from leonardite; LS1 − LS5 = specialty lignosulfonates.

#### *2.6. Statistical Analysis of Data*

The correlation analysis evidenced significant relationships between the parameters analyzed in maize plants subjected to treatment with Hs (Table S1). The root dry weight, which was more stimulated than the leaf dry weight by Hs, positively correlated with SPAD, total phenols, GS and GOGAT root activity, and RubisCO activity, whereas it negatively correlated with the content of glucose in roots. SPAD index values displayed a positive correlation with the content of N metabolites (proteins and phenols), the activity of GOGAT, GS (only in roots), RubisCO, and the leaf fructose content. However, SPAD negatively correlated with the root glucose content. Total phenols showed positive correlation with GS activity in leaves and roots and GOGAT activity in roots. The activity of GS in leaves did not show any correlation with the other parameters analyzed, but GS activity in roots positively correlated with the activity of GOGAT in leaves and roots. The activity of both N enzymes also positively correlated with RubisCO activity. The activity of all three enzymes, GS (in roots), GOGAT, and RubisCO, negatively correlated with glucose content in roots. RuBisCO positively correlated with the leaf glucose content and fructose content in both leaves and roots, whereas it negatively correlated with the root glucose.

With respect to PCA analysis, three factors accounted for 91% of the total variance. Factor 1 explained 53.6% of the variance and positively correlated with GS and GOGAT activity in roots, SPAD, total phenols, while it negatively correlated with glucose content in roots. Factor 2 explained 22.7% of the variance and was positively correlated with GOGAT activity in leaves, protein content, and leaf fructose amount. Factor 3 explained the remaining 14.8% of the variance and was correlated with the content of fructose in roots and GS activity in leaves. Plotting data reported in Table S2 according to PC1 and PC2 allowed three clusters to be identified (Figure S1A,C); a main group constituted by plants 1, 2, 3, 4, 5, 6, and 7, corresponding to LS1, LS2, LS4, LS5, PH, and LH, and the other two by control (untreated, 8) and LS3 (3). In particular, LS1, LS2, LS4, LS5, PH, and LH were characterized by high values of GS and GOGAT activity in roots, SPAD, and total phenols, whilst LS3 had high values of GOGAT activity in roots and protein. The control plants had higher values of glucose content in roots. Plotting PC1 and PC2 also revealed that, among plants treated with humates LS1, LS2, LS4, LS5, PH and LH, those treated with LH tended to be at the bottom of the cloud, and PH was at the top, along the axis 2. It should be also noted that plotting PC1 and PC3, LH (7) differed from the other treatments for high GOGAT activity in leaves.

#### **3. Discussion**

Humates can differ in composition depending on the source material and process type employed for their production. Therefore, they can show significant variation in biostimulant properties. In this study, we assayed seven humates (a commercial lignosulfonate-based product, a commercial humate extracted from leonardite, and five specialty lignosulfonates provided by Borregaard's company) by determining their elemental content and dissecting the major functional groups occurring in their formulation. Then, in order to determine the plant-growth promoting potential of Hs, we evaluated differences in their capacity to promote plant biomass production, N assimilation into organic compounds (chlorophylls, proteins and phenols), and photosynthesis.

We found that all products were able to stimulate plant growth and the metabolic responses typically triggered by biostimulants. Therefore, untreated plants were different from plants treated with tested Hs in terms of performance, as revealed by PCA analysis. However, LS1, LS2, LS3, and LS5 appeared to be the most effective in this respect, being able to induce the greatest increments (up to 184%) of most physiological parameters (dry weight, root GS activity, GOGAT activity, RuBisCO activity) and targeted-biochemical markers (SPAD, proteins, phenols, fructose content) in maize, compared to the untreated plants. A general overview of such increments is depicted in the heat map of plant-associated parameters influenced by individual humates, reported in Figure S2. LS2 and LS3 contained a similar percent content of total C and N, as well as LS1 and LS5. The spectroscopic characteristic of all samples and especially LS2 and LS5 revealed the presence of cellulose residues and

aromatic groups. LS4 and PH contained the highest percentage in aromatic groups according to the deconvolution process of FT-IR spectra, while for LS1 the functional group distribution appeared to be a mixture of the same groups observed in LS4 and PH, but with a considerable hydrophilic feature (see the band at 1632 cm−1). Therefore, the C and N composition and profile in functional groups of specialty products LS1, LS2, LS3, and LS5 could explain their better efficiency as biostimulants compared to the lignosulfonate LH.

Overall, root growth was more stimulated (+51–140%) than leaf growth by all Hs (+5–35%), with more pronounced effects observed in plants treated with LS1 and LS5. These results are in line with the current literature that reports early root growth as a typical response of plants treated with humic substances, while the stimulation of leaf growth is generally recognized as a delayed response [10,17,39]. One possible explanation of this effect is that humic substances can act on root development by influencing the hormonal balance within the plants and nitric oxide distribution, either directly or indirectly, and by modifying the nutrient uptake by plants and the activity of root membrane H+-ATPase [8,39,40]. Early root development could also be ascribed to the biological properties of humic substances, whose hormone-like activity has been previously described [41,42], and that Hs tested in this study might possess as well. Ertani et al. [17], in particular, reported the auxin-like and gibberellin-like activity of two lignosulfonates, and the gibberellin-like activity of a leonardite humic acid. The hormone-like activity of humic substances and commercial humates are likely due to their content in auxin-like substances, as well as to the presence of phenol-C groups with biological activity [43,44].

Hs were also effective in promoting N metabolism. In particular, LS1, LS2, and LS5 determined the highest increases in the activity of N assimilation enzymes, i.e., glutamine synthetase (GS) and glutamate synthase (GOGAT), in roots. This finding could explain why plants treated with these products developed their roots more. In this respect, the root dry weight of maize plants positively correlated with GS and GOGAT root activity. In general, all Hs enhanced the activity of GS and GOGAT more in roots than in leaves, which may suggest that early root growth stimulation in maize by Hs was also a result of a more pronounced N metabolism enhancement and decreased N storage. Similar findings and hypothesis have been previously reported by Jannin et al. [45]. Higher activity of N enzymes in roots might be due to metabolic changes related to differences in the root/shoot nitrate balance occurring under LH treatment [39]. In leaves, GOGAT activity was significantly stimulated by all Hs, while GS activity was stimulated by only four of them. Such differences could be ascribed to distinct mechanisms of regulation of N enzymes induced by several factors, including N metabolites (e.g., ammonium, glutamine, and glutamate) that are known to exert feedback effects [46–48]. In this respect, those Hs determining the highest increases in leaf protein accumulation were responsible for the least increases in GS leaf activity. Interestingly, they also stimulated the accumulation of phenolic compounds as the other Hs, but to a less extent. This observation seems to suggest that when plants are treated with Hs, two preferential metabolic pathways can be mainly stimulated, i.e., the N primary metabolism that produces proteins and the secondary metabolism involved in the synthesis of phenolics. These two metabolic pathways have been previously identified as principal targets of humic substances and other biostimulants, including lignosulfonate-humates, in maize and other plant species [17,49]. With respect to phenolic compounds, the increase in content of a number of them, especially in leaves, to levels that were not injurious to plants, can be deemed as an important result because these phytochemicals have recognized health beneficial properties, are implied in the plant defense responses against stress conditions, and mediate plant relationships with ecological partners [50–53].

The positive effects of all Hs on plant metabolism was also confirmed by the increased activity of RuBisCO, i.e., the enzyme responsible for CO2 fixation in the Calvin cycle. Indeed, measuring the RuBisCO activity allowed for knowing whether Hs stimulated the photosynthetic efficiency of plants, because higher activity generally corresponds to higher photosynthetic rates and productivity. The increased activity of RuBisCO in plants under treatment by humic substances could be due to

increased number of chloroplasts per cell, as proposed by Jannin et al. [45]. RuBisCO activity positively correlated with the SPAD index values and the leaf content of reducing sugars. Similar results were previously reported by Ertani et al. [17].

In our study, we observed a reduction in glucose and fructose accumulation in roots of maize plants. Glucose is mainly produced in the cytosol from triose-phosphate precursors produced during the Calvin cycle and its accumulation in cells is influenced by different factors, like the photosynthetic rate, the need of glucose for energy-dependent processes, and the metabolic fate of the precursor glutaraldehyde 3-P (including the synthesis of starch). In roots, the level of carbohydrates depends on the source of N they receive (NO3, NH4, or amino acids), the rate of transport of photosynthates and the quantity of reserves that are stored in the root tissues. The different distribution of glucose between leaves and roots also depends on the need of the plant to use glucose in a specific organ for a metabolic requirement. The decrease of glucose in the roots, for instance, may be indicative of a high demand for ATP-dependent nutrient transport and other energy-requiring processes in the root cells, including growth processes, and could be associated with the increased need of C-skeleton for the synthesis of N compounds. A similar reasoning can be made for fructose.

#### **4. Materials and Methods**

#### *4.1. Elemental Composition and Spectroscopic Analysis of Hs*

Seven humates (Hs) were tested in this study for their biostimulant properties. All these products completely dissolved in H2O without leaving insoluble clumps. The carbon (C), nitrogen (N), and sulfur (S) contents of Hs were determined via dry combustion conducted in the element analyzer vario MACRO CNS (Hanau, Germany).

The Fourier transform infrared (FTIR) spectra of these products were recorded using an ALPHA FTIR spectrometer (Bruker Optics, Ettlingen, Germany) equipped with an ATR (attenuated total reflectance) sampling device containing diamond crystals. The absorbance spectra were recorded between 4000 cm−<sup>1</sup> and 400 cm−1, at a spectral resolution of 4 cm−1, with 64 scans co-added and averaged. A background spectrum of air was recorded under the same procedure conditions before each series of measurements. Spectra were processed with the Grams/386 spectroscopic software (version 6.00, Galactic Industries Corporation, Salem, NH). Overlapping peaks were resolved using a peak fitting analysis in the spectral region from 1800 to 1000 cm−<sup>1</sup> by using the Grams/386 spectroscopic software (version 6.00, Galactic Industries Corporation, Salem, NH). The overlapping bands were resolved with a Gaussian function. The best fitting parameters were determined by minimization of the reduced Chi square (χ2). Good agreement between experimental and calculated profiles was obtained, with coefficients of determination, R2, ranging from 0.999 to 0.988 and the standard error, SE, from 0.001 to 0.003. All data are expressed as percentage area.

FT-Raman spectra of Hs were recorded in solid state with a Multiram FT-Raman spectrometer (Bruker Optics, Ettlingen, Germany) equipped with a cooled Ge-diode detector. The excitation source was a Nd-YAG laser (1.064 nm, about 30 mW laser power on the sample) in the backscattering (180◦) configuration. The low laser power was due to the brown color of the samples, which burned out using a higher laser power. As a consequence of burning, it was possible to record only the spectra of LS2 and LS5.

#### *4.2. Plant Material and Experimental Design*

Seeds of *Zea mays* L. (P1921, Pioneer HI-BRED, Italia Sementi S.r.l.) were soaked in distilled water overnight and then surface-sterilized in 5% (v/v) sodium hypochlorite for 10 min while shaking. Seeds were germinated on filter paper wetted with distilled water for 60 h in the dark at 25 ◦C. Seedlings were then transferred into 3 L pots in the presence of a thoroughly aerated Hoagland solution, with a density of 24 plants per pot. The nutrient solution was renewed every 48 h and contained the following salts (μM): KH2PO4 (40), Ca(NO3)2 (200), KNO3 (200), MgSO4 (200), FeNaEDTA (10), H3BO3 (4.6), CuCl2·2H2O (0.036), MnCl2·4H2O (0.9), ZnCl2 (0.09), and NaMoO·2H2O (0.01). Plants were grown inside a chamber with 14 h of light per day, in air temperatures of 21 ◦C (night) and 27 ◦C (day), at a relative humidity of 70/85%, and with a photon flux density of 280 mol m−2s−1. After twelve days of growth in hydroponics, each Hs was added in a unique application to the nutrient solution at 1 mg C L−<sup>1</sup> (for each treatment with single Hs, 3 pots were prepared). After 48 h from the addition of Hs, plants were harvested. The choice of this short incubation time was dictated by results obtained in several previous studies, where a period of 24–48 h was found to induce early molecular responses and morpho-physiological changes in both roots and leaves. Plants that were not added with Hs served as controls (3 pots, 24 plants per pot).

At the end of the treatment, plants were randomly harvested and then carefully washed and dried with blotting paper. A sub-sample of the plant material was immediately frozen with liquid nitrogen and kept at −80 ◦C, to be used for biochemical analyses. For dry weight measurement, 10 plants randomly harvested were used (ten per treatment from each pot). The samples were placed in a drying oven for 2 d at 70 ◦C and allowed to cool for 2 h inside a closed bell jar. The dry weight of individual roots and leaves was measured for each plant.

#### *4.3. Determination of the SPAD Index*

The relative chlorophyll content was determined using a non-destructive method that employed light transmission across a leaf, at two wavelengths, to quantify the greenness and thickness of leaves. The ratio of the transmission of the two wavelengths provides a chlorophyll content index that is also named the SPAD index. The analyses were performed using a SPAD (Soil Plant Analysis Development) chlorophyll meter (SPAD-502 model, Minolta Camera Co, Ltd., Osaka, Japan) and the SPAD index was measured on the last expanded leaf of maize plants. The determination was carried out on 5 measurements per leaf from 10 plants per each treatment.

#### *4.4. Analysis of Soluble Proteins and Reducing Sugars*

For protein extraction, frozen foliar tissues (100 mg) of five plants per pot were ground in liquid nitrogen and vortexed in the presence of 5 mL buffer (100 mM Tris-HCl pH 7.5, 1 mM Na2EDTA, 5 mM DTT) and centrifuged at 14,000 g. The supernatants were mixed with 10% (w/v) trichloroacetic acid and then centrifuged. The pellets were finally re-suspended in 0.1 N NaOH. The protein concentration was determined using the Bradford method through a UV/VIS spectrophotometer (Lambda 1, Perkin-Elmer, Monza, Italy) at λ = 595 nm. Protein concentration was expressed as mg of protein g−<sup>1</sup> fresh weight (FW).

For reducing sugar analysis, foliar tissues (100 mg) of five plants per pot were dried for 48 h at 80 ◦C, ground to obtain a fine powder, and then extracted with 2.5 mL 0.1 N H2SO4. Samples were incubated in a heating block for 40 min at 60 ◦C and then centrifuged at 6000 g for 10 min at 4 ◦C. Supernatants were filtrated (0.2 μm, Membra-Fil® Whatman Brand, Whatman, Milan, Italy) and further analyzed via HPLC (Perkin Elmer 410). Soluble sugars were separated using a Biorad Aminex 87 C column (300 <sup>×</sup> 7.8 mm) with H2O as eluent at a flow rate of 0.6 mL min<sup>−</sup>1. Sugar concentration was expressed as mg g−<sup>1</sup> dry weight (DW).

#### *4.5. Analysis of Total and Individual Phenolic Compounds*

The content of total phenols in plant samples was quantified using the Folin–Ciocalteu method. For individual phenol detection, extraction from frozen plant material of five plants (1 plant = 1 biological replicate) was performed using water/methanol (1:1 v/v), filtered at 0.45 μm. Phenols were separated via an HPLC 2700 (Thermo Finnigan, San Jose, CA, USA) coupled with an 1806 UV/Vis (Thermo Finnigan, San Jose, CA, USA) detector. The column was a TM-LC 18 (Supelcosil) equipped with pre-column TM-LC 18 (Pelliguard, Supelco). Elution was conducted at a flow rate of 1.2 mL min−<sup>1</sup> using a mixture of water/ n-butanol/ acetic acid (80.5:18:1.5 v/v) as the mobile phase. The injection volume of each sample was 20 μL. Detection was performed at λ = 275 nm and the identification of compounds was obtained by comparison of their retention time values with those of corresponding

standards. The calibration curve and quantification were performed considering the relationship between peak areas vs. standard concentrations at four concentrations (*n* = 4). A linear fitting with an R squared value of (*R*2) = 0.99 was obtained.

#### *4.6. Determination of GS, GOGAT and RuBisCO Activity*

For the assay of glutamine synthetase (GS) and glutamate synthase (GOGAT) enzyme activity, fresh root and leaf tissues (1 g) were ground in a mortar with 10 mL of 100 mM Hepes-NaOH solution at pH 7.5, 5 mM MgCl2 solution, and 1 mM dithiothreitol. For the RuBisCO enzyme, the extraction protocol was the same as for GS and GOGAT, but the enzyme activity in this case was measured in leaves only and the ratio of plant material to buffer was 1:3 (w/v). The extracts were filtered through two layers of muslin and centrifuged at 20,000 g for 15 min at 4 ◦C. The supernatants were used for enzymatic assays.

For the glutamine synthetase (GS EC 6.3.1.2) assay, each mixture contained 90 mM imidazole-HCl (pH 7.0), 60 mM hydroxylamine (neutralized), 20 mM KAsO4, 3 mM MnCl2, 0.4 mM ADP, 120 mM glutamine, and enzyme extract. The assay was performed in a final volume of 750 μL. The enzymatic reaction was developed for 15 min at 37 ◦C. The α-glutamyl hydroxamate was colorimetrically determined by addition of 250 μL of a mixture (1:1:1) of 10% (w/v) FeCl3·6H2O in 0.2 M HCl, 24% (w/v) trichloroacetic acid and 50% (w/v) HCl. The optical density was measured at λ = 540 nm. Enzyme activity was expressed in μmol−<sup>1</sup> g−<sup>1</sup> FW, representing the amount of enzyme catalyzing the formation of 1 nmole γ-glutamyl-hydroxamate min<sup>−</sup>1.

The glutamate synthase (GOGAT EC 1.4.7.1) assay contained 25 mM Hepes-NaOH (pH 7.5), 2 mM L glutamine, 1 mM α-ketoglutaric acid, 0.1 mM NADH, 1 mM Na2EDTA, and 100 μL of enzyme extract. GOGAT activity was measured spectrophotometrically by monitoring NADH oxidation at λ = 340 nm. The enzyme activity was expressed in μmol−<sup>1</sup> g−<sup>1</sup> FW, representing the amount of enzyme catalyzing the oxidation of 1 nmole NADH min<sup>−</sup>1.

The activity of RuBisCO (EC 4.1.1.39) was determined spectrophotometrically in a coupled assay by measuring the production of 3-phosphoglycerate following a 5 min period of incubation with 2 mL of 10 mM MgCl2 and 20 mM NaHCO3 [54].

For each enzyme activity assay, analyses were conducted in three biological replicates (1 plant = 1 biological replicate) per treatment and the absorbance in the samples was measured using a JASCO V-530 UV/VIS spectrophotometer.

#### *4.7. Statistical Analysis*

For all determinations, the analysis of variance (ANOVA) was performed using the SPSS software version 19.0 (SPSS Inc. 1999), which was followed by pair-wise post hoc analyses (Student–Newman–Keuls test) to determine which means differed significantly at *p* < 0.05 (±SD). The number of biological replicates varied depending on the analysis performed and is indicated in the figure and table legends. Correlations between variables were determined using Pearson's coefficient. To identify the structure of the interdependences between the main parameters, a joint principal component analysis (PCA) was performed on the following variables, considering both untreated plants (control) and plants treated with the different humates: Root dry weight, leaf dry weight, SPAD, proteins, total phenols, leaf GS, root GS, leaf GOGAT, root GOGAT, RuBisCO, leaf glucose, leaf fructose, root glucose, and root glucose. The standardized variables were subjected to PCA. Rotated orthogonal components (varimax method of rotation) were extracted and the relative scores were determined. Only PCs with an eigenvalue > 1 were considered for the discussion. Statistics were performed using SPSS software version 25.0 (SPSS, Chicago, IL).

#### **5. Conclusions**

In conclusion, the current study provides clear evidence that all tested products acted as biostimulants. Additionally, the specialty lignosulfonates provided by Borregaard's company were apparently the most effective in this role, likely because of the novel process employed for their production and the products' chemical features (e.g., different C content values and presence of functional groups). These results support the importance of setting up new technologies and advanced industrial processes for the production of novel commercial humates and lignosulfonates with better formulation performance, which can be used as efficient biostimulants during crop cultivation in the framework of sustainable agriculture. Future studies could be performed in field trials and using other crop species, including horticultural crops, to definitely confirm the positive characteristics of these products under varying and/or stress conditions.

**Supplementary Materials:** The following are available online at http://www.mdpi.com/2073-4395/9/8/445/s1, Table S1: Correlations between variables determined using Pearson's coefficient. Asterisks indicate significant correlation at *p* < 0.05 (\*) or *p* < 0.01 (\*\*). r = root, l = leaf, dw = dry weight, TP = total phenols, GS = glutamine synthetase, GOGAT = glutamate synthase, FRU = fructose, GLU = glucose, PROT = proteins, Table S2: Loadings values of the plant variables on the axes identified by principal components (PC) analysis for the different types of treatment and control. r = root, l = leaf, dw = dry weight, TP = total phenols, GS = glutamine synthetase, GOGAT = glutamate synthase, FRU = fructose, GLU = glucose, PROT = proteins. Figure S1: Position of the treated and untreated plants (1 = LS1, 2 = LS2, 3 = LS3, 4 = LS4, 5 = LS5, 6 = PH, 7 = LH, and 8 = control) in the reduced space of the first two principal components (PC1 and PC2) (A) and on PC1 and PC3 (B); variables projected in the plane determined by PC1 and PC2 (D) and PC1 and PC3 (C). r = root, l = leaf, dw = dry weight, TP = total phenols, GS = glutamine synthetase, GOGAT = glutamate synthase, FRU = fructose, GLU = glucose, PROT = proteins, Figure S2: Heat map of plant-associated parameters influenced by individual humates. Different colors indicate different levels of induction/repression (more red more repression, more blue more induction). r = root, l = leaf, dw = dry weight, TP = total phenols, GS = glutamine synthetase, GOGAT = glutamate synthase, FRU = fructose, GLU = glucose, PROT = protein.

**Author Contributions:** O.F. and A.T. performed the spectroscopic analysis and wrote the relative part in the ms; A.E. performed the physiological analyses, bioassays and chemical analyses of the products and wrote the ms; M.S. wrote the ms; D.P. critically read the ms and performed the statistical analyses of data; S.N. designed the study. All the authors critically reviewed the ms.

**Funding:** This study was funded by Borregaard (Hjalmar Wessels vei, Sarpsborg, Norway).

**Acknowledgments:** We would like to thank Federica Zanellato for the precious help in conducting the physiological analyses.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


© 2019 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (http://creativecommons.org/licenses/by/4.0/).

### *Article* **Biostimulant Application Enhances Fruit Setting in Eggplant—An Insight into the Biology of Flowering**

#### **Alicja Pohl, Aneta Grabowska, Andrzej Kalisz and Agnieszka S ˛ekara \***

Department of Vegetable and Medicinal Plants, University of Agriculture in Krakow, 29-Listopada 54, 31-425 Kraków, Poland

**\*** Correspondence: agnieszka.sekara@urk.edu.pl; Tel.: +48-12-662-52-16

Received: 15 July 2019; Accepted: 23 August 2019; Published: 26 August 2019

**Abstract:** Eggplant (*Solanum melongena* L.) is a warm climate crop. Its cultivation extends to temperate regions where low temperatures can affect the course of the generative phase, which is primarily sensitive to abiotic stress. The novelty of the present investigation consisted of characterising the heterostyly, pollination, and fertilisation biology of eggplants in field cultivations, which provided a basis for explaining the effect of a protective biostimulant on these processes. We aimed to investigate the flowering biology of three eggplant hybrids treated with Göemar BM-86®, containing *Ascophylum nodosum* extract, to determine the crucial mechanisms behind the increased flowering and fruit set efficiency and the final effect of increased yield. The flower phenotype (long, medium or short styled), fruit setting, and the number of seeds per fruit were recorded during the two vegetation periods. The numbers of pollen tubes and fertilised ovules in ovaries were evaluated during the generative stage of development to characterise the course of pollination and fertilisation for all types of flowers depending on the cultivar and biostimulant treatment. The expression of heterostyly depended on the eggplant genotype, age of the plant, fruit load, and biostimulant treatment. Domination by long-styled flowers was observed, amounting to 41%, 42%, and 55% of all flowers of "Epic" F1, "Flavine" F1, and "Gascona" F1, respectively. This flower phenotype contained the highest number of pollen tubes in the style and the highest number of fertilised ovules. The biostimulant had a positive effect on the flower and fruit set numbers, as well as on the pollination efficiency in all genotypes. *Ascophylum nodosum* extract could be used as an efficient stimulator of flowering and fruit setting for eggplant hybrids in field conditions in a temperate climatic zone.

**Keywords:** *Ascophyllum nodosum*; *Solanum melongena*; heterostyly; pollination efficiency

#### **1. Introduction**

Eggplant is a photoperiodically inert plant with bisexual and partially self-pollinating flowers, although cross-pollination increases the effectiveness of fruit setting [1].

The downward-facing flowers are born solitary or in clusters. The eggplant produces three types of flowers: With a long-style pistil, where the stigma is localised above the anthers; with a medium-style pistil, where the stigma is at the same level as the anthers; and with a short-style pistil, where the stigma is below the anthers (Figure 1). This flower character promotes outcrossing between morphs via delivery and uptake of pollen by pollinators [2,3]. The stamen pores of the long- and medium-styled pistils are localised above or close to the stigma, favouring self-pollination. On the contrary, the stigmas of short-styled pistils are inside the downward-facing anther cone, making self-pollination difficult [4–6].

**Figure 1.** Stylar heteromorphism in eggplant: The flower with long-styled pistil of "Flavine" F1 (**a**), medium-styled pistil of "Gascona" F1 (**b**), short-styled pistil of "Epic" F1 (**c**).

Anthers are ready to release pollen and the stigma is receptive from the first opening of the flower (Figure 2). Stigma receptiveness gradually decreases with the plant's age, and by the fifth day of flowering, receptiveness is negligible, and the stigma turns brown [2].

**Figure 2.** Stigma in eggplant pistil with visible papillae in a receptive phase, (**a**), and pollen grains (**b**) in "Epic" F1 by scanning electron microscopy.

All types of flowers are found in the same plant and even within the same cluster. The expression of heterostyly depends on the plant genotype, the age of the plant, fruit load, environmental conditions, and growing practices. Generally, domination by long-styled flowers has been reported, amounting to 50–100% of all flowers [7–9]. The higher fruit setting efficiency of this phenotype results from well-developed nodules with high pollen absorption capacity. However, the development of the ovules and their position in the placenta, as well as pollen grain shape, size, and amount in anthers, were nondifferentiated among long-, medium-, and short-styled pistils [6,10], although Wang et al. [11] demonstrated that lower fruit setting from short-styled flowers resulted from stigma-pollen incompatibility. The bumblebee (*Bombus terrestris*) is the most effective eggplant pollinator for plants under covers. Yield increase and better fruit quality are considered to be the major benefits of bumblebee application as compared to self-pollination or inflorescence vibrating [8,12]. Optimisation of eggplant yield in unfavourable conditions could also be achieved by introducing the cultivation of parthenocarpic cultivars [13]. Pollination leading to fruit and seed formation is associated with the production of endogenous growth regulators such as auxins. In this respect, the use of fruit-setting using auxin-based growth regulators has also been recommended to enhance fruit setting under suboptimal temperatures [5,14]. Investigations on the control of eggplant flowering through growth regulators have been successively performed since the end of the 20th century, but their results have been inconclusive [15,16]. Eggplant tolerance to biotic and abiotic stresses can be

managed through grafting. The effects of rootstock/scion combinations on eggplant performance were investigated in terms of yield and fruit quality [17,18]. It can be assumed that this technique affects the flowering biology as well, but this issue needs future investigation. In Poland, eggplants are cultivated mainly under unheated foil covers from spring to autumn. To lower costs, cultivation is also performed in open fields where air temperatures may fall below the optimum, causing a reduction in flowering and fruit setting [19]. A promising way to control eggplant generative development could be biostimulant application. Biostimulants have been a focus of global interest of the scientific community since the end of the 20th century, giving promising results in different branches of agriculture as stimulators of crops growth, stress tolerance and yield [20,21]. Seaweed extracts (SWE) are among the main biostimulants, recognised as nontoxic, nonpolluting and nonhazardous to various organisms [22,23]. The majority of the SWE formulations are based on the extract of the brown algae *Ascophyllum nodosum* (L.) Le Jolis. Although seaweed extracts are heterogeneous in nature, the leading companies standardise their chemical composition to ensure consistent product quality [24,25]. Some authors have reported the stimulatory effect of seaweed extracts on eggplant yield [26,27], but there are no references on the flowering biology of this species as affected by SWE biostimulation. SWE action is extremely complex, but interdisciplinary investigation of biostimulant vs plant interactions may shed new light on the effective utilisation of these promising bioproducts in horticulture.

We hypothesise that seaweed extract affects the flowering and fruit setting of eggplant in a multidirectional manner. The reaction of plants to biostimulant treatment depends on the flowering biology of the cultivars, particularly the proportions of different flower phenotypes and their fertility. We aimed to investigate the flowering biology of three eggplant hybrids treated with seaweed extract Göemar BM-86® (Arysta LifeScience North America, LLC) to determine the crucial mechanisms behind the final effect of increased yield.

#### **2. Materials and Methods**

#### *2.1. Experimental Arrangement*

A two-factorial experiment was set up using randomised blocks in three replications, in the years 2013 and 2015, at the University of Agriculture in Krakow, Poland. The investigated eggplant hybrids, "Epic" F1 (Seminis Vegetable Seeds), "Flavine" F1 (Gautier Semences), "Gascona" F1 (Gautier Semences), were selected on the basis of preliminary studies evaluating their performance in field cultivation under temperate climate conditions [26–28], determined by the earliness, vigour, and yield potential of those plants. Biostimulant Göemar BM-86® (Arysta LifeScience North America, LLC) was applied three times in two week intervals as a foliar application, in a dose of 1.5 dm<sup>3</sup> ha−1. Control plants were sprayed with distilled water. Goemar BM 86® is standardised *Ascophyllum nodosum* (L.) Le Jolis extract, which provides a constant and balanced formulation containing (in %): N, 5.0; Mg, 2.4, S, 3.2, B, 2.07; and Mo, 0.02 [29].

#### *2.2. Cultivation Procedures*

Eggplant seeds were sown on 1 March 2013 and 3 March 2015 in seed boxes. After three weeks, the seedlings with one fully developed leaf were transplanted into black 40-cell multipots (VEFI, Norway) with a single cell volume of 0.23 dm3. Seedlings were grown in a greenhouse, in temperatures of 20/17 ± 2 ◦C day/night. The growing medium was peat substrate KlasmanTS2 (Klasmann-Deilmann GmbH, Geeste, Germany). The foliar fertiliser Kristalon Green (Yara, Szczecin, Poland) was applied twice in a dose of 10 g dm−<sup>3</sup> water during seedling production. A gradual decrease in temperature and irrigation was used for the hardening of seedlings seven days before being transplanted to the experimental field (50◦04 N, 19◦51 E) on 7 May 2013 and 15 May 2015, with spacing of 0.75 × 0.60 m. Experimental plots covered 15 plants per treatment for observations of flowering and fruit setting and an additional 15 plants per treatment for flower collection for microscopic observations. Plots were surrounded by shelterbelts. The soil of the experimental field was Fluvic Cambisol (Humic) according

to the FAO (Food and Agriculture Organization of the United Nations) classification with a Corg level of 2% and pHKCl 6.11. Before the field experiment was established, the soil samples were analysed, and doses of fertilisers were applied to achieve a stable content of nutrients (in mg dm3): N, 100; P, 90; K, 220; Ca, 1,100; Mg, 70. Cultivation procedures of weeding, irrigation, and plant protection were performed according to the standard recommendations for eggplant cultivated in field conditions in Poland, described by S ˛ekara [30].

#### *2.3. Weather Conditions*

The climate of the experimental station is humid continental (Dfb) according to the Köppen's classification. Detailed data concerning the mean air temperature, photosynthetically active radiation (PAR), and the total rainfall during the vegetation seasons in 2013 and 2015 are presented in Table 1. Data were collected from automatic HOBO Pro RH/Temp loggers to assess temperature and a HOBO Weather Station (Onset Comp. Corp., Cape Cod, USA) to assess light characteristics and rainfall at the experimental site. The growing season in 2015 was generally warmer than that in 2013 regarding mean monthly temperatures, with the exception of June. In 2015, precipitation was distributed evenly, while in 2013, 45% of rainfall was recorded in June. A cool September in both years and low PAR caused a continuous decline in eggplant yield (Table 1).


**Table 1.** Mean monthly temperature, photosynthetically active radiation (PAR) and sum of rainfall in vegetation seasons 2013 and 2015.

#### *2.4. Procedures for Flowering and Fruit Setting Observations*

The observations were conducted on 5 plants per replication (15 plants per treatment and cultivar) during the flowering period, from June to September. Single flowers were labeled according to the order of appearance on each plant. The numbers of flowers of particular phenotypes (with long-styled, medium-styled, or short-styled pistil) were recorded after the opening of petals. Then, the number of fruits set from flowers of a particular phenotype was also recorded about one week after fertilisation, when fruit sets reached 1–2 cm in diameter. Flowers which did not set fruits were naturally aborted. Fruits in a stage of harvest maturity were picked to reflect the standard cultivation conditions and to exclude excessive metabolite sink by ripening fruits. Data were calculated and presented as a sum of flowers and fruits per plant per month for the two experimental years separately.

#### *2.5. Procedures for Microscopic Observations*

At full flowering, 20 pollinated flowers of each phenotype per treatment and cultivar were collected in 2013 and 2015. Data are presented as a sum of observations for investigated seasons, *N* = 40. The styles were isolated and fixed in FAA (formalin-acetic-alcohol), according to Martin's method [31] adapted by S ˛ekara [30]. The germination of pollen on stigmas, growth of pollen tubes, and fertilisation of ovules were examined under fluorescence microscopy with the use of SteREO LUMAR V12 microscope (Carl Zeiss AG, Jena, Germany) (Figure 3). The number of pollen tubes in half of the style and the number of fertilised ovules were evaluated. The numbers of pistils having a number of pollen tubes in the ranges 0–100; 100–200; 200–300; 300–400; 400–500; 500–600; 600–700; 700–800, 800–900; 900–1000 were determined. For fertilised ovules, the following ranges were included: 0–50; 50–100; 100–150; 150–200; 200–250; 250–300; 350–400.

**Figure 3.** Germination of the pollen on stigmas (**a**) and fertilisation of ovules (**b**) of eggplant observed under fluorescence microscopy after Martin's aniline blue fluorescence technique.

#### *2.6. Statistical Analyses*

Statistical analyses were performed using the Statistica 12.0 software package (StatSoft Inc., Tulsa, OK, USA). A three-way analysis of variance followed by Tukey's honest significance test was used to determine the main effects of the type of flower, biostimulant, and time of sampling, as well as interactions between main effects, at the *p* ≤ 0.05 significance level. Data shown in the tables and figures are averages of three replicates.

#### **3. Results**

In the conditions of the present experiment, the eggplants started flowering at the beginning of June, while the period of the most intensive flowering fell in August. The investigated hybrids showed flower heterostyly—the presence of long-, medium-, and short-styled flowers was observed for all investigated plants. Moreover, heterostyly expression significantly depended on biostimulant treatment and the age of the plants (Table 2).


**Table 2.** Chosen aspects of flowering and fruit setting of eggplant as depended on fruit type and biostimulant treatment.


**Table 2.** *Cont*.

\* C, control; B, biostimulant; \*\* Means within rows, followed by different letters, are significantly different at *p* ≤ 0.05, *N* = 3. Comparisons were performed with the use of Tukey's honest significance test.

Among 23 flowers set by "Epic" F1 plant during one vegetation period, 55% had long-style pistils, 30% had medium-style pistils, and 15% had short-style pistils. "Epic" F1 plants produced 7 fruits during the vegetation season, on average; 57% of these were from long-styled flowers, 26% from medium-styled flowers, and 17% from short-styled flowers. Biostimulant treatment significantly increased the number of only medium-styled flowers in 2013 and the number of fruits set by long- and medium-styled flowers in both vegetation periods. The most effective in fruit setting were long-styled flowers. The biostimulant positively affected the percentage of fruits set by all flower phenotypes and the number of seeds, with the exception of flowers with short-styled pistils in 2013. The first fruits were collected at the end of July. The highest number of long- and medium-styled flowers was observed in August; the lowest was observed in September (Figure 4, Table 3).

**Figure 4.** The course of flowering and fruit setting of "Epic" F1 eggplant as depended on fruit type and biostimulant treatment. C, control; B, biostimulant. Bars represent mean number of flowers per plant in 2013 (**a**), 2015 (**b**) and fruits per plant in 2013 (**c**), and 2015 (**d**) (error bars indicate SE).


**Table 3.** Results of ANOVA for parameters of flowering and fruit setting of "Epic" F1 eggplant presented in Figure 4.

Levels of significance for ANOVA: \* *p* ≤ 0.05; \*\* *p* ≤ 0.01; \*\*\* *p* ≤ 0.001; ns, not significant; *N* = 3. Comparisons were performed with the use of Tukey's honest significance test.

The number of short-styled flowers increased in line with aging of the plants. The number of fruits set from long- and medium-styled flowers increased from July to August, then decreased in September. We observed, on average, 106 pollen tubes in the short-styled pistils, 422 in medium-styled pistils, and 610 in long-styled pistils collected from control plants and 129, 490, and 778 pollen tubes, respectively, collected from biostimulant-treated plants (Figure 5). The ovaries of the short-styled flowers contained approximately 36 fertilised ovules, and more fertilised ovules were found in the remaining types of flowers: 199 and 225 in medium- and long-styled flowers, respectively, produced by control plants. The flowers of biostimulant-treated plants contained 39%, 32%, and 36% more fertilised ovules in the ovaries of short-, medium-, and long-styled flowers, respectively.

**Figure 5.** *Cont*.

**Figure 5.** Number of pollen tubes (**a**) and fertilised ovules (**b**) in the styles of different flower types in "Epic" F1.

The "Flavine" F1 plants produced 22 flowers during the vegetation period; 41% had long-styled pistils, 38% had medium-styled pistils, and 19% had short-styled ones (Table 2). The number of fruits collected from a plant was 6 on average; 42% of these were from long-styled flowers, 34% from medium-styled flowers, and 24% from short-styled flowers. Biostimulant treatment significantly increased the number of long-styled flowers in 2015, the numbers of medium-styled flowers in 2013 and 2015, and the numbers of fruits set by long- and medium-styled flowers in 2015. The most effective in fruit setting were long-styled flowers. The biostimulant positively affected the percentage of fruits set by all flower phenotypes and the number of seeds in fruits born by long-styled flowers in 2015 and by medium-styled flowers in both years of the experiment. The highest number of long- and medium-styled flowers was observed in August; the lowest was in September (Figure 6, Table 4). The number of fruits set from long-styled flowers was the highest in August. We observed, on average, 149 pollen tubes in the middle of the style in the short-styled flowers of control plants, 410 in medium-styled flowers, and 595 in long-styled ones (Figure 7). In biostimulant-treated flowers, a 23% higher number of pollen tubes was observed in short-styled pistils, 16% higher in medium-styled pistils, and 20% higher in long-styled pistils. The ovaries of the short-styled flowers collected from the control plants contained, on average, 54 fertilised ovules; more fertilised ovules were found in the remaining types of flowers: 208 and 243 in medium- and long-styled flowers, respectively. The numbers of fertilised ovules in analogous types of flowers collected from biostimulant-treated plants were 102%, 24%, and 23% higher, respectively.

**Figure 6.** *Cont*.

**Figure 6.** The course of flowering and fruit setting of "Flavine" F1 eggplant as depended on fruit type and biostimulant treatment. C, control; B, biostimulant. Bars represent mean number of flowers per plant in 2013 (**a**), 2015 (**b**) and fruits per plant in 2013 (**c**), and 2015 (**d**) (error bars indicate SE).

**Table 4.** Results of ANOVA for parameters of flowering and fruit setting of "Flavine" F1 eggplant presented in Figure 6.


Levels of significance for ANOVA: \* *p* ≤ 0.05; \*\* *p* ≤ 0.01; \*\*\* *p* ≤ 0.001; ns, not significant; *N* = 3. Comparisons were performed with the use of Tukey's honest significance test.

**Figure 7.** *Cont*.

**Figure 7.** Number of pollen tubes (**a**) and fertilised ovules (**b**) in the styles of different flower types in "Flavine" F1.

"Gascona" F1 plants produced 18 flowers during the vegetation period, with 42% of these being long styled, 38% medium styled, and 20% short styled (Table 2). The number of fruits collected from a plant was 6, on average; 46% of these were from long-styled flowers, 34% from medium-styled flowers, and 20% from short-styled flowers. Biostimulant treatment significantly increased the number of medium-styled flowers in 2013 and 2015 but did not affect the number of fruits. The most effective in fruit setting were long-styled flowers. The biostimulant positively affected the percentage of fruits set by long-styled flowers in 2015 and medium-styled flowers in both years, but it negatively affected the effectiveness of fruit setting by short-styled flowers. Biostimulant treatment positively affected the seed number (Table 2). The highest number of long-, and medium-styled flowers was observed in August; the lowest was in September (Figure 8, Table 5). The number of fruits set from long- and medium-styled flowers increased from July to August, then decreased in September. Long-, medium-, and short-styled flowers were analysed regarding the number of pollen tubes in the styles. Differences in the course of pollination and fertilisation between investigated cultivars concerned the number of pollen tubes and fertilised ovules in the pistils. For control "Gascona" F1 plants, we observed, on average, 119 pollen tubes in the middle of the style in the short-styled flowers, 418 in medium-styled flowers, and 595 in long-styled ones (Figure 9). The ovaries of the short-styled flowers contained approximately 0–50 fertilised ovules, while more fertilised ovules were found in the remaining types of flowers: 200–400 in medium- and long-styled flowers. The flowers of control plants contained lower numbers of both pollen tubes and fertilised ovules in all types of flowers.

**Table 5.** Results of ANOVA for parameters of flowering and fruit setting of "Gascona" F1 eggplant presented in Figure 8.


Levels of significance for ANOVA: \* *p* ≤ 0.05; \*\*\* *p* ≤ 0.001; ns, not significant; *N* = 3. Comparisons were performed with the use of Tukey's honest significance test.

**Figure 8.** The course of flowering and fruit setting of "Gascona" F1 eggplant as depended on fruit type and biostimulant treatment. C, control; B, biostimulant. Bars represent mean number of flowers per plant in 2013 (**a**), 2015 (**b**) and fruits per plant in 2013 (**c**), and 2015 (**d**) (error bars indicate SE).

**Figure 9.** *Cont*.

**Figure 9.** Number of pollen tubes (**a**) and fertilised ovules (**b**) in the styles of different flower types in "Gascona" F1.

#### **4. Discussion**

#### *4.1. Heterostyly Expression in Eggplant as A*ff*ected by Biostimulant Treatment and Cultivar*

The recent research aimed to develop an overview of the heterostyly phenomenon in eggplant, and its implications on fruit setting biology. We demonstrated the presence of three phenotypes of flowers and the differentiated fertility of them, specific to the investigated hybrids. Generally, long-styled flowers dominated, but the fruit setting efficiency was not directly determined by the flower phenotype. A study by Srinivas et al. [32] indicated that for two eggplant hybrids of Indian breeding, with long, green and round, purple fruits, 80% of fruits were set by long-styled flowers, whereas 20% of the fruits were set by medium-styled flowers and no fruit by short-styled flowers. The partial sterility of short-styled flowers demonstrated in the cited research was due to small stigmas with under-developed papillae on which pollen grains failed to germinate. The short-styled flowers of the eggplant hybrids which are the subject of the present investigations were fertile, although the lowest number of pollen tubes and fertilised ovules was observed in this flower phenotype. Despite this fact, "Epic" F1 and "Gascona" F1 plants set about 20% of fruits from short-styled flowers. For "Flavine" F1, the percentage of fruits set by this mentioned flower phenotype was 30%. S ˛ekara and Bieniasz [6] determined that the ovules of short-styled pistils were typically developed, but that their fruit setting efficiency was low. On the contrary, results by Hazra et al. [33] indicated full sterility of short-styled flowers due to some problem related to ovary development. Observations with the use of a fluorescence microscope allowed us to verify the correct growth of pollen tubes in the styles of all types of pistils but their number was significantly affected by flower type and biostimulant treatment and by cultivar to a lesser extent. This observation is contrary to the results of Wang et al. [11], who determined that the structure of the stigmatic surface in short-styled flowers inhibited pollen germination. On the grounds of highly genotype-dependent heterostyly expression in eggplant, results on short-styled pistil performance may be divergent.

Application of Göemar BM-86® caused an increase in the numbers of pollen tubes and fertilised ovules. This phenomenon was common for all types of flowers and is directly attributable to pistil characteristics. Biostimulant-treated and control plants were not isolated, so they both could act as pollen donors. The effect of biostimulants on pollen production and fertility should also be an object of future research. Based on the available literature, we can only conclude that a wide pool of bioactive seaweed extract compounds provided balanced development and enhanced the flowering and fruiting of the investigated eggplant hybrids. The biostimulant-treated plants could be able to develop a better canopy for effective interception of light and—through a significant reduction in interplant competition for solar energy and nutrients—build suitable carbohydrate reserves earlier. Such mechanisms beyond increased flowering and fruit setting in seaweed extract treated plants were proposed by Arthur et al. [34] for bell peppers and by Helaly et al. [35] for tomatos.

The increasing number of short-styled flowers in line with plant aging, in the conditions of the present experiment, could be the result of increasing fruit load with the vegetation season's flow. Having well-developed anthers, short-styled flowers act as pollen donors to provide reproductive success. Araméndiz Tatis et al. [9] demonstrated that short-styled flowers of the "Lilac" eggplant landrace and "Long Purple" increased male fitness and thus produced an imbalance in functioning between male and hermaphrodite flowers. According to Khah et al. [36], fruit load negatively affected style length but not anther cone length in eggplant, even under favourable climatic conditions. The investigated hybrids could reduce energy outlines by creating flowers with reduced pistil and decreased fertility at the end of the vegetation period, but do so while producing pollen in the normally shaped anthers, promoting male behaviour. Short-styled flowers could be borne by fruit-loaded plants as a source of pollen for insects. The construction of the eggplant stamens is an expression of adaptation to pollination through vibrations. Such an adaptation limits the potential pollinators to species that are able to introduce vibrations into anthers, including bumblebees [13]. Bumblebees commonly visited eggplant flowers in the conditions of the presented experiment.

#### *4.2. Biostimulant-A*ff*ected Flower and Fruit Set E*ff*ectiveness*

Biostimulants have shown promising results in promoting flowering and reducing the fruit drop agents in many fruit trees, like apple, avocado, clementine, orange, olive, and pomegranate [37–40]. In this respect, seaweed extracts enriched in microelements are the most effective [41]. Vegetables with edible fruits are characterised by competition between the flowers and fruits at different stages of growth and in different positions in relation to inflowing assimilates [42]. Dropping of flowers, typical for eggplant, could be the mixed effect of lack of pollination or limited inflow of assimilates and the phenomenon of domination of fruit producing growth regulators. In the present research, hybrids treated with seaweed extract bore more flowers and fruits than did untreated ones. More intensive flower setting was elicited either by improved plant growth through seaweed extract application or by endogenous components, especially cytokinins, which enhance nutrient partitioning in vegetative plant organs and increase in the transport of assimilates to the growing fruits. A similar effect was observed for eggplant treated with seaweed extract by Abd El-Gawad and Osman [43]. Under the conditions of the presented experiment, biostimulant application also increased the number of pollen tubes and fertilised ovules in all types of flowers of the investigated cultivars. The overall positive influence of seaweed extracts on the plants resulted in better reproductive effectiveness and increased fruit yield and quality, described in detail by Pohl et al. [26,27]. Gómez-Cadenas et al. [44] investigated the effect of a biostimulant product containing macronutrients on citrus fruit set abscission. The beneficial effects of the biostimulant resulted from an increase in the photosynthetic efficiency which led to better transport of carbohydrates from leaves to fruit sets. Seaweed-treated apple trees also showed higher leaf chlorophyll contents and increased rates of photosynthesis and respiration due to treatment decreasing the oscillations in yield between "on" and "off" years and increasing the average fruit weight on plants affected by too high a crop load [45]. Pollination and fertilisation are very stress-sensitive stages of development [1]. Based on research on tomato, low temperatures, especially during the night, are not detrimental to ovule development but could affect stigma and style function [5]. Pollen viability is the highest at 20–22 ◦C [14], while the mean temperatures for the flowering period (July–September) were 16.8 and 18.8 ◦C in 2013 and 2015, respectively. Pollen development and viability depend on carbohydrate supply [5], so the increased photosynthetic performance of biostimulant-treated plants could improve sugar partitioning to developing pollen grains. The bioactive compounds of seaweed

extracts enhance the tolerance of eggplant to abiotic stresses [46] and this tolerance can also cover the generative reproduction of this crop in temperate regions.

#### **5. Conclusions**

Eggplant is a warm climate crop and is cultivated for fruits, widely used in many world cuisines because of their unique taste and dietetic value. Nonoptimal growing conditions, especially in temperate climatic zones, affect plant flowering and fruit setting. Biostimulant application in the experiments presented herein affected the flowering biology of eggplant cultivars in different ways. Generally, the biostimulant positively affected the percentages of the most fertile medium- and long-styled flowers and the effectiveness of fruit setting by all flower phenotypes. Increased numbers of pollen tubes and fertilised ovules in all types of flowers of the investigated cultivars were noted. The overall positive influence of Göemar BM-86® on the plants resulted in increased reproductive effectiveness. Biostimulant application seems to be a promising solution to improve eggplant flowering and fruit setting in unfavourable growing conditions.

**Author Contributions:** Conceptualization, A.P. and A.S.; methodology, A.P. and A.S.; software, A.P. and A.S.; validation, A.P. and A.S. and A.K.; formal analysis, A.P.; investigation, A.P. and A.G.; resources, A.P. and A.S.; data curation, A.P. and A.S.; writing—original draft preparation, A.P. and A.S.; writing—review and editing, A.P. and A.S. and A.K.; visualization, A.P. and A.S.; supervision, A.S.

**Funding:** This research was funded by the Ministry of Science and Higher Education of the Republic of Poland.

**Conflicts of Interest:** The authors declare no conflict of interest.

#### **References**


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