Figure 7 shows the variations of saponin contents for processed seeds in response to several pearling durations. According to statistical analysis, saponin content in processed seeds was affected only by pearling duration; no significant difference was recorded between varieties. However, the interaction between variety and pearling duration factors was significant, which indicates that the tested varieties responded differently to the pearling process. For instance, the ICBA-Q5 variety responded very well to pearling, as it had the highest saponin content in raw seeds (0 min) and the lowest in processed seeds (6 min), and consequently the highest saponin removal.

**Figure 7.** Variations of saponin contents in processed seeds in response to pearling duration. Error bars indicate the standard deviation. Treatments without a common letter are significantly different at *p* < 0.05.

#### *2.4. Pearson Correlation*

Pearson's correlation analysis was conducted for all investigated nutrients in processed seeds and seed bran separately, by correlating all the physicochemical parameters (productibility and growth) with the varieties used. The obtained results are shown in Figure 8.

**Figure 8.** Pearson correlations among all chemical parameters for processed seeds (**a**) and seed brans (**b**).

The analysis revealed a strong positive correlation between groups of parameters G1 (N, protein content) and G2 (Na, Mg, Mn, Ca, ash content, and K). Strong negative correlations were found between moisture and P, saponin, and time. On the other hand, no correlation between time, Mn, K, saponin, Ca, and K was observed (Figure 8a).

For the obtained seed brans (Figure 8b), a strong correlation was found between N, protein content, and total fat. A strong negative correlation was observed between time, ash content, K, moisture, P, and S. No correlation between moisture and Ca was observed.

#### *2.5. Principal Component Analysis (PCA)*

Results of PCA (Figure 9) indicate that the first two components represent over 69% of the variability. PCA axis 1 was largely determined by B, Mn, S, Ca, Na, and K. Among them, B and Ca had the highest positive correlation (r = 0.927–0.975). Variation along PCA axis 2 was highly determined by P, fat content, N, and protein content.

The PCA graph of variables showed that the pearling duration was correlated negatively with saponin, moisture, Ca, Na, Mg, and Ca. On the other hand, this key parameter was correlated positively with the rest of the parameters.

**Figure 9.** PCA analysis of investigated parameters for processed seeds.

#### **3. Discussion**

Since its introduction to Morocco in the 2000s, Quinoa has been seen as a rustic and stress-tolerant crop with several potentialities to replace cereals and other traditional crops in marginal environments, mainly those affected by drought and salinity. A deep understanding of the nutritional profiling of quinoa and the high amount of saponin reveal the importance of processing in order to remove saponins and produce a palatable seed. Regarding the morphological aspects of quinoa seeds, the quinoa seed is disc-shaped, approximately 2 mm in diameter, and 0.5 mm in thickness. The size of a quinoa seed depends on several factors, such as quinoa cultivar and abiotic conditions (climate, soil, etc.). In fact, the whole quinoa seed is divided into three main structural components: perisperm, embryo, and bran. The bran is defined as the out layer that surrounds the embryo. The embryo entirely covers the perisperm, the innermost layer, as a headband. The bran is eliminated during abrasion (manual or mechanical) because it contains 86% saponins compared to other grain fractions [29]. The embryo and perisperm contain 11% and 3% of saponins, respectively [29].

The special structure of quinoa seeds requires special scarification methods, and the dehulling rates change when milled for the same rate of duration using different equipment [30]. Hence, it is necessary to have a balance between the pearling duration and the nutritional contents of quinoa. For all used varieties, the thousand-seed weight of quinoa seeds decreased with increase pearling duration due to the removal of the external layer of quinoa [31]. Some research works have mentioned that the average diameter can reveal some quinoa characteristics, such as hardness coefficient and saponin content. Otherwise, the higher the hardness coefficient of quinoa, the more saponins are present [32]. The average thousand-seed weight was negatively correlated with the pearling duration. Our finding indicates that after 2 min of mechanical abrasion, a high percentage of saponins were removed.

There are two main processes of saponin removal. The first one is a wet method based on washing. It increases seed moisture, which then requires drying to prevent mold growth, and enhances the seed's shelf life (while raising the cost of seed processing) [10]. The second method, which is the commonest one, is a dry method based on abrasion that removes the outer bran layer by mechanical scarification. Often, processing operations involve a preliminary washing step following abrasion to further reduce the level of saponins [33]. Nevertheless, for some bitter quinoa varieties that contain higher amounts of saponins, if a wash operation is skipped, the embryo may be damaged as the next outer layer after the bran, which is totally eliminated [34]. Quinoa's embryo is the richest nutritional source; it contains 57% of the protein, 49% of the fat, 20% of the sugar, 45% of the dietary fiber, and 51% of the ash of the whole grain [29]. Hence, removal techniques that keep the embryo intact preserve the complete nutritional profile of quinoa. Abrasion alone after 30 s reduced saponin levels from 1.19% to 0.45%. However, after an additional 2 min, the saponin level was reduced slowly to 0.21% [33]. Our results showed the same tendency of reducing saponins for the three studied varieties. Two minutes of mechanical abrasion was enough to reduce saponins below 0.12% which is the accepted threshold for quinoa consumption [35]. On the other hand, washing alone augmented the saponin percentage of oleanolic acid, while diminishing phytolaccagenic acid. Although abrasion decreased the percentage of oleanolic acid, it increased the percentage of phytolaccagenic acid [33]. These variations are correlated with the chemical structure of the specific saponin and the location within the bran layer. Furthermore, hydrating the quinoa without manual abrasion increased the saponin levels from 3.3% to 3.6% [32]. The mechanical polishing permitted a reduction of saponins from 2% to 0.4% for Titicaca variety, and from 1.4% to 0.5% for Puno, decreases of 80% and 64% of the initial saponin respectively [36].

The protein contents of the studied quinoa cultivars were equal to 14.4%, 12.8%, and 7.9% (dry weight basis) for Puno, Titicaca, and ICBA-Q5, respectively. Puno and Titicaca's protein contents were consistent with the protein content (9.5–15.7%, dry weight) reported by Nowak Verena [37]. However, protein content for each variety did not change during the mechanical abrasion. The protein loss increased in terms of seed weight when the pearling duration was higher (after 4 min of mechanical abrasion). The decrease in weight can be explained by the fact that the protruding scutellum of the pericarp of quinoa was scoured away [38,39]. Furthermore, when pearling duration was greater than 6 min, protein loss became greater, since it is primarily present in an embryo, which accounts for 57% of total quinoa protein [37]. Our finding indicates that the embryo and endosperm were milled at this stage of the pearling process.

The fat contents of quinoa cultivars were equal to 3.7%, 4.5%, and 2.2% (dry weight basis) for Puno, Titicaca, and ICBA-Q5, respectively. All contents fit within the range of the total lipid content of quinoa [40].  Moreover, we noticed a decrease in fat content after 6 min of mechanical abrasion, as the bran was removed and the embryo was ground. The obtained results are aligned with another study showing that a dehulling rate of 8.6% can decrease fat content by 7.6% [41].

The ash contents for the three studied varieties fit within the range reported in the literature [40]. The ash content of treated seeds decreased with the increase of the pearling duration. After 2 min of mechanical abrasion, the ash content decreased sharply for all studied varieties. The decrease of ash content is explained by the fact that the pericarp, where most of the minerals are mostly concentrated, was removed. On the other hand, the seed bran obtained from two minutes of mechanical abrasion is highly rich in minerals, and the ash contents were equal to 18.0%, 17.3%, and 16.7% (dry weight basis) for Puno, Titicaca, and ICBA-Q5, respectively. In another study, Ando et al. [29] reported that the ash content was about 9.2% (dry weight basis) in the bran after pearling. That is relatively lower than the content found in this work, which can be explained by the use of different pearling processes and equipment.

A suitable pearling duration is the one that results in saponin content lower than the threshold required from the CODEX standard [35]. Thus, two minutes of mechanical abrasion was the best pearling duration. In this study, we found that K was the most abundant macroelement for Puno, Titicaca, and ICBA-Q5. The remaining macroelements, by abundance, were P, followed by Mg, Ca, S, and then Na. Regarding micronutrients, Fe was the most abundant element with concentrations in raw seeds of 55.2 mg/kg for Puno, 61.8 mg/kg for Titicaca, and 36.81 mg/kg for ICBA-Q5. Our findings in this regard are in agreement with results reported by many other studies where K and P were the dominant macroelements, followed by Mg and then Ca. Fe was the dominant micronutrient [29,36,42]. The quinoa cultivated in Morocco is characterized by higher amounts of K, Ca, and Mg compared to other studies focusing on mineral profiling of quinoa seeds [29,42,43]. Mhada et al. [36] explained that these high contents are mainly due to calcareous soil rich in K, Ca, and other elements.

Our results suggest that protein and mineral contents were not affected by pearling, which indicates that protein and most of minerals are mostly located in the endosperm and embryo, which confirm the finding reported by Prego et al. [44]. According to Konishi et al. [45], the P, K, and Mg are located in embryonic tissue. In particular, P's origin is attributed to phytic acid, and the origins of Mg and K to phytate. The same authors also reported that K and Ca were present in the pericarp [45]. Furthermore, they found that Ca is scarcely found in embryonic tissues of quinoa seeds. It occurred mostly in the pericarp and seed coat, as well as the boundary between perisperm and embryo. It is associated with carboxyl groups of pectin molecules in the cell wall to form Ca-pectin complexes [44], which confirms the finding of this study that showed decreases in Ca of 58%, 64%, and 14% for Puno, Titicaca, and ICBA-Q5, respectively. Moreover, the decreases in K (56%, 56%, and 16% for Puno, Titicaca, and ICBA-Q5, respectively) can be explained by the fraction located in the pericarp; the K fraction located in the embryo was not eliminated. According to Ando et al. [29], milled quinoa had lower K, Mg, and Ca, and higher P. Nevertheless, the authors also found that Fe, Zn, Cu, and Mn contents remained stable [29]. This disagreement may be explained by some differences in terms of genetic material, since they worked with another variety (quinoa Real) and a lower degree of polishing by the pearling machines used for seed processing [29,36]. According to a previous study conducted with the same pearling machine, the authors found that the pearling process had no effect on macronutrients and protein content, except for Ca, which declined by 28% for Puno variety [10,11]. However, micronutrient contents were significantly reduced. The pearling process resulted in significant losses for most micronutrients—by 60%, 57%, 2%, and 6% for Cu, B, Fe, and Zn, respectively; however, there was no significant reduction in terms of Mn [10,11].

During the pearling process, an important portion of the nutrients is wasted in the produced seed bran; therefore, its valorization is essential. All seed brans generated from the three studied varieties are rich in nutrients compared to the raw seeds. This is explained by the fact that the pericarp has high mineral content, as all produced seed brans contain high ash content. In fact, the research findings demonstrate the richness of quinoa seed brans. However, their valorization will mainly depend on several parameters (saponin contents, microelement contents, and microelement contents), and their usage scope is vast: they can be valorized in food, cosmetic, pharmaceutical, or agronomic industries. Recently, many studies were conducted to valorize saponins in different sectors. For instance, the effect of saponins on pests was studied. Some recent studies suggested the implementation of saponins as an ingredient for the storage of food and grains in order to preserve their sanitary quality. In fact, saponins rupture the internal mucous cells of pest's intestines by forming complexes with digestive enzymes such as proteases [46]. According to Zegarra [47], the acaricidal activity was evaluated from saponins present in bitter quinoa varieties. The author indicated that the Markjo variety shows bioactivity close to a commercial acaricide product. Furthermore, saponins have a tremendous inhibitory effect on fungi such as botrytis cinerea [48]. Quinoa saponins and polyphenols have high antioxidant effects and free radical scavenging activities, as well as an anti-inflammatory effect [49]. Drugs that contain saponins improve the body's immune response significantly through the enhancement of absorption and their hemolytic properties. Saponins grabbed worldwide attention due to their anti-carcinogenic characteristics and cholesterol-lowering effects [50]. In food industries, saponins have a positive effect on the physical stabilization of oil/water (O/W) emulsions. Moreover, they contribute to preserving liquids from oxidation, and they can be incorporated as foaming agents in beverage industries [50,51].

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

#### *4.1. Seed Material*

Quinoa seeds (*Chenopodium quinoa* Willd) grown in Berrechid, Morocco (2018 harvest) were used in this experiment. Seeds of three varieties of quinoa (Puno, Titicaca, and ICBA-Q5) were used after threshing and winnowing operations to study the effects of several pearling durations on seed nutritional and anti-nutritional properties. Quinoa seeds were not unisized before the pearling process, since it is not important to ensure this additional postharvest operation as well as simulating the real conditions that will be adopted by farmers.

#### *4.2. Quinoa Pearling Machine Design*

The pearling machine used in the experiment was locally manufactured by Benrim farm (Figure 10). It is equipped with two motors. The first one is designed to turn a drum with a rotation speed of 750 rpm. The second one is more powerful (3000 rpm) and designed to extract the fine dust produced during the pearling process. The rotating drum is made of an 80 cm long perforated inox steel and has six baffles distributed throughout the drum.

**Figure 10.** Design of the quinoa pearling machine used in the present experiment.

#### *4.3. Pearling Treatments and Machine Operating Mode*

The machine operates on a semi-industrial scale—a minimal quantity of 8 kg per sample is required. In this trial, five pearling durations were tested (0, 2, 4, 6, 7, and 8 min) for the three selected varieties; 0 min corresponds to raw seeds. Each combination pearling duration x variety was repeated 3 times. For each duration, a 500 g sample was taken, and a quantity of the seed bran recovered for each step was measured. During seed processing, the speed of rotation and friction (seed-seed, seed-drum, and seed-baffles) gradually increased the temperature of the seeds to 40 ◦C and decreased the moisture from 13% to 10%. After carrying out the pearling process, we collected 99 samples, including 45 processed quinoa samples, 45 seed bran samples, and 9 samples of the untreated raw seeds of the three tested varieties.

#### *4.4. Physical Analysis*

A precision electronic balance (reading to 0.001 g) was used to determine the 1000 SW by weighing 100 seeds in triplicate and then extrapolating this weight to get 1000 SW [52]. Pictures of seeds were taken using optical microscopy Nikon Eclipse Lv100nd.

#### *4.5. Chemical Analysis*

After processing, quinoa samples were ground to a fine powder using FOSS CT 293 Cyclotec grinder with a range of screens of 0.3 mm.

The moisture content was measured by drying 100 g of the sample at 105 ◦C for 48 h according to ICC Standard 110/1. The protein, fat, and ash contents were determined according to AOAC procedures [53].

For the evaluation of protein content, a macro-Kjeldahl method (N × 6.25) was used. The fat content was determined by Soxhlet method, using hexane solvent for the extraction for 4 h [54].

The ash content was determined according to the AOAC 923.03 standard. Micronutrients were determined after sample mineralization. Representative samples (0.25 g) were digested with 7.5 mL of nitric acid (HNO3) in a DigiPrep system for two hours at 100 ◦C. After digestion, the solutions were filtered through 45 µm filters, and the filtrates were diluted to 50 mL with deionized water and acidified (2% HNO3) in order to undergo the analysis by ICP-OES using Agilent technologies 5110 ICP-OES [10].

All measurements were performed in triplicate.

#### *4.6. Saponin Determination by Liebermann–Buchard Colorimetric Assay*

The protocol allows the determination of triterpene saponins from quinoa seeds. The principle of the assay is based on the reaction between the Liebermann–Burchard (LB) reagent and the triterpene molecules by developing a color [55].

#### 4.6.1. Determining the Wavelength for the Dosage

A series of absorbance readings were taken in different wavelengths to get the wavelength corresponding to the maximum absorbance. We found that the maximal absorbance corresponds to 528 nm.

#### 4.6.2. Dilutions for Range Preparation

According to Table 3, a series of standard solutions (0.2, 0.3, 0.5, and 0.8 mL) were taken, and the mixed solutions were prepared by the method described above for measuring absorbance values.

**Table 3.** Dilution series for the standard curve determination.

#### 4.6.3. Saponin Extraction

Five grams of the defatted quinoa powder (Soxhlet extraction by using hexane solvent for 4 h and repeated three times) was taken in 50 mL of ethanol. The mixture was then shaken vigorously for 30 min to extract total saponins. Then, the mixture was filtered using a filter paper (*Fironi standard—Plisse*). The filtrate was collected and topped up to 50 mL with ethanol.

#### 4.6.4. Development of the color

The saponin content determination was carried out according to the method applied by Irigoyen et al. [55], by adding 1 mL of the diluted solution to 3.5 mL of the Lieberman-Buchard reagent (16.7% acetic anhydride in concentrated sulfuric acid). The solution was vortexed and stored in the dark for 30 min at room temperature. The absorbance of the solution was measured at 528 nm in a spectrophotometer (Agilent 8453 Spectrophotometer, Los Angeles, CA, USA). Figure 11 shows the development of the color for each dilution.

**Figure 11.** Different concentrations of the colored solutions.

4.6.5. Calibration Curve Preparation

Regarding the preparation of the stock solution of the standard, oleanolic acid (97%) (Sigma-Aldrich, St. Louis, MO, USA) was used as the pure standard for triterpene saponin; 10 mg of OA was dissolved in 10 mL of pure ethanol to obtain a stock solution concentration of 1 mg/mL [56].

The range of absorbance obtained after dilutions was between 0.2 and 1 mg/mL which corresponds to an absorption of 0.246 to 1.055 (Figure 12). Beer–Lambert's law is applicable in the straight part of the trace, and the range of linearity is between 0.2 and 0.8 absorbance units.

**Figure 12.** Calibration curve of oleanolic acid at 528 nm.

From the calibration curve, X is calculated as the following:

$$
\chi \left( \frac{mg}{ml} \right) = \frac{y - 0.0789}{1.0627} \tag{1}
$$

The concentration of saponins is calculated according to the following equation:

$$\text{Concentration} \left(\frac{\text{g}}{100\text{g}} DM\right) = \frac{\text{x} \left(\frac{\text{mg}}{\text{ml}}\right) \times V\_{\text{D}} \times 10^{-3} \times 100 \times 100}{(1 + \text{x}(\text{Fat})) \times \text{Sample weight } (\text{g}) \times (100 - \text{Moisture})} \tag{2}$$

#### *4.7. Statistical Analysis*

Statistical analysis was carried out using R version 3.6.2 software. A two-way analysis of variance (ANOVA) was used to assess the effects of the pearling process on each variety (physical and chemical parameters). The level of significance was set to *p* < 0.05.

Pearson's correlation coefficient was evaluated according to the linear correlations between any two parameters. If the obtained coefficient was −1 or +1, there was a perfect negative or positive linear relationship, respectively. The value of 0 denotes no correlation between the two variables. Generally, a correlation could be statistically significant at a 95% confidence interval (*p* < 0.05).

Principal component analysis (PCA) was used to investigate the potential correlation matrix between all the measured parameters and the pearling time, as well as the data groupings after the pearling treatment.

#### **5. Conclusions and Recommendations**

One of the limiting factors facing quinoa valorization and consumption in Morocco and other countries is the seed content of saponin. Furthermore, traditional ways of removing saponins using manual abrasion and washing require intensive labor and thus increase production costs. Therefore, developing mechanical dry pearling tools for saponin removal will solve two interlinked challenges—reducing the processing costs and saving water, thereby increasing farmer's income.

In our study, we tested three quinoa varieties grown in Morocco and polished with a locally manufactured machine. As most cleaning and saponin removal is done under wet or humid methods, our method by dry abrasion had advantageous results, which can be beneficial for farmers and processors.

The findings of this study clearly indicate that two-minute pearling was enough to remove saponins and keep their concentration below the consumption threshold, while at the same time preserving both physical and nutritional quality of the seeds. This study highlights the potentialities of the locally manufactured machine to be used in other regions in Morocco or in other countries, especially as it is a low-cost tool and could be easily operated by smallholder farmers.

**Author Contributions:** S.R. investigated physical parameters, conducted chemical analysis, analyzed the data, and wrote the manuscript; M.R. supervised chemical analysis and saponin analysis assays; J.P.R. contributed to manuscript writing-review and editing; S.A. carried out macronutrient analysis; A.E. conducted data statistical analysis; M.E.G. supervised micronutrient analysis at AITTC laboratory; S.K. conducted saponin analysis and interpretation; R.C.-A. supervised morphological aspects' results and wrote the manuscript; A.H. supervised the research activities, analyzed the data, and wrote the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This study was carried out within an IDRC (International Development Research Center-Canada) funded project entitled: Scaling up quinoa value chain to improve food and nutritional security in poor rural communities of Morocco under the agreement number 108637-001.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data generated within this work are open access and available to be shared with interested persons according to signed agreement with IDRC.

**Acknowledgments:** First, we thank our sponsor, the International Development Research Center (IDRC)—Canada, for providing fund to conduct the project activities. We also appreciate the efforts deployed by the technical team of UM6P—Agricultural Innovation and Technology Transfer Center (AITTC), and the technical support of the International Center of Biosaline Agriculture (ICBA). Special thanks to Benrim Farm (Haj Mohamed Ibrahimi) for his cooperation. Our gratitude goes to our local partners and facilitators (farmers, female cooperatives, startups, OCP Group, DRA Marrakech-Safi, DPA Rehamna, ONCA Rehamna, ONSSA Rehamna. etc.).

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

#### **References**

## *Article* **Free and Conjugated Phenolic Profiles and Antioxidant Activity in Quinoa Seeds and Their Relationship with Genotype and Environment**

**Fabiana Antognoni 1,\*, Giulia Potente <sup>1</sup> , Stefania Biondi <sup>2</sup> , Roberto Mandrioli <sup>1</sup> , Lorenzo Marincich <sup>1</sup> and Karina B. Ruiz <sup>3</sup>**

**Abstract:** The nutraceutical interest in quinoa (*Chenopodium quinoa* Willd.) seeds is associated with the presence of macronutrients, micronutrients, minerals, vitamins, and polyphenols. In particular, polyphenols contribute to the health-promoting effects of this food crop, and their levels are influenced by environmental conditions. Production of quinoa is recently being explored in temperate climate areas, including Italy. The aim of this research was to assess the profile of bioactive compounds in seeds of two quinoa varieties, Regalona-Baer and Titicaca, grown in northern Italy, compared to that of seeds of those varieties grown in Chile and Denmark, respectively. Highperformance liquid chromatography-diode array detector (HPLC-DAD) analysis of phenolic acid and flavonoid profiles, both in their free and soluble conjugated forms, showed that the main differences between Regalona grown in Chile and Italy were for the free vanillic acid and daidzein contents, while the two Titicaca samples mainly differed in quercetin derivative levels. The total phenolic index was comparable in Titicaca and Regalona, and only a slight decrease in this parameter was found in seeds of the two varieties grown in Italy. The in vitro antioxidant activity of seed extracts, evaluated by means of three different assays, indicated that it correlated with flavonol (quercetin derivative) levels. In conclusion, the results indicate that, although environmental conditions alter the polyphenolic profile and biological activities, it is possible to grow good-quality quinoa in northern Italy.

**Keywords:** antioxidant activity; *Chenopodium quinoa*; conjugated phenolics; flavonoids; phenolic acids; nutraceutical properties

#### **1. Introduction**

Quinoa (*Chenopodium quinoa* Willd.) seeds have an excellent nutritional profile [1,2], due to their remarkably high protein content, well-balanced content of essential amino acids comparable, in terms of quality, to that of casein [3], and the presence of dietary fiber, vitamins, and unsaturated fatty acids, e.g., linoleic and α-linolenic acids [2–6]. Quinoa seeds are considered an excellent example of a "functional food", able to exert healthpromoting effects also due to the presence of secondary metabolites [6]. These include phenolic acids and flavonoids [7], as well as terpenoids, steroids, and large amounts of α- and γ-tocopherol, which are known antioxidants [8]. For some of these specialized metabolites, a clear role in preventing various human diseases, such as neurological and other chronic disorders, has been recognized, as a result of their effects on cell signalling and metabolism [6,9,10].

**Citation:** Antognoni, F.; Potente, G.; Biondi, S.; Mandrioli, R.; Marincich, L.; Ruiz, K.B. Free and Conjugated Phenolic Profiles and Antioxidant Activity in Quinoa Seeds and Their Relationship with Genotype and Environment. *Plants* **2021**, *10*, 1046. https://doi.org/10.3390/ plants10061046

Academic Editors: Cataldo Pulvento and Didier Bazile

Received: 23 April 2021 Accepted: 19 May 2021 Published: 21 May 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

Quinoa cultivation remained, for a long time, exclusive to Andean populations, especially from Bolivia and Peru [11], and, to a lesser extent, from Chile and Ecuador. The wide genetic diversity of this species, resulting from its fragmented and localized cultivation over the centuries in the Andean region, together with its high tolerance to extreme environmental conditions and abiotic stresses [12,13], has led to the differentiation of five major ecotypes, based on their ability to adapt to specific agroecological conditions [11]. Nevertheless, in recent decades, with the boom in demand for "superfoods", the interest in this ancient Andean halophytic seed-producing crop has become global, and its cultivation has been growing accordingly. In less than twenty years, the global demand for quinoa grew to such an extent that it led to the triplication of the Andean areas dedicated to its cultivation [14]. Moreover, the adaptability and stress tolerance of this plant is being exploited to establish cultivations outside the Andean territories [11,15]. As a consequence, the number of countries growing quinoa has rapidly risen starting from the 1980s, and, nowadays, more than 95 countries are cultivating or testing quinoa as a crop [16]. In the early 1990s, quinoa began to be cultivated in Europe, and the crop has been successfully tested in several Mediterranean countries, such as Greece, Morocco, Spain, and Italy [17–20].

The main concern about the introduction of this crop was related to its high sensitivity to photoperiods during the reproductive phase [21]. The best results were achieved with the sea level/coastal ecotype from central and southern Chile, since it is the best adapted to temperate environments. Using these Chilean quinoa lines, a new variety, called Titicaca, was bred in Denmark [22]. Titicaca is one of four European quinoa cultivars (with Puno, Jessie, and Zeno) originating from different gene pools and already tested in various countries, including Germany [23] and Italy, where several field trials have been carried out at different latitudes to test the adaptability of the crop to varying environments [21,23,24]. Pulvento et al. [25] reported good plant performance and tolerance to high temperatures and water deficit, typical conditions of southern Italy, for cultivars Regalona-Baer and Titicaca.

Several studies point to the importance of environmental and/or agronomical factors in affecting the nutritional properties of quinoa. For example, Reguera et al. [26] showed that amino acid profiles, total protein content, mineral composition, and phytate content varied in seeds of three cultivars (Salcedo-INIA, Titicaca, Regalona) grown in three different countries (Spain, Peru, and Chile). However, the impact of agroecological conditions and crop management techniques on quinoa seed quality in terms of functional bioactive compounds (e.g., phenolics) has been little explored. Miranda et al. [27] reported that seeds of several quinoa genotypes grown in different geographical locations in Chile contained variable amounts of phenolics and flavonoids. Considering that these secondary metabolites are plastically produced in plant tissues as an acclimation response to environmental cues, their concentrations in plant tissues are the combined result of several factors, including genetic components, environmental conditions, and the complex interplay between the two [28–30]. In the present work, we checked this hypothesis by analysing the phenolic profiles of quinoa seeds of the same variety from different agroecological environments and verified if the antioxidant activity (AA) of seed extracts likewise changed. Such analyses are useful to identify the environmental conditions that modulate the health-promoting characteristics of quinoa seeds. To this purpose, we evaluated the composition in bioactive free and soluble-conjugated phenolic acids and flavonoids and the AA of seeds of two quinoa varieties, the Chilean Regalona-Baer and the Danish-bred Titicaca, grown in northern Italy and compared them with those of seeds of the same varieties grown in Chile and Denmark. The present results could help to establish the best conditions (genotype/geographical cultivation zone) leading to seed enrichment in functional compounds and to reinforce the notion that good-quality quinoa can be grown in Italy.

**2. Results and Discussion**

*(TFC) in Quinoa Seed Extracts*

#### **2. Results and Discussion** Phenolic acids and flavonoids in seed extracts were quantified by analysing the free

*Plants* **2021**, *10*, 1046 3 of 15

#### *2.1. Free and Soluble-Conjugated Total Phenolic Content (TPC) and Total Flavonoid Content (TFC) in Quinoa Seed Extracts* and soluble‐conjugated forms (and not the insoluble ones) because they represent the frac‐

Phenolic acids and flavonoids in seed extracts were quantified by analysing the free and soluble-conjugated forms (and not the insoluble ones) because they represent the fractions of dietary phenolic compounds that are more rapidly absorbed by the stomach and small intestine and for which important health benefits have been reported [31]. tions of dietary phenolic compounds that are more rapidly absorbed by the stomach and small intestine and for which important health benefits have been reported [31]. Total phenolic content (TPC) and total flavonoid content (TFC) were evaluated in extracts of the four seed samples enriched in the solvent‐extractable (free; F) and in both

Total phenolic content (TPC) and total flavonoid content (TFC) were evaluated in extracts of the four seed samples enriched in the solvent-extractable (free; F) and in both the acid-hydrolysed (AH) and base-hydrolysed (BH) soluble-conjugated forms (Figure 1). the acid‐hydrolysed (AH) and base‐hydrolysed (BH) soluble‐conjugated forms (Figure 1).

**Figure 1.** Total phenolic content (**A**) and total flavonoid content (**B**) in free (F), acid‐hydrolysed (AH), and base‐hydrolysed (BH) fractions of quinoa seed extracts. (**A**) TPC is expressed as mg gallic acid equivalents (GAE)/g DW seed powder. (**B**) TFC is expressed as catechin equivalents (CE)/g DW seed powder. RI: Regalona grown in Italy; RC: Regalona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in Denmark. Data are the means ± standard error of two inde‐ **Figure 1.** Total phenolic content (**A**) and total flavonoid content (**B**) in free (F), acid-hydrolysed (AH), and base-hydrolysed (BH) fractions of quinoa seed extracts. (**A**) TPC is expressed as mg gallic acid equivalents (GAE)/g DW seed powder. (**B**) TFC is expressed as catechin equivalents (CE)/g DW seed powder. RI: Regalona grown in Italy; RC: Regalona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in Denmark. Data are the means ± standard error of two independent determinations with three biological replicates. Different letters within the same parameter indicate statistically significant differences at *p* ≤ 0.05.

pendent determinations with three biological replicates. Different letters within the same parame‐ ter indicate statistically significant differences at *p ≤* 0.05. As concerns TPC (Figure 1A), AH was the richest fraction, with levels reaching 1.7– 1.8 mg GAE/g DW of seed powder, followed by the BH fraction (0.6–0.8 mg GAE/g DW), while much lower levels were observed in the free form (0.2–0.3 mg/g DW). In both the F and BH fractions, no significant differences in TPC were observed between RI, RC, TI, and TD. By contrast, the Danish‐grown Titicaca seeds (TD) had about twice the concentration of AH soluble‐conjugated phenolics as compared with the same cultivar grown in Italy. As regards Regalona, the TPC in this fraction (AH) was very similar in both Chilean‐ and Italian‐grown seeds, and not significantly different from that of TD (Figure 1A). This sug‐ gests that the conjugated form of soluble phenolics is mainly linked through ether bonds, such as the glycosidic one, which is acid hydrolysable. The distribution of TPC into the three fractions in the quinoa seeds analysed here is in line with that reported by Tang et al. [32] in a white seed quinoa, but not with that by Gómez‐Caravaca et al. [33] for geno‐ types Kancholla and Witulla. These authors reported that most phenolic compounds pre‐ As concerns TPC (Figure 1A), AH was the richest fraction, with levels reaching 1.7–1.8 mg GAE/g DW of seed powder, followed by the BH fraction (0.6–0.8 mg GAE/g DW), while much lower levels were observed in the free form (0.2–0.3 mg/g DW). In both the F and BH fractions, no significant differences in TPC were observed between RI, RC, TI, and TD. By contrast, the Danish-grown Titicaca seeds (TD) had about twice the concentration of AH soluble-conjugated phenolics as compared with the same cultivar grown in Italy. As regards Regalona, the TPC in this fraction (AH) was very similar in both Chilean- and Italian-grown seeds, and not significantly different from that of TD (Figure 1A). This suggests that the conjugated form of soluble phenolics is mainly linked through ether bonds, such as the glycosidic one, which is acid hydrolysable. The distribution of TPC into the three fractions in the quinoa seeds analysed here is in line with that reported by Tang et al. [32] in a white seed quinoa, but not with that by Gómez-Caravaca et al. [33] for genotypes Kancholla and Witulla. These authors reported that most phenolic compounds present in their seed extracts were in the free form, but it should be underlined that these authors used a different analytical method, i.e., HPLC-DAD-ESI-TOF-MS, which could explain the discrepancies. Although Repo-Carrasco-Valencia et al. [7] observed a huge variation in the proportion of soluble (free and conjugated) phenolics (ranging from 7 to 61%), among the ten genotypes taken into consideration, we did not observe large differences in the total amounts of soluble phenolics between the two T and R varieties. It should be pointed out that the TPC in the AH fraction might be overestimated, since several authors [32,34] indicated that, under acid hydrolysis, the degradation of free

variation in the proportion of soluble (free and conjugated) phenolics (ranging from 7 to 61%), among the ten genotypes taken into consideration, we did not observe large differ‐ ences in the total amounts of soluble phenolics between the two T and R varieties. It should be pointed out that the TPC in the AH fraction might be overestimated, since sev‐ eral authors [32,34] indicated that, under acid hydrolysis, the degradation of free sugars can give rise to furan derivatives, which react with the Folin–Ciocalteu's reagent. Thus,

sent in their seed extracts were in the free form, but it should be underlined that these

sugars can give rise to furan derivatives, which react with the Folin–Ciocalteu's reagent. Thus, the distribution of phenolic compounds between the free and AH conjugated forms, resulting from the TPC determination, only provides a rough estimate, which must be confirmed through more specific analytical techniques. the distribution of phenolic compounds between the free and AH conjugated forms, re‐ sulting from the TPC determination, only provides a rough estimate, which must be con‐ firmed through more specific analytical techniques. As concerns TFC, both the relative distribution in the three forms and the pattern

As concerns TFC, both the relative distribution in the three forms and the pattern within each fraction were similar to those of TPC; the conjugated forms were more abundant than the free ones, even though, in this case, the levels of AH and BH soluble flavonoids were comparable (Figure 1B). In plants, it is known that flavonoids are mostly conjugated to various types of molecules. The abundance of flavonoids in both the AH and BH fractions indicates that these were linked via both ether and ester bonds; in the former category, glycosides represent the most widespread form, while the latter suggest their linkage to organic acids and proteins, as demonstrated by Koistinen et al. [35]. As concerns the comparison among samples, the pattern of TFC in the free and AH fractions was very similar to that of TPC, with AH forms again higher in TD than in TI. Danish-grown Titicaca showed the highest TFC in the BH fraction, while no differences were found between the other three samples (Figure 1B). Based on these results, both genotype and environmental conditions seem to exert their effect mainly on the conjugated form of phenolics, leaving the free form almost unchanged. Moreover, they indicate that TI had the lowest level of phenolics conjugated through ether linkages. within each fraction were similar to those of TPC; the conjugated forms were more abun‐ dant than the free ones, even though, in this case, the levels of AH and BH soluble flavo‐ noids were comparable (Figure 1B). In plants, it is known that flavonoids are mostly con‐ jugated to various types of molecules. The abundance of flavonoids in both the AH and BH fractions indicates that these were linked via both ether and ester bonds; in the former category, glycosides represent the most widespread form, while the latter suggest their linkage to organic acids and proteins, as demonstrated by Koistinen et al. [35]. As concerns the comparison among samples, the pattern of TFC in the free and AH fractions was very similar to that of TPC, with AH forms again higher in TD than in TI. Danish‐grown Titi‐ caca showed the highest TFC in the BH fraction, while no differences were found between the other three samples (Figure 1B). Based on these results, both genotype and environ‐ mental conditions seem to exert their effect mainly on the conjugated form of phenolics, leaving the free form almost unchanged. Moreover, they indicate that TI had the lowest level of phenolics conjugated through ether linkages.

#### *2.2. Chromatographic Analysis of Free and Soluble-Conjugated Phenolic Compounds in Quinoa Seed Extracts 2.2. Chromatographic Analysis of Free and Soluble‐Conjugated Phenolic Compounds in Quinoa Seed Extracts*

The three fractions were subjected to high-performance liquid chromatographydiode array detector (HPLC-DAD) analysis to identify and quantify single phenolic compounds. These include phenolic acids (both hydroxycinnamic and dihydroxybenzoic acids), flavonols, flavan-3-ols, and isoflavones (Figure 2). The three fractions were subjected to high‐performance liquid chromatography‐di‐ ode array detector (HPLC‐DAD) analysis to identify and quantify single phenolic com‐ pounds. These include phenolic acids (both hydroxycinnamic and dihydroxybenzoic ac‐ ids), flavonols, flavan‐3‐ols, and isoflavones (Figure 2).

**Figure 2.** Chromatogram of HPLC‐DAD separation of (**A**) standard phenolic acids and flavonoids, and (**B**) representative chromatographic profiles of free (upper line), base‐hydrolysed (middle line), and acid‐hydrolysed (bottom line) fractions of one seed extract (TI). **Figure 2.** Chromatogram of HPLC-DAD separation of (**A**) standard phenolic acids and flavonoids, and (**B**) representative chromatographic profiles of free (upper line), base-hydrolysed (middle line), and acid-hydrolysed (bottom line) fractions of one seed extract (TI).

In the free fraction, vanillic and ferulic acids were the major hydroxycinnamic acids detected in all samples, the former being about 10‐fold more concentrated than the latter (Figure 3). Seed extracts of Chilean Regalona had a vanillic acid content about twice that of the same genotype grown in Italy, and of both TD and TI, whose concentrations were very similar. As for ferulic acid levels, no differences were observed between the two cul‐ tivars, no matter where they were grown (Figure 3). In the free fraction, vanillic and ferulic acids were the major hydroxycinnamic acids detected in all samples, the former being about 10-fold more concentrated than the latter (Figure 3). Seed extracts of Chilean Regalona had a vanillic acid content about twice that of the same genotype grown in Italy, and of both TD and TI, whose concentrations were very similar. As for ferulic acid levels, no differences were observed between the two cultivars, no matter where they were grown (Figure 3).

**Figure 3.** Free phenolic acid and flavonoid concentrations in quinoa seed extracts. RI: Regalona grown in Italy; RC: Rega‐ lona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in Denmark. Data are the means ± standard error of two independent determinations with three biological replicates. Different letters within the same parameter indicate sta‐ tistically significant differences at *p ≤* 0.05*.* **Figure 3.** Free phenolic acid and flavonoid concentrations in quinoa seed extracts. RI: Regalona grown in Italy; RC: Regalona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in Denmark. Data are the means ± standard error of two independent determinations with three biological replicates. Different letters within the same parameter indicate statistically significant differences at *p* ≤ 0.05.

The prevalence of vanillic acid as the main representative of the phenolic acid class in the free form is in line with the results of Tang et al. [32] in a white seed quinoa, even though significant variations in the relative amounts of single phenolic acids in different quinoa genotypes and varieties were also reported by Repo‐Carrasco‐Valencia et al. [7]. Nevertheless, a comparison with the results from these authors is not easy, since they re‐ The prevalence of vanillic acid as the main representative of the phenolic acid class in the free form is in line with the results of Tang et al. [32] in a white seed quinoa, even though significant variations in the relative amounts of single phenolic acids in different quinoa genotypes and varieties were also reported by Repo-Carrasco-Valencia et al. [7]. Nevertheless, a comparison with the results from these authors is not easy, since they reported the soluble phenolic acid content without distinguishing between the free and the soluble-conjugated fraction.

ported the soluble phenolic acid content without distinguishing between the free and the soluble‐conjugated fraction. As concerns flavonoids, epicatechin, quercetin, and kaempferol derivatives (querce‐ tin‐3‐*O*‐rutinoside, quercetin‐3‐*O*‐glucoside, kaempferol‐3‐*O*‐glucoside, and kaempferol‐ 3‐*O*‐rutinoside) and the isoflavones daidzein and genistein were detected in the free frac‐ tion (Figure 3). Significant differences among samples were observed only in some cases. In particular, TD seeds showed higher levels of quercetin‐3‐*O*‐glucoside and quercetin‐3‐ *O*‐rutinoside compared to seeds of the same cultivar grown in Italy (two‐ and four‐fold, respectively), and compared to both Regalona extracts. In Regalona seeds, a higher level of these two quercetin derivatives was found in Italian samples as compared to Chilean ones. Conversely, the latter showed the highest daidzein levels among all samples, and no differences in this isoflavone were found among other seeds. All seed extracts con‐ tained very low amounts of kaempferol derivatives, whose differences were of no biolog‐ ical relevance, nor were any differences observed as concerns genistein and epicatechin As concerns flavonoids, epicatechin, quercetin, and kaempferol derivatives (quercetin-3-*O*-rutinoside, quercetin-3-*O*-glucoside, kaempferol-3-*O*-glucoside, and kaempferol-3-*O*rutinoside) and the isoflavones daidzein and genistein were detected in the free fraction (Figure 3). Significant differences among samples were observed only in some cases. In particular, TD seeds showed higher levels of quercetin-3-*O*-glucoside and quercetin-3- *O*-rutinoside compared to seeds of the same cultivar grown in Italy (two- and four-fold, respectively), and compared to both Regalona extracts. In Regalona seeds, a higher level of these two quercetin derivatives was found in Italian samples as compared to Chilean ones. Conversely, the latter showed the highest daidzein levels among all samples, and no differences in this isoflavone were found among other seeds. All seed extracts contained very low amounts of kaempferol derivatives, whose differences were of no biological relevance, nor were any differences observed as concerns genistein and epicatechin (Figure 3). Thus, the agroclimatic conditions in which the Chilean Regalona seeds were produced seem to be particularly favourable for the accumulation of daidzein since, in this sample, the highest daidzein/genistein ratio (=6) was found. This value is very close to that observed by Lutz et al. [36] in some commercial quinoa seeds and suggests that, under certain environmental conditions, a seed with a good phytoestrogen-like activity can be obtained.

(Figure 3). Thus, the agroclimatic conditions in which the Chilean Regalona seeds were produced seem to be particularly favourable for the accumulation of daidzein since, in In the AH conjugated fraction, *p*-coumaric and ferulic acids did not vary considerably among samples (Figure 4). The ferulic acid concentration was very similar to that found

this sample, the highest daidzein/genistein ratio (=6) was found. This value is very close

under certain environmental conditions, a seed with a good phytoestrogen‐like activity

In the AH conjugated fraction, *p*‐coumaric and ferulic acids did not vary considerably among samples (Figure 4). The ferulic acid concentration was very similar to that found in the free fraction, while *p*‐coumaric acid was not detected in the free form. As concerns

can be obtained.

0.0

2.0

4.0

6.0

g/g DW

a

a

a

flavonoids, the aglycones quercetin and kaempferol were found as the main representa‐

*Plants* **2021**, *10*, 1046 6 of 15

Phenolic acids and flavonoids were also detected in the BH soluble conjugated frac‐ tion. Both *p*‐coumaric and ferulic acids were present, and the latter was detected at higher Phenolic acids and flavonoids were also detected in the BH soluble conjugated fraction. Both *p*-coumaric and ferulic acids were present, and the latter was detected at higher levels than in the AH fraction (Figure 5).

levels than in the AH fraction (Figure 5). RI b c b a b RC TI a a b b TD a a a a Vanillic acid was present at similar levels to those of the free fraction, while conjugated ferulic acid was over ten-fold more concentrated than in the free form. Thus, the results indicate that a relevant portion of these polyphenols is conjugated through ester bonds with soluble cellular components, such as short peptides and/or low-molecular weight oligosaccharides. It is known that ferulic and *p*-coumaric acids are covalently bound to mixed-linkage (1→3, 1→4)-β-D-glucans and to hemicelluloses, forming the soluble dietary fiber [37]. Being a dicot species, quinoa cell walls contain mainly xyloglucans in their matrix. The extractability of hemicelluloses during the preparation of crude seed extracts using sonication has been demonstrated [38] as well as the binding of ferulic acid to shortchain hemicelluloses [39]. Thus, the crude extracts used in this study probably contain soluble feruloylated and/or coumaroylated xylo-oligosaccharides, which could explain the presence of high levels of aglycone components in our BH fraction. Several healthpromoting activities have been reported for these short-chain feruloylated oligosaccharides, including an immunomodulatory effect [40]. As concerns the comparison among samples, the results indicate that concentrations of vanillic and *p*-coumaric acids did not differ significantly between Regalona and Titicaca, no matter where they were grown, while for ferulic acid, a 55% higher concentration was found in RC compared to seeds of the same genotype grown in Italy; no differences were found between it and TD or TI seeds.

a

a

a a

Vanillic acid Ferulic acid p-Coumaric acid Que-3-O-rut Kae-3-O-glu

**Figure 5.** Phenolic acid and flavonoid concentrations in the base‐hydrolysed solubleconjugated fraction of quinoa seed extracts. RI: Regalona grown in Italy; RC: Regalona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in

c

a

g/g DW

4).

a

a

a

c

**Figure 4.** Phenolic acid and flavonoid concentrations in the acid‐hydrolysable soluble‐conjugated fraction of quinoa seed extracts. RI: Regalona grown in Italy; RC: Regalona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in Denmark. Data are the means ± standard error of two independent determinations with three biological replicates. Differ‐

<sup>a</sup> <sup>a</sup>

p-Coumaric acid Ferulic acid Quercetin Kaempferol

ent letters within the same parameter indicate statistically significant differences at *p ≤* 0.05*.*

a

a

levels than in the AH fraction (Figure 5).

**Figure 5.** Phenolic acid and flavonoid concentrations in the base‐hydrolysed solubleconjugated fraction of quinoa seed extracts. RI: Regalona grown in Italy; RC: Regalona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in **Figure 5.** Phenolic acid and flavonoid concentrations in the base-hydrolysed solubleconjugated fraction of quinoa seed extracts. RI: Regalona grown in Italy; RC: Regalona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in Denmark. Data are the means ± standard error of two independent determinations with three biological replicates. Different letters within the same parameter indicate statistically significant differences at *p* ≤ 0.05.

Quercetin-3-*O*-rutinoside (or rutin) and kaempferol-3-*O*-glucoside were identified as the main flavonols in the BH fraction. As regards the former, the pattern was similar to those in the free and AH conjugated fractions, with DT being richer than the Italiangrown counterpart, while RI had a slightly, but significantly, higher rutin content than RC; no differences were observed for kaempferol-3-*O*-glucoside among the four samples (Figure 4).

flavonoids, the aglycones quercetin and kaempferol were found as the main representa‐ tives, and TD was confirmed to be the richest in these flavonols (about three‐fold higher than in TI), while no differences were detected between the two Regalona samples (Figure

b

c

c

a

RI

RC

TI

TD

a

Phenolic acids and flavonoids were also detected in the BH soluble conjugated frac‐ tion. Both *p*‐coumaric and ferulic acids were present, and the latter was detected at higher

b

b

c

Some authors have investigated the phenolic composition of different quinoa varieties, and phenolic acids and flavonoids were reported to exist both in their aglycone and glycosidic forms [32,41,42]. On average, the concentrations of these compounds found in our work were similar to those found by other authors [41,43], but lower compared to other reports [33]. These discrepancies are not surprising, since several parameters were demonstrated to have a great impact on the extraction efficiency of these metabolites from plant material, such as sampling, solvent-to-solid ratio used for extraction, duration of extraction, and sonication treatment applied, among others [44]. The use of more advanced separative techniques, such as the HPLC-DAD-ESI-TOF-MS utilized by Gómez-Caravaca et al. [33], allowed these authors to identify more compounds, which are not detectable with other techniques, and this can also explain the differences.

Hirose et al. [45] reported that quinoa and buckwheat contain several quercetin and kaempferol derivatives, which are not present in any of the widely consumed cereals. They also compared the flavonoid pattern and in vitro antioxidant capacity of seeds cultivated in Japan with those of seeds from Bolivia and Peru. Results indicated that the major differences between the two groups of seeds were found for quercetin and kaempferol derivatives, which were significantly higher in Japanese-grown seeds compared to South American ones, and this was accompanied by a significantly higher antioxidant capacity. Seasonal variations in the pattern of flavonol glycosides observed by these authors allowed them to hypothesize that sunlight was the factor that mostly influenced the accumulation of quercetin glycosides [45]. Thus, it is possible that variations in quercetin and its derivatives between Titicaca and Regalona grown at different latitudes may be due to differences in

day length and light quality/intensity. Even though the effect of latitude on flavonoid accumulation has yet to be fully clarified, due to the complexity of latitude itself as a parameter to be investigated [29], many studies have concluded that northern climates may have a positive impact on the biosynthesis of flavonoids in plants, although there are variations in the response among species and within individual flavonoid groups. Flavonols, especially moieties of quercetin or kaempferol, have been reported to accumulate in response to increased UV-B radiation [46], which typically characterizes the higher latitudes in the Northern Hemisphere. The prominent role for this subclass of flavonoids in regulating plant–environment interactions has been demonstrated, particularly as concerns the acclimation of plants to different light exposures [47]. In general, flavonoids may play prominent roles as scavengers of reactive oxygen species (ROS) generated by adverse conditions [48,49], and quercetin derivatives, due to their chemical characteristics, are particularly efficient in buffering alterations in ROS homeostasis [28]. Moreover, several reports indicate that they strongly affect phytohormone (auxin and ABA) signalling, due to their ability to affect the activity of a wide range of proteins [50,51]. Thus, the higher accumulation of quercetin derivatives in Titicaca seeds grown in Denmark compared to seeds of the same cultivar grown in Italy and to both Regalona samples might be the result of an acclimation mechanism to long light exposures, typical of the higher latitudes. The bioavailability and therapeutic potentials of quercetin and its derivatives in plant-based foods have been extensively investigated [52], and several studies demonstrated that they play relevant roles both in the prevention and treatment of chronic diseases, including cardiovascular and neurodegenerative ones and some types of cancer [53–55]. Thus, the enrichment of quinoa seeds in flavonol derivatives could represent an added value able to reinforce the well-known nutraceutical properties of this plant. vonoids in regulating plant–environment interactions has been demonstrated, particu‐ larly as concerns the acclimation of plants to different light exposures [47]. In general, flavonoids may play prominent roles as scavengers of reactive oxygen species (ROS) gen‐ erated by adverse conditions [48,49], and quercetin derivatives, due to their chemical char‐ acteristics, are particularly efficient in buffering alterations in ROS homeostasis [28]. Moreover, several reports indicate that they strongly affect phytohormone (auxin and ABA) signalling, due to their ability to affect the activity of a wide range of proteins [50,51]. Thus, the higher accumulation of quercetin derivatives in Titicaca seeds grown in Denmark compared to seeds of the same cultivar grown in Italy and to both Regalona samples might be the result of an acclimation mechanism to long light exposures, typical of the higher latitudes. The bioavailability and therapeutic potentials of quercetin and its derivatives in plant‐based foods have been extensively investigated [52], and several stud‐ ies demonstrated that they play relevant roles both in the prevention and treatment of chronic diseases, including cardiovascular and neurodegenerative ones and some types of cancer [53–55]. Thus, the enrichment of quinoa seeds in flavonol derivatives could rep‐ resent an added value able to reinforce the well‐known nutraceutical properties of this *2.3. Antioxidant Activity of Quinoa Seed Extracts*

there are variations in the response among species and within individual flavonoid groups. Flavonols, especially moieties of quercetin or kaempferol, have been reported to accumulate in response to increased UV‐B radiation [46], which typically characterizes the higher latitudes in the Northern Hemisphere. The prominent role for this subclass of fla‐

#### *2.3. Antioxidant Activity of Quinoa Seed Extracts* The AA of the three fractions of Regalona and Titicaca seed extracts was evaluated

plant.

The AA of the three fractions of Regalona and Titicaca seed extracts was evaluated using three different in vitro assays, i.e., FRAP, TEAC, and ORAC (Figure 6). using three different in vitro assays, i.e., FRAP, TEAC, and ORAC (Figure 6).

*Plants* **2021**, *10*, 1046 8 of 15

**Figure 6.** Antioxidant activity, assayed by TEAC (**A**) and ORAC (**B**), in free (F), acid‐hydrolysed (AH), and base‐hydro‐ lysed (BH) fractions of seed extracts from different quinoa samples. Results are expressed as Trolox equivalents (TE)/g DW seed powder. RI: Regalona grown in Italy; RC: Regalona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in Denmark. Data are the means ± standard error of two independent determinations with three biological repli‐ **Figure 6.** Antioxidant activity, assayed by TEAC (**A**) and ORAC (**B**), in free (F), acid-hydrolysed (AH), and base-hydrolysed (BH) fractions of seed extracts from different quinoa samples. Results are expressed as Trolox equivalents (TE)/g DW seed powder. RI: Regalona grown in Italy; RC: Regalona grown in Chile; TI: Titicaca grown in Italy; TD: Titicaca grown in Denmark. Data are the means ± standard error of two independent determinations with three biological replicates. Different letters within the same parameter indicate statistically significant differences at *p* ≤ 0.05.

cates. Different letters within the same parameter indicate statistically significant differences at *p* ≤ 0.05. The results obtained by the TEAC and ORAC assays univocally show that the frac‐ tions enriched in soluble‐conjugated phenolic acids and flavonoids possess a higher AA compared to the free fraction (Figure 6A,B). FRAP provided the same results (data not shown). Although this may derive, at least in part, from the above‐cited overestimation The results obtained by the TEAC and ORAC assays univocally show that the fractions enriched in soluble-conjugated phenolic acids and flavonoids possess a higher AA compared to the free fraction (Figure 6A,B). FRAP provided the same results (data not shown). Although this may derive, at least in part, from the above-cited overestimation due to the formation of 5-hydroxymethyl-2-furfural, which displays AA [34], it has been widely demonstrated that soluble-conjugated polyphenols and/or feruloy-

due to the formation of 5‐hydroxymethyl‐2‐furfural, which displays AA [34], it has been

of grains [56,57]. As concerns the comparison among samples, the results obtained with the TEAC/FRAP and ORAC assays are slightly different. For all fractions (free, AH, BH), the former assay (Figure 6A; FRAP data not shown) revealed that TD had a slightly, but significantly, higher AA compared to TI and to both Regalona extracts, which, conversely, *Plants* **2021**, *10*, 1046 9 of 15

lated/coumaroylated short oligosaccharides are able to greatly contribute to the antioxidant capacity of grains [56,57]. As concerns the comparison among samples, the results obtained with the TEAC/FRAP and ORAC assays are slightly different. For all fractions (free, AH, BH), the former assay (Figure 6A; FRAP data not shown) revealed that TD had a slightly, but significantly, higher AA compared to TI and to both Regalona extracts, which, conversely, did not differ from each other. According to ORAC, no differences among samples were observed in the free fraction, while for the AH one, TD was confirmed to have the highest AA; no differences were found between the two Regalona samples grown in Chile and in Italy (Figure 6B). In the BH fraction, a statistically significant difference emerged between the Italian-grown Regalona and both RC and TD, even though the biological relevance of this difference is questionable. Thus, it can be concluded that the higher amount of flavonol derivatives in Danish-grown Titicaca might contribute to the higher antioxidant activity, given the strong redox capacity associated to this subgroup of flavonoids. AA; no differences were found between the two Regalona samples grown in Chile and in Italy (Figure 6B). In the BH fraction, a statistically significant difference emerged between the Italian‐grown Regalona and both RC and TD, even though the biological relevance of this difference is questionable. Thus, it can be concluded that the higher amount of flavo‐ nol derivatives in Danish‐grown Titicaca might contribute to the higher antioxidant activ‐ ity, given the strong redox capacity associated to this subgroup of flavonoids. For an overall comparison among samples, the total phenolic index (TPI) and the total antioxidant index (TAI) were calculated as the sum of the individual phenolic com‐ pounds and AA, respectively, detected in all fractions. As shown in Figure 7A, despite a different distribution between the free and soluble conjugated forms, the results clearly indicate that the Chilean Regalona and the Danish Ti‐

did not differ from each other. According to ORAC, no differences among samples were observed in the free fraction, while for the AH one, TD was confirmed to have the highest

For an overall comparison among samples, the total phenolic index (TPI) and the total antioxidant index (TAI) were calculated as the sum of the individual phenolic compounds and AA, respectively, detected in all fractions. ticaca show a very similar TPI, while seeds of these cultivars grown in Italy respond to the different climatic/environmental conditions with a 20% lower accumulation of the main

As shown in Figure 7A, despite a different distribution between the free and soluble conjugated forms, the results clearly indicate that the Chilean Regalona and the Danish Titicaca show a very similar TPI, while seeds of these cultivars grown in Italy respond to the different climatic/environmental conditions with a 20% lower accumulation of the main phenolic compounds compared to their counterparts. The results of the two-way ANOVA indicate that the area of cultivation significantly contributed to the variance of TPI (78%, *p* = 0.005), while neither the genotypes nor their interaction had a significant impact on the variation (*p* = 0.13 and 0.56, respectively). As for the TAI resulting from the ORAC assay (Figure 7B), it was only slightly lower (15%) in RI compared to RC, while a greater difference (40%) was observed between TD and TI; similar results were found for the TEAC assay (data not shown). These results confirm that the total antioxidant activity only weakly correlates with the TPI, as it is more tightly related to the specific composition in antioxidants; in fact, single compounds or classes of compounds can provide a greater contribution compared to others. The two-way ANOVA indicated that the total antioxidant index did not vary significantly depending on genotype (*p* = 0.61), while the cultivation area, as well as its interaction with genotype, provided a significant contribution (*p <* 0.0001 and *p* = 0.0031, respectively). phenolic compounds compared to their counterparts. The results of the two‐way ANOVA indicate that the area of cultivation significantly contributed to the variance of TPI (78%, *p* = 0.005), while neither the genotypes nor their interaction had a significant impact on the var‐ iation (*p* = 0.13 and 0.56,respectively). As forthe TAIresulting from the ORAC assay (Figure 7B), it was only slightly lower (15%) in RI compared to RC, while a greater difference (40%) was observed between TD and TI; similar results were found for the TEAC assay (data not shown). These results confirm that the total antioxidant activity only weakly correlates with the TPI, as it is more tightly related to the specific composition in antioxi‐ dants; in fact, single compounds or classes of compounds can provide a greater contribu‐ tion compared to others. The two‐way ANOVA indicated that the total antioxidant index did not vary significantly depending on genotype (*p* = 0.61), while the cultivation area, as well as its interaction with genotype, provided a significant contribution (*p <* 0.0001 and *p* = 0.0031, respectively*).*

**Figure 7.** Total phenolic index (TPI, **A**) and total antioxidant index (TAI, **B**) in the four seed ex‐ tracts of quinoa. TPI was calculated as the sum of individual phenolic compounds detected, and TAI as the sum of ORAC‐assayed AA in each fraction. **Figure 7.** Total phenolic index (TPI, **A**) and total antioxidant index (TAI, **B**) in the four seed extracts of quinoa. TPI was calculated as the sum of individual phenolic compounds detected, and TAI as the sum of ORAC-assayed AA in each fraction.

type, and were bred from gene pools originating from southern/central Chile. The results also indicate that agroecological conditions can, to some extent, alter these phytochemical profiles and the biological activities; in particular, light/UV‐B intensity may have contrib‐ uted to higher flavonol levels in seeds from Denmark [58,59]. Nonetheless, the changes are quite small and corroborate previous data [25], demonstrating that both Titicaca and

In conclusion, the present results suggest that, both in terms of phenolic profiles and

In conclusion, the present results suggest that, both in terms of phenolic profiles and AA, genotype-dependent differences are not very relevant as far as Titicaca and Regalona are concerned, both of which are adapted to temperate climates, belong to the coastal ecotype, and were bred from gene pools originating from southern/central Chile. The results also indicate that agroecological conditions can, to some extent, alter these phytochemical profiles and the biological activities; in particular, light/UV-B intensity may have contributed to higher flavonol levels in seeds from Denmark [58,59]. Nonetheless, the changes are quite small and corroborate previous data [25], demonstrating that both Titicaca and Regalona can be successfully cultivated in Italy, where they maintain good growth, productivity, and nutritional properties.

#### **3. Materials and Methods**

#### *3.1. Chemicals and Reagents*

The following chemicals and reagents were purchased from Sigma-Aldrich Italia (Milan, Italy): Folin–Ciocalteu's phenol reagent, 6-hydroxyl-2,5,7,8-tetramethyl-chroman-2 carboxylic acid (Trolox), 2,2-azinobis (3-ethylbenzothiazoline-6-sulfonic acid; ABTS), AlCl3, NaNO2, FeCl2, FeCl3, FerroZine®, fluorescein, 2,2-azobis(2-amidinopropane) dihydrochloride (AAPH), phosphoric acid (85–87%, *w*/*w*), hydrochloric acid (37%, *w*/*w*), monobasic sodium phosphate (>98%), sodium hydroxide beads (>98%), and HPLC-grade solvents. Pure standards of phenolic acids (4-hydroxybenzoic, gallic, caffeic, chlorogenic, ferulic, *p*-coumaric, sinapic, syringic, trans-cinnamic, and vanillic acids) and flavonoids (quercetin, quercetin-3-*O*-glucoside, quercetin-3-*O*-rutinoside, quercetin-3-*O*-galactoside, kaempferol, kaempferol-3-*O*-rutinoside, catechin, epicatechin, daidzein, and genistein) were purchased from Extrasynthese (Genay Cedex, France). The IUPAC names for these compounds are indicated in Table S1. All standards (>99.5% purity in powder form) were prepared as stock solutions at 1 mg/mL in methanol and stored in the dark at −18 ◦C for less than three months.

#### *3.2. Plant Material and Extraction Procedure*

Seeds of quinoa cultivars Regalona-Baer (opaque white) and Titicaca (pale yellow) harvested in 2017 were used. Seeds of cv. Regalona collected from plants grown in Italy (RI) were purchased from Dall'Ara & Lolli farm (Campiano, Ravenna, Italy; 44◦1801000 N, 12◦1200700 E; mean annual temperature 14.4 ◦C, max 19.0 ◦C, min 8.3 ◦C; total annual precipitation 646.0 mm), while seeds of the same cultivar grown in Chile (RC) were purchased from the seed company Semillas Baer (Temuco, Chile; 38◦4402300 S, 72◦3600000 W; mean annual temperature 10.9 ◦C, max 17.3 ◦C, min 5.9 ◦C; total annual precipitation 1355.2 mm). Seeds of quinoa cv. Titicaca grown in Italy (TI) were kindly supplied by D. Vanuzzi (Tuttoquinoa) and collected from plants grown in Sale (Alessandria, Italy; 44◦5805400 N, 8 ◦4803700 E; mean annual temperature 13.5 ◦C, max 18.6 ◦C, min 8.5 ◦C; total annual precipitation 501.5 mm), while Titicaca seeds from plants grown in Denmark (TD) were kindly supplied by S. Jacobsen (Copenhagen, Denmark; 55◦3804600 N, 12◦1705300 E; mean annual temperature 8.9 ◦C, max 19.2 ◦C, min −3.4 ◦C; total annual precipitation 848.8 mm).

The extractions were performed in triplicate, following the procedure described by Tang et al. [32], with slight modifications. Seeds were ground in a knife mill for 4 × 30 s (A11 basic, IKA Werke GmbH & Co. KG, Staufen, Germany) and in a mortar to obtain a fine and homogeneous powder. The seed powder was then subjected to the "coning and quartering" sampling procedure, and three technical replicates were carried out. A 3-g aliquot of fine powdered sample was transferred in a 50-mL tube, mixed with 10 mL of 70% MeOH acidified with HCl (0.1%, *v*/*v*), and kept on an orbital shaker (Duomax 1030, Heidolph Instruments, Schwalbach, Germany) at 200 rpm for 3 h at RT; after a 15-min ultrasound extraction (Elma Schmidbauer GmbH, Singen, Germany), the mixture was centrifuged for 30 min at 4000 rpm, and the supernatant, containing all extractable compounds, was collected. The procedure was repeated twice, and supernatants were combined to form the crude extract (CE).

#### Preparation of Free (F), Base-Hydrolysed (BH), and Acid-Hydrolysed (AH) Soluble Fractions

To obtain the free (F) fraction, the CE was evaporated to dryness, re-suspended with 2 mL of acidified water (pH = 2), and subsequently extracted with 2 mL of a diethyl ether/ethyl acetate mixture (1:1, *v*/*v*) three times. The organic phases were merged, evaporated to dryness, re-suspended in 2 mL of 70% MeOH, filtered through 0.2-µm nylon syringe filters, and stored at −80 ◦C until analysis.

The aqueous phases were also pooled and subjected to base and acid hydrolyses to obtain the base-hydrolysed (BH) and acid-hydrolysed (AH) fractions, respectively. A 2-mL volume of 10 N NaOH was added to 8 mL of aqueous phase, and the mixture was stirred for 1 h under N<sup>2</sup> flow and brought to pH 2 with concentrated HCl. The resulting solution was then subjected to the extraction with diethyl ether/ethyl acetate mixture as previously described. The combined organic phases were evaporated to dryness, re-suspended with 2 mL of 70% MeOH, filtered through 0.2-µm nylon syringe filters, and stored at −80 ◦C until analysis.

The same volume of the aqueous phase was subjected to acid hydrolysis, by adding 1.6 mL of 12 N HCl, and the mixture was stirred for 1 h at 85 ◦C in a water bath. The resulting solution was extracted three times as previously described, and the combined organic phases were evaporated to dryness, re-suspended with 2 mL of 70% MeOH, filtered through 0.2-µm nylon syringe filters, and stored at −80 ◦C until analysis.

#### *3.3. Determination of Total Phenolic Content and Total Flavonoid Content*

The total phenolic content (TPC) assay was carried out using the Folin–Ciocalteu reagent according to the procedure described by Singleton and Rossi [60], with modifications. A mixture containing 100 µL of diluted extract fractions or standard and 440 µL of Folin–Ciocalteu's reagent (diluted 1:10 with water) was incubated for 10 min at RT. Then, 440 µL of 7.5% sodium carbonate was added and the mixture was incubated in the dark for 60 min at RT. Gallic acid was used as a standard, and a calibration curve was built, in the 1–25 ppm range. Absorbance was measured at 765 nm using a double-beam spectrophotometer (V-630 Jasco, Jasco Europe s.r.l., Cremella, Italy) and used to calculate TPC, expressed as mg gallic acid equivalents (GAE) g−<sup>1</sup> DW of seed powder.

The total flavonoid content (TFC) assay was carried out according to the procedure described by Zou et al. [61], with modifications. A mixture containing 100 µL of diluted extract fractions or standard and 440 µL of 0.066 M NaNO<sup>2</sup> was left to react for 5 min at RT. Then, 60 µL of 0.75 M AlCl<sup>3</sup> was added, and the mixture was incubated for 5 min. Lastly, 400 mL of 0.5 M NaOH was added, and the mixture was incubated for 6 min at RT. The absorbance was measured at 510 nm, and the TPC of samples was calculated by interpolating with the calibration curve built with catechin as a standard, in the 1–25 ppm concentration range. Results are expressed as mg catechin equivalents (CE) g−<sup>1</sup> DW of seed powder.

#### *3.4. HPLC Determination of Phenolic Compounds*

The chromatographic method was adapted from Tang et al. [32], with some modifications. The extracts were injected into a Jasco (Tokyo, Japan) HPLC-DAD system, which consisted of a PU-4180 pump, an MD-4015 PDA detector, and an AS-4050 autosampler. The stationary phase was an Agilent (Santa Clara, CA, USA) Zorbax Eclipse Plus C18 reversephase column (100 × 3 mm I.D., 3.5 µm). The mobile phase was a mixture of solvent A (water/formic acid 95/5, *v*/*v*) and solvent B (methanol/acetonitrile 95/5, *v*/*v*), with a composition gradient ranging from 95% to 5% of solvent A and flowing at 0.7 mL/min. Injection volume was 20 µL for all determinations, and analyte detection was carried out with a diode array detector (DAD) by monitoring at 280, 329, and 360 nm. Quantification

was performed with pure standards using calibration curves ranging between 1.25 and 30 µg mL−<sup>1</sup> (r<sup>2</sup> <sup>≥</sup> 0.9634).

#### *3.5. In Vitro Antioxidant Activity Assays*

The ABTS assay was carried out essentially as described by Thaipong et al. [62]. After incubating 950 µL of 7.4 mM 2,2-azinobis(3-ethylbenzothiazoline-6-sulfonic acid) (ABTS) in methanol with either 50 µL of methanolic Trolox solution at different concentrations (0.05–1.00 mM), methanol (blank solution), or the diluted sample in the dark at RT for 2 h, the absorbance of the solution was read at 734 nm in the V630 spectrophotometer. Calibration curves were set up by plotting the discoloration ratio (i.e., [Abs without TX/Abs with TX] − 1) as a function of Trolox concentration. The antioxidant capacity of the sample, expressed as Trolox equivalents (TEs), was calculated by interpolating with the calibration curve. The oxygen radical absorbance capacity (ORAC) assay was carried out on sample fractions essentially as described by Moore et al. [63], using a Viktor X3 multilabel plate reader (Perkin Elmer, Turku, Finland). Trolox equivalents (TEs) were calculated from the relative area under the curve of the emission intensity vs. time plots.

The ferric reducing antioxidant power-ferrozine (FRAP-FZ) assay was performed as described by Mandrone et al. (2015) [64].

#### *3.6. Statistical Analysis*

Statistical analysis was performed using GraphPad (San Diego, CA, USA) Prism Software v. 5.0. Comparison among samples was conducted through one-way analysis of variance (ANOVA) with Tukey's multiple comparison test, and values of *p* ≤ 0.05 were considered statistically significant. Two-way ANOVA was carried out to analyse the genotype x cultivation area interaction.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10.3 390/plants10061046/s1, Table S1: Common and IUPAC names of the phenolic compounds analysed in the study.

**Author Contributions:** Conceptualization, F.A., R.M., and K.B.R.; data curation, F.A., G.P., and R.M.; funding acquisition, F.A., S.B., R.M., and K.B.R.; investigation, G.P. and L.M.; methodology, G.P. and L.M.; supervision, F.A. and R.M.; writing—original draft, F.A. and G.P.; writing—review and editing, S.B. and R.M. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by Ricerca Fondamentale Orientata (RFO) grants from the University of Bologna to F.A., R.M., and S.B. FONDECYT INI 11190834 grant from ANID (Chile) to K.B.R. also funded the research.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Data presented in this study are available upon request.

**Acknowledgments:** The authors would like to thank Dario Vanuzzi (Tuttoquinoa, Italy), S.E. Jacobsen (Copenhagen), Giorgio Dall'Ara (Quinoa Romagna, Italy), Ingrid von Baer and Ricardo Bonert (Semillas Baer, Chile) for kindly providing Titicaca and Regalona seeds. The authors also thank Giuseppe Gabellini and Elena Lombardini (undergraduate students) for their technical contribution.

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

#### **References**

## *Article* **Heat Stress Impact on Yield and Composition of Quinoa Straw under Mediterranean Field Conditions**

**Javier Matías 1,\*, Verónica Cruz <sup>1</sup> and María Reguera <sup>2</sup>**

**Abstract:** Quinoa (*Chenopodium quinoa* Willd.) is receiving increasing attention globally due to the high nutritional value of its seeds, and the ability of this crop to cope with stress. In the current climate change scenario, valorization of crop byproducts is required to support a climate-smart agriculture. Furthermore, research works characterizing and evaluating quinoa stems and their putative uses are scarce. In this work, straw yield and composition, and the relative feed value of five quinoa varieties, were analyzed in two consecutive years (2017–2018) under field conditions in Southwestern Europe. High temperatures were recorded during the 2017 growing season resulting in significantly decreased straw yield and improved feed value, associated with compositional changes under elevated temperatures. Crude protein, ash, phosphorus, and calcium contents were higher under high temperatures, whereas fiber contents decreased. The relative feed value was also higher in 2017 and differed among varieties. Differences among varieties were also found in straw yield, and contents of phosphorus, potassium, and calcium. Overall, the results presented here support a sustainable quinoa productive system by encouraging straw valorization and shedding light on the mechanisms underlying heat-stress responses in this crop.

**Keywords:** quinoa; stems; high temperatures; food security; climate smart agriculture; quinoa by-products

#### **1. Introduction**

In the twenty-first century, climate change has become one of the most critical global challenges, and is particularly crucial for the agricultural and livestock sectors [1–4]. This phenomenon is associated with increments of atmospheric greenhouse gas (GHG) emissions and the increase in the mean global temperature, in addition to alterations in the precipitation regime, affecting food and feed production, and its nutritional composition [1,3,5–8]. Because these changes represent a serious threat to global food security, it is necessary to understand how crops respond to elevated temperatures and how their tolerance to heat stress can be improved [9]. In particular, the Mediterranean climate comprises the western edges of continents between latitudes of 30◦ and 45◦ [10–12].  Because of climate change, severe droughts, increased temperatures, and salinity problems are predicted to be more frequent in the near future in these areas [13].

In addition, the current rapid growth in the global population is expected to lead to a sharp increase in demand for food in coming decades [14]. Furthermore, available land for agriculture and water resources is increasingly scarce following soil degradation and other land uses [4,15]. Hence, agriculture in the twenty-first century must meet increasing food demand with fewer resources, while GHG emissions should be reduced. Within this context, it is necessary to introduce crops with lower irrigation requirements (e.g., in South

**Citation:** Matías, J.; Cruz, V.; Reguera, M. Heat Stress Impact on Yield and Composition of Quinoa Straw under Mediterranean Field Conditions. *Plants* **2021**, *10*, 955. https://doi.org/10.3390/plants 10050955

Academic Editor: Cataldo Pulvento

Received: 9 April 2021 Accepted: 7 May 2021 Published: 11 May 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

Europe alone, more than 60% of freshwater is used by agriculture [13]) and better adapted to less favorable soil conditions, in addition to valorizing crop byproducts. The latter can be achieved if these materials are first well characterized, shedding light on the effects of climate change on the nutritional value of plants.

Quinoa (*Chenopodium quinoa* Willd.) is an ancient crop from South America that has expanded globally because of the increasing interest in the nutritional composition of its seeds. This has contributed to its selection as one of the crops destined to contribute to food security in the next century, according to the FAO (Food and Agricultural Organization) [16]. Quinoa has also been studied for its potential use as a forage crop because of the high nutritional value of the whole plant for livestock [17]. However, this use has been little explored. In contrast, quinoa seeds are well known for being gluten-free, because of their high-quality protein and because they are one of the few plant foods that contain all nine essential amino acids, with a wider amino acid profile than cereals or legumes [18]. Quinoa is an annual C3 plant belonging to the Amaranthaceae family [19], with remarkable adaptability to unfavorable growing conditions [20]. It is a facultative halophytic plant species that is also tolerant to the combined effect of high temperatures and salinity [21–23], and is considered an environmental stress-resilient crop plant [17].

Quinoa has been cultivated for thousands of years in the Andean region comprising Peru, Bolivia, Colombia, and Ecuador although currently its cultivation has expanded globally [6,24]. Indeed, there is a remarkable interest in growing quinoa at European latitudes [25]. In the Mediterranean region, quinoa has been cultivated in Spain, Italy, Greece, and Portugal. In Spain, quinoa cultivation has been expanded significantly in recent years, particularly in the southern part of the country. Interestingly, one of the consequences of the massive expansion of quinoa cultivation is the generation of residues composed of quinoa plant straw, for which possible uses have been little explored [15,26]. Only recently has the use of quinoa straw been evaluated for animal feed [15,17] or generation of renewable energy [15,26].

Thus, in this study, aiming to further explore possible uses of quinoa crop byproducts, and therefore contribute to the implementation of sustainable quinoa cultivation under Mediterranean conditions, the straw production and composition of five quinoa varieties were evaluated in a field located in Southwestern Spain, where episodes of elevated temperatures are frequent. Differences in straw biomass and composition were expected to occur based on the developmental differences of the genotypes tested. Furthermore, the impact of high temperatures under field conditions was analyzed. These results will contribute to a better understanding of how temperature stress might influence straw production.

#### **2. Results**

#### *2.1. Straw Yield and Biomass Partitioning*

As observed in Figure 1, the straw yield varied significantly according to the year and variety. Interaction between both factors was also significant. The mean straw yield was significantly higher in 2018 (2.2 t ha−<sup>1</sup> ) than that in 2017 (1.7 t ha−<sup>1</sup> ), except for Roja and Duquesa, which achieved lower yields in 2018. Marisma reached the highest mean stem yield among varieties (2.8 t ha−<sup>1</sup> ), and was remarkably high in 2018 (3.9 t ha−<sup>1</sup> ). Biomass partitioning was evaluated by determining the HI index, which was significantly influenced by the variety and the year (Figure 2). The average HI in 2017 (0.49) was lower than that in 2018 (0.53), mainly due to the 20% HI reduction achieved in 2017 by Roja (0.40) and Duquesa (0.45). The highest average HI (0.55) was achieved by Jessie (short cycle), whereas the lowest HI was obtained by the varieties with longer life cycles (Roja: 0.48 and Duquesa: 0.50). HI was not correlated with straw yield.

**Figure 1.** Straw yield (t. ha−<sup>1</sup> ) of five quinoa varieties during the two years (2017 and 2018) of field experiments. Error bars represents the standard deviation. Different uppercase letters in the same variety indicate a significant difference between years according to Student's *t*-test at *p <* 0.05. In each year, different lowercase letters indicate significant differences among varieties according to Tukey's test at *p <* 0.05.

**Figure 2.** Harvest index (HI) of five quinoa varieties during the two years (2017 and 2018) of field experiments. Error bars represent the standard deviation (SD). Different uppercase letters in the same variety indicate a significant difference between years according to Student's *t*-test at *p <* 0.05. In each year, different lowercase letters indicate significant differences among varieties according to Tukey's test at *p <* 0.05.

#### *2.2. Crude Protein (CP), Crude Fibre (CF), and Ash Content*

The CP and CF contents were significantly influenced by the year, whereas no significant differences were found according to the variety (Table 1). The CP content in 2017 (12.8%) was 52.3% higher than that in 2018 (8.4%). On the contrary, the CF content was significantly higher in 2018 (33.8%) compared to 2017 (28.0%). The average ash content differed significantly according to the variety. The average ash content varied from 11.5% (Roja) to 16.6% (Jessie) with no different found between years.

**Table 1.** Crude protein (CP), crude fiber (CF), and ash content (%) of straw of five quinoa varieties during the two years (2017 and 2018) of field experiments.

CP<sup>1</sup> : crude protein; CF<sup>2</sup> : crude fiber. Variety means denoted by different lowercase letters in the same column are significantly different at *p* < 0.05 according to Tukey's test. Year means followed by different uppercase letters are significantly different at *p* < 0.05 according to Tukey's test. HSD: critical value for comparison. n.s.: not significant; significant at \* *p* < 0.05; \*\* *p* < 0.01, respectively.

#### *2.3. Fibre Composition*

Significant higher neutral detergent fiber (NDF), acid detergent fiber (ADF), and acid detergent lignin (ADL) contents were determined in 2018 (55.4%; 40.5%; 6.5%, respectively) compared to 2017 (44.1%; 30.8%; 5.2%, respectively), as shown in Table 2. The variety showed a significant influence on the ADF and the ADL content. The average ADF content ranged from 31.6% (Jessie) to 40.9% (Roja), whereas the average ADL content varied from 5.3% (Marisma) to 6.5% (Roja). Interactions between the year and variety were not significant. The hemicellulose (HEM) contents only showed significant differences in 2017 according to the variety, ranging from 11.2% (Marisma) to 15.2% (Jessie). Significant differences were also found in cellulose (CEL) contents in 2017 according to the variety, ranging from 20.6% (Jessie) to 29.2% (Pasto). The mean CEL content was significantly affected by the year and the variety. It was higher in 2018 (34.0%) than in 2017 (25.6%), and ranged from 25.3% in Jessie to 34.5% in Roja. Neither the HEM nor the CEL contents were significantly affected by the year × variety interaction.

**Table 2.** Fiber composition of straw of five quinoa varieties during the two years (2017 and 2018) of field experiments.

NDF<sup>1</sup> : neutral detergent fiber, ADF<sup>2</sup> : acid detergent fiber, ADL<sup>3</sup> : acid detergent lignin, HEM<sup>4</sup> : hemicellulose, CEL<sup>5</sup> : cellulose. Variety means denoted by different lowercase letters in the same column are significantly different at *p* < 0.05 according to Tukey's test. Year means followed by different uppercase letters are significantly different at *p* < 0.05 according to Tukey's test. HSD: critical value for comparison. n.s.: not significant; significant at \* *p* < 0.05; \*\* *p* < 0.01, respectively.

### *2.4. Mineral Composition*

As can be observed in Table 3, the N, P, and Ca contents differed significantly according to year, and all of them were higher in 2017. The variety had a significant influence on the P, K, and Ca contents, but not on the N and Mg average contents, which were 1.7% and 0.64%, respectively. The average P content ranged from 0.17% (Roja and Duquesa) to 0.24% (Jessie). K showed average contents from 4.4% (Roja) to 6.0% (Jessie), and the average of Ca content varied between 0.8% (Roja) and 1.6% (Pasto).

**Table 3.** Mineral composition of straw of five quinoa varieties during the two years (2017 and 2018) of field experiments.

Variety means denoted by different lowercase letters in the same column are significantly different at *p* < 0.05 according to Tukey's test. Year means followed by different uppercase letters are significantly different at *p* < 0.05 according to Tukey's test. HSD: critical value for comparison. n.s.: not significant; significant at \* *p* < 0.05; \*\* *p* < 0.01 and \*\*\* *p* < 0.001, respectively.

> Interactions between year and variety were significant for the P and Ca contents. In addition, the P content was relatively similar in both years in the short cycle variety (Jessie), whereas in the other varieties, the P content was considerably higher in 2018, especially in Roja (medium-long cycle), for which differences were found to be higher than 60%. Regarding the Ca content, differences between years were small in Jessie and Pasto, whereas in 2018 the Ca content decreased by more than 18% in the other varieties.

#### *2.5. Relative Feed Value*

The year showed a significant influence on the digestible dry matter (DDM), the dry matter intake (DMI) and on the relative feed value (RFV), which were higher in 2017 (Table 4). The variety significantly influenced the DMI and RFV values, achieving the highest values in the short cycle variety Jessie (64.3 and 131.0, respectively), and the lowest values in the medium-long cycle varieties (Roja; 57.1 and 99.4, respectively). Interaction between the year and the variety was not significant.

**Table 4.** Straw forage quality of five quinoa varieties during the two years (2017 and 2018) of field experiments.

DDM<sup>1</sup> : digestible dry matter; DMI<sup>2</sup> : dry matter intake; RFV<sup>3</sup> : relative feed value. Variety means denoted by different lowercase letters in the same column are significantly different at *p* < 0.05 according to Tukey's test. Year means followed by different uppercase letters are significantly different at *p* < 0.05 according to Tukey's test. HSD: critical value for comparison. n.s.: not significant; significant at \* *p* < 0.05; \*\* *p* < 0.01, respectively.

#### **3. Discussion**

The use of quinoa straw has the potential to contribute to the development of a cleaner agriculture. This byproduct has been used and studied for agricultural purposes as feed and bedding material, in addition to bioenergy or biomaterial production [27,28]. Quinoa cultivation has experienced a rapid increase in the past decade, in parallel with its expansion to many different geographical areas around the world [29]. However, quinoa straw use has been little explored because the interest in this crop has focused mainly on seed yield. Thus, the current study aimed to evaluate variations in quinoa

straw characteristics linked to genetic differences under Mediterranean field conditions, to implement alternative uses of this crop and therefore contribute to the development of sustainable agriculture. This study was performed in two consecutive years showing different climatic conditions, which included average differences in Tmax during the quinoa growth period of 5 ◦C. Generally, elevated temperatures inhibit quinoa plant growth by impacting quinoa flowering [19,30–32]. Heat stress can limit the source and sink capacity of plants, reducing growth and development [33]. Indeed, the higher temperatures registered during the first year resulted in a reduction of straw yield in 2017 (1.7 t ha−<sup>1</sup> ) compared to 2018 (2.1 t ha−<sup>1</sup> ) (Figure 1), especially in short-medium cycle varieties, such as Marisma, which doubled its yield in 2018. On the contrary, long-cycle varieties, such as Roja or Duquesa, showed lower straw yields in 2018, which may point to a differential response depending on the developmental stage affected by high temperatures. In cereals, such as wheat, it has been reported that heat stress is an important factor that reduces straw yield, because photosynthesis is altered [34,35]. Nonetheless, further analysis should be performed to determine the exact impact of heat stress on the vegetative growth and its influence on straw yield, particularly in quinoa. It should be noted that the high straw yield achieved by Marisma (2.8 t ha−<sup>1</sup> , on average), especially in 2018 (3.9 t ha−<sup>1</sup> ), was well correlated with a higher seed yield, as previously reported [36], which is interesting from an agronomical perspective because seed yield penalties should be avoided. Intriguingly, the two-year average straw yield (1.9 t ha−<sup>1</sup> ) achieved in this work was lower than those previously reported for Mediterranean field conditions (3.0 t ha−<sup>1</sup> for Titicaca; 7.4 t ha−<sup>1</sup> for Regalona) [37], and similar to those obtained by Asher et al. (2020), although at the bottom of the range (0.5–9.1 t ha−<sup>1</sup> ). The results reported earlier can be partially explained by the lower HI obtained in these works, which resulted in higher straw yield per seed yield. This was probably a consequence of the different environmental conditions, and differences in the planting density and/or in the genotypes used, that resulted in detrimental effects on seed yield but positive effects on vegetative growth [38,39]. In the current study, the higher temperatures registered in 2017 also had a significant impact on the HI in medium-long cycle varieties (Roja and Duquesa). Due to the longer vegetative stage of Roja and Duquesa, flowering coincided with higher Tmax and lower RH, increasing the probability of flower damage causing a HI decrease (Figure 2). On the contrary, the shortest-cycle genotype Jessie showed the highest HI (0.55) in 2017, which was related to its ability to escape from high temperatures at flowering stage, which resulted in the maintenance of its sink capacity but larger straw yield penalties. In general, as shown in Figure 2, the average HI was 0.51, which is similar to the HI reported in modern wheat varieties (0.3–0.6) [39,40], and higher than those reported for grain legumes and canola [41]. Nonetheless, no correlation between straw yield and HI was found, in contrast to the findings of previous studies [42,43].

The straw composition differed between years as observed in Tables 1–4, probably influenced by the high temperatures of 2017. This effect has been previously studied in the main products of different crops [44–48], although the impact on byproducts, such as straw, has been little explored. In fact, the current study is pioneering in its evaluation of the effect of high temperatures on quinoa straw yield and composition. Furthermore, the CP content range in the straws obtained in this study (6.9–13.5%) was slightly higher than that obtained by Asher et al. (2020) under Mediterranean conditions (5.1–10.6%).

As can be observed in Table 1, the CP content was considerably higher in 2017 (52.7%). Because of the higher average straw yield of 2018, the lower straw CP content achieved in 2018 could have been a consequence of a dilution effect. However, in 2017 the CP content was also higher in the medium-long cycle varieties, which also achieved higher straw yields. Therefore, high temperatures could lead to higher straw CP content independently of the straw yield. Interestingly, no correlation was found between CP content and the other parameters analyzed in this study, except for the straw N content. It should be noted that the high CP content observed here (particularly in 2017) was also higher than the contents found in cereal (2.9%) or legume (7.4%) straws [49], which is important when the aim is to use this byproduct for animal feed. In contrast, the CF content was

reduced about 20% in 2017 without a significant effect linked to the variety (Table 1). Plant fiber includes the cell wall used for providing mechanical support to the plant, and the vascular system in which fluids are transported [50,51]. The response observed in the CF content was likely related to a change in composition due to the heat stress impact on the biosynthesis-related pathways of cell wall components [52–54]. Actually, a steeper decrease was detected in the ADF content (with an average reduction of 24%), and in the cellulose content (which showed an average decrease of 25%) (Table 2). The two-year average of NDF was slightly lower than the values determined by Asher et al. 2020 (41.4–63.6% and 44.4–71.3%, respectively), whereas the lignin was higher than the values reported by these authors (5.3–7.0% and 5.03–7.83%, respectively). This effect raises an interesting aspect related to the straw composition as lignocellulosic biomass, which is now considered an important fiber resource for renewable energy and biomaterial production [55]. By comparison, the ash content did not change between years, achieving values close to 15%, restricting quinoa straw's use as a solid biofuel (for which ash should not exceed 1.5%) [56].

When analyzing the increased mineral and protein contents in 2017, the changes could point to a detrimental effect of higher temperatures in nutrient translocation into the seed as observed in wheat [33]. However, the protein and mineral contents were also higher in quinoa seeds in 2017 [36]. It is known that the environmental conditions can affect nutrient levels of quinoa seeds [57]. In the current study, the straw N and P contents were significantly lower in 2018, which could be related to a dilution effect linked to higher yields, similar to that described in cereal grains [58]. However, those differences were even higher in Roja and Duquesa, which achieved higher yields in 2017. Furthermore, the straw Ca content was significantly lower in 2017, especially in Roja and Duquesa. Therefore, differences in mineral nutrient contents were probably more related to a still unknown heatinduced adaptation mechanism (and/or to the effect caused by the interaction between nutrients) than to a direct effect caused by nutrient dilution. The effect of heat stress on cellular osmotic adjustments due to the increased transpiration rates when elevated temperatures occur should also not be discounted; this could result in increments in the mineral and protein content [59,60]. Interestingly, the mineral composition of quinoa straw was richer from a nutritional perspective compared with the mineral composition of cereal straws commonly used for animal feed [61]. For instance, Ca or Mg contents in wheat straws were reported to be, on average, 0.18% and 0.06%, respectively whereas quinoa contents were, on average, 1.2% and 0.64%, respectively (Table 3).

The relative feed value (RFV) developed by the Universities of Minnesota and Wisconsin and the American Forage and Grassland Council (AFGC), is an index widely used to determine the forage quality, which combines significant nutritional factors (including voluntary intake and digestibility) [62,63]. When evaluating the changes in composition of the quinoa straw, a significant impact on the RFV was detected, which resulted in the increase in this index in 2017, due to the lower NDF and ADF, improving the straw nutrient intake and digestibility (Table 4). Considering the AFGC classification method [62], the mean value for the RFV in 2017 (138.5) would indicate that this byproduct would be classified as a forage type I (125–151), which is considered acceptable. In 2018, the mean value (98.5) would correspond to a low-quality forage (being classified as a type IV forage (87–102). Therefore, based on these results, in addition to the higher CP and mineral contents (higher than those found in winter cereals, which is the straw most frequently used in animal feed), it can be considered that the quinoa straw obtained in 2017 possesses a higher nutritional value. Nonetheless, further research is required to determine the exact impact of heat stress on straw yield and composition. Based on the results presented here, elevated temperatures, which are expected to continue increasing due to the continuation of the warming period [64], may positively impact straw composition in a genotype-dependent manner.

Overall, the results here presented show that Marisma is the variety with the best cultivation potential in terms of straw yield for this particular area (Southwestern Spain), despite suffering important yield penalties linked to the elevated temperatures suffered in 2017 [36]. Furthermore, the straw composition analysis performed shows that quinoa

straw is a valuable resource for animal feed, biofuel, or biomaterial production. To enhance quinoa straw valorization, it is crucial to evaluate the straw yield and compositional changes related to the genotype and environment, and to understand the relationships among biomass, seed, and straw yield. Our results, together with the previously reported changes in chemical composition of cereal straws, highlight that quinoa straw yield and composition may be affected by agronomic and genotypic factors, and environmental and climate conditions [50,55]. Within the current climate context in which more frequent episodes of elevated temperatures in the Mediterranean area are expected to occur, and considering that the effects of heat stress on plants trigger complex responses that result in the alteration of growth and development, thereby changing physiological functions and reducing seed/grain formation and plant yield [33,65], further studies should be undertaken to evaluate the impact of heat stress on straw yield and composition, with the aim of selecting the best adapted quinoa cultivars for a particular area of cultivation. More importantly, this work points to the many possibilities offered by the use of quinoa straw as an agricultural byproduct that can greatly contribute to sustainable agriculture.

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

#### *4.1. Location, Climate and Soil Characteristics of the Experimental Site*

A two-year field experiment was conducted during 2017–2018 at the experimental farm of the Center for Scientific and Technological Research of Extremadura (CICYTEX), located in Southwest Spain (lat. 38◦5101000 N; long. 6◦3901000 W). Data of monthly mean minimum and maximum temperature (Tmin and Tmax), and the rainfall during the crop cycle (Supplementary Figure S1) were obtained from the weather station located at the experimental farm. The soil was a sandy loam, neutral (pH 6.9), presenting 0.38% organic matter, 0.24% total N, and 93.4 ppm and 57.9 ppm of available P and K, respectively.

#### *4.2. Layouts of Experiments, Plant Material and Crop Management*

Five European varieties of quinoa (Pasto, Marisma, Jessie, Roja, Duquesa) were evaluated in a randomized complete block design with four replications: Jessie (short cycle; ~120 d); Pasto and Marisma (medium cycle; ~135 d); Roja and Duquesa (medium-long cycle; ~145 d). The plot size was formed by four rows 0.75 m apart and 10 m long. Sowing was conducted mechanically in early February, at a dose of 4 kg ha−<sup>1</sup> . Weeding was carried out by hand when required. Irrigation was carried out by sprinkling to maintain the soil under non-limiting water conditions. The crop was fertilized at the rate of 150, 100, and 100 kg/ha of N, P2O5, K2O, respectively. Plants were harvested by hand at physiological maturity (in the middle of June for Jessie, and early July for the other four varieties). The sampling area was 3 m<sup>2</sup> . A stationary thresher was used to collect the seed. The straw samples were ground through a 1 mm screen for further analysis.

#### *4.3. Analysis and Measurements*

Data were expressed on a dried weight (dW) basis. Analysis of moisture, crude protein (CP), and ash contents were conducted following the AOAC Official Methods [66]. The mineral content was assessed following the official methods of the Spanish Ministry of Agriculture [67]. P was analyzed by a UV-VIS spectrophotometer (Hitachi U-2810). K was determined using flame atomic emission spectroscopy, and Ca and Mg by flame atomic absorption (SpectrAA 110, Agilent). The crude fiber (CF), neutral detergent fiber (NDF), acid detergent fiber (ADF), and acid detergent lignin (ADL) values were analyzed following the Ankom procedure (ANKOM Technology, Fairport, NY, USA), using a fiber analyzer (ANKOM 2000) and F57 Ankom filter bags (porosity: 25 µm). Hemicellulose (Hem) and cellulose (Cel) contents were calculated as follows:

Hemicellulose (Hem) = [NDF% − ADF%]

Cellulose (Cel) = [ADF% − ADL%]

Relative feed value (RFV) was calculated as calculated from the digestible dry matter (DDM) and the dry matter intake (DMI) (live weight: LW, %) according to the following equations:

$$\text{DDM} \left( \% \right) = \text{88.9} - \left[ 0.779 \times \text{ADF\%} \right]$$

$$\text{DMI} \left( \text{Live Weight}: \text{ LW\%} \right) = 120/[\text{NDF\%}]$$

$$\text{RFV} = \left[ \text{DMD} \times \text{DMI} \right] / 1.29$$

Harvest index (HI) was calculated as the ratio between the grain yield (G) and the yields of straw plus grain (G + S), in order to determine the biomass partitioning index, as previously reported [36].

#### *4.4. Statistical Analysis*

All measured and derived data were processed using a two-way analysis of variance (ANOVA), including the year, the variety, and their interactions in the model. The year was treated as a fixed factor. Normality and equal variances could be assumed, according to the results of the Shapiro Wilk test and Levene's test, respectively. For better interpretation of the data, additional one-way ANOVA was carried out for each year separately. When the F ratio was significant (*p* < 0.05), Tukey's test was performed and used to compare means. Student's *t*-test was used to determine statistically significant differences between years for each variety. Analyses were performed using the Statistix 8 analytical software.

**Supplementary Materials:** The following are available online at https://www.mdpi.com/article/10 .3390/plants10050955/s1, Figure S1: Monthly rainfall and maximum (Tmax) and minimum (Tmin) mean temperature registered at the experimental station during the quinoa growth period of 2017 and 2018.

**Author Contributions:** Conceptualization, J.M. and M.R.; methodology J.M., V.C. and M.R.; software, J.M., V.C. and M.R.; validation, J.M. and M.R.; formal analysis, J.M. and M.R.; investigation, J.M., V.C. and M.R.; resources, J.M., V.C. and M.R.; data curation, J.M., V.C. and M.R.; writing—original draft preparation, J.M., V.C. and M.R.; writing—review and editing, J.M., V.C. and M.R.; visualization, J.M., V.C. and M.R.; supervision, J.M. and M.R. All authors have read and agreed to the published version of the manuscript.

**Funding:** The authors gratefully acknowledge the financial support received from the Junta de Extremadura, the European FEDER Funds (MESOCEX), the Ministerio de Ciencia e Innovación (MICINN, Spain) (PID2019-105748RA-I00) and the Comunidad de Madrid-Universidad Autónoma de Madrid (SI1/PJI/2019-00124).

**Data Availability Statement:** The data that support the findings of this study are available upon request.

**Acknowledgments:** The authors greatly thank Susana Vilariño (Algosur, Spain) for her support helping to conceive and plan the experiments and for providing the quinoa seeds used in this study.

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

#### **References**

## *Article* **The Importance of Non-Diffusional Factors in Determining Photosynthesis of Two Contrasting Quinoa Ecotypes (***Chenopodium quinoa* **Willd.) Subjected to Salinity Conditions**

**José Delatorre-Herrera 1,\*, Karina B. Ruiz <sup>2</sup> and Manuel Pinto <sup>3</sup>**

**\*** Correspondence<EMAIL_ADDRESS>
**Abstract:** The broad distribution of quinoa in saline and non-saline environments is reflected in variations in the photosynthesis-associated mechanisms of different ecotypes. The aim of this study was to characterize the photosynthetic response to high salinity (0.4 M NaCl) of two contrasting Chilean genotypes, Amarilla (salt-tolerant, *salares* ecotype) and Hueque (salt-sensitive, coastal ecotype). Our results show that saline stress induced a significant decrease in the K+/Na<sup>+</sup> ratio in roots and an increase in glycine betaine in leaves, particularly in the sensitive genotype (Hueque). Measurement of the photosynthesis-related parameters showed that maximum CO<sup>2</sup> assimilation (Amax) in control plants was comparable between genotypes (ca. 9–10 µmol CO<sup>2</sup> m−<sup>2</sup> s −1 ). However, salt treatment produced different responses, with Amax values decreasing by 65.1% in the sensitive ecotype and 37.7% in the tolerant one. Although both genotypes maintained mesophyll conductance when stomatal restrictions were removed, the biochemical components of Amarilla were impaired to a lesser extent under salt stress conditions: for example, the maximum rate of ribulose-1,5-bisphosphate carboxylase/oxygenase (RubisCO; Vcmax) was not as affected in Amarilla, revealing that this enzyme has a higher affinity for its substrate in this genotype and, thus, a better carboxylation efficiency. The present results show that the higher salinity tolerance of Amarilla was also due to its ability to control non-diffusional components, indicating its superior photosynthetic capacity compared to Hueque, particularly under salt stress conditions.

**Keywords:** Na<sup>+</sup> , K<sup>+</sup> , CO<sup>2</sup> assimilation; stomatal restrictions; non-diffusional; diffusional; RubisCO activity

### **1. Introduction**

At present, about one-third of the world's irrigated land [1] is affected by salinity, which reduces plant growth and crop yield [2]. As the main food crops are rather sensitive to this stress, tolerance to salinity has become an important agronomical trait for breeders, physiologists, and agronomists [3]. The salinity of soils is predominantly caused by salt transported by irrigation water [4]. On the other hand, due to climate change, well water is becoming the main water resource for irrigation in many parts of the world, and several studies have indicated that this method is linked to potential risks of soil salinization [5–7]. An increase in the soil concentration of Na<sup>+</sup> and other ions, such as Ca2+, Cl−, and K<sup>+</sup> , causes a decrease in the soil water potential, which limits the water absorbed by roots and induces water stress [8]. Specific ion toxicities are due to the accumulation of sodium, chloride, and/or boron in the tissue of transpiring leaves to damaging levels. The accumulation of injurious ions may inhibit photosynthesis and protein synthesis, inactivate enzymes, and damage chloroplasts and other organelles [9]. Under normal physiological conditions,

**Citation:** Delatorre-Herrera, J.; Ruiz, K.B.; Pinto, M. The Importance of Non-Diffusional Factors in Determining Photosynthesis of Two Contrasting Quinoa Ecotypes (*Chenopodium quinoa* Willd.) Subjected to Salinity Conditions. *Plants* **2021**, *10*, 927. https://doi.org/10.3390/ plants10050927

Academic Editors: Cataldo Pulvento and Didier Bazile

Received: 19 March 2021 Accepted: 27 April 2021 Published: 6 May 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 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 (https:// creativecommons.org/licenses/by/ 4.0/).

<sup>3</sup> Plant Physiology Laboratory, Institute of Agronomic and Veterinary Sciences, Universidad de O´Higgins, Rancagua 2820000, Chile<EMAIL_ADDRESS>
plants maintain a high cytosolic K+/Na<sup>+</sup> ratio. Given the difference in negative membrane potential at the plasma membrane (–140 mV), a rise in extracellular Na<sup>+</sup> concentration will establish a large electrochemical gradient that favors the passive transport of Na<sup>+</sup> into cells through the activation of K<sup>+</sup> transporters/channels and through non-selective channels that are sensitive to Ca2+. The permeation of Na<sup>+</sup> via voltage-independent cation (VIC) channels is inhibited by an increase in extracellular Ca2+ concentration [10]. Overaccumulation of Na<sup>+</sup> in the cytosol inhibits protein synthesis, enzyme activity [1], and many photosynthetic processes [11,12]. Therefore, maintaining its water supply and excluding Na<sup>+</sup> from photosynthetic organs are crucial mechanisms used by tolerant plants to ensure an adequate rate of carbon fixation under salt stress [13]. It is well known that a reduction in stomatal conductance negatively affects the CO<sup>2</sup> assimilation rate as well as the water balance in leaves [3,14].

Excess salt affects plant growth due to an increase in soil osmotic pressure and interference with plant nutrition. A high salt concentration in the soil solution reduces the ability of plants to acquire water, which is referred to as the osmotic or water-deficit effect of salinity [9]. A decrease in CO<sup>2</sup> assimilation is a widely reported effect of salt [15]. In many cases, this decrease is associated with the negative effect of salinity on diffusional mechanisms. These mechanisms depend on the gradient between the external and internal CO<sup>2</sup> concentrations (mesophyll and chloroplasts), on the stomatal conductance in the gaseous phase (gs), and on the mesophyll conductance in the liquid phase (gm) [15–18].

However, salt also affects non-diffusional mechanisms, which are typically associated with electron transport and the activity of enzymes in carboxylation cycles; key enzymes in these processes include ribulose-1,5-bisphosphate carboxylase/oxygenase (RubisCO) [19,20] and those involved in the regeneration of ribulose-1,5-bisphosphate (RuBP) [21,22].

Most authors agree that diffusional factors are predominant in affecting the CO<sup>2</sup> assimilation rate [15,23]. For example, Flexas et al. [24] reported that the conductance of the mesophyll in grapevine was strongly correlated with the rate of photosynthesis. In olive trees (*Olea europaea*), this occurs in the leaves of stressed plants with a conductance greater than 50 µmol m−<sup>2</sup> s −1 [22]. According to the aforementioned study, the combined reduction in stomatal and mesophyll conductance in different salt-stressed olive cultivars increases the difference in CO<sup>2</sup> concentration between the environment and chloroplasts. These results indicate that the low concentration of CO<sup>2</sup> in chloroplasts caused by decreased stomatal and mesophyll conductance is the main limiting factor of photosynthesis.

Quinoa (*Chenopodium quinoa* Willd.) has evolved a number of adaptive responses to saline stress conditions, such as the increased elasticity of cell walls, low water potential, osmotic adjustment, reduction in the foliar area, the presence of *papillae* with calcium oxalate, the production of organic solutes [25–28], the adaptation of physiological functions such as photosynthesis [2,29], the regulation of the water status [29], ionic partitioning [30,31], and changes in stomatal conductance [3,16,27].

There are an increasing number of studies on CO<sup>2</sup> assimilation in quinoa [2,16,26,32,33], although few have characterized the importance of diffusional and non-diffusional mechanisms. This species is particularly diverse at the genetic level [34], a consequence of its broad distribution in both saline and non-saline environments. This diversity could allow some ecotypes to thrive under saline stress conditions due to improved rates of photosynthesis [30].

Thus, it is possible to postulate that saline-tolerant ecotypes have better control of CO<sup>2</sup> diffusion mechanisms than those that are less tolerant to this stress. Therefore, the aim of this study was to evaluate the differences in the photosynthetic processes between two contrasting quinoa ecotypes, as well as to analyze the relative importance of non-diffusional factors in the two ecotypes when subjected to salinity conditions.

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

#### *2.1. Plant Material*

The two quinoa ecotypes used in this study, Amarilla and Hueque, have been previously characterized as tolerant and sensitive to salinity, respectively [30]. The Amarilla ecotype comes from the salt flat area in the northern highlands, and Hueque originates from the wet area at sea level in southern Chile.

Disinfected seeds (2% sodium hypochlorite for 7 min) were germinated in 330 mL pots using perlite as the substrate. Pots containing 4 plants each were arranged over a tray containing water or solution nutritive and connected to it by means of a cotton wick to ensure water supply by capillarity. The plants were watered with sodium-free nutrient solution or 0 M (0.25 dS m−<sup>1</sup> ) until the third pair of leaves developed (45 days after sowing (DAS)). At this stage, half of the pots of each ecotype were transferred to a modified Hoagland 2 solution containing 0.4 M NaCl (38.1 dS m−<sup>1</sup> ). This concentration corresponds to the LD50max for quinoa, as determined previously by Delatorre-Herrera and Pinto [30]. To avoid a saline shock, 0.4 M NaCl was applied in increments of 0.1 M per day. Control plants were irrigated with solution nutritive in a semi-hydroponic system (Schlick and Bubenheim [35], modified by Delatorre and Pinto [30]); for this purpose, filtered water with an electrical conductivity of 0.25 dS m−<sup>1</sup> with a pH of 6 was used. When the plants developed their fourth or fifth pair of true leaves (70–75 DAS), they were placed in a shaded field, where the average temperature of the day was 20 ± 5 ◦C, and the maximum light intensity was 1500 µmol m−<sup>2</sup> s <sup>−</sup><sup>1</sup> of photosynthetically **a**ctive **r**adiation (PAR).

#### *2.2. Na<sup>+</sup> and K<sup>+</sup>*

Sodium (Na<sup>+</sup> ) and potassium (K<sup>+</sup> ) contents were determined in samples of root, stems, and leaves taken from the bottom, middle, and top parts of the stem. Once collected, the material was dried at 70 ◦C for 48 h and then finely ground. Na<sup>+</sup> and K<sup>+</sup> were extracted by digesting 0.1 g of each sample in 15 mL of 0.5 M HCl for 2 days. The concentrations of Na<sup>+</sup> and K<sup>+</sup> were determined according to the procedure described by Hunt [36] using a flame photometer (Jenway Model PFP 7, Cole Parmer, Vernon Hills, IL, USA).

#### *2.3. Proline and Glycine Betaine*

Proline (Pro) content was determined spectrophotometrically (Spectrophotometer, Genesys, Thermo Scientific, Waltham, MA, USA) in tissue with a dry weight of 100 mg using the method described by Bates et al. [37]. A standard curve was constructed with proline (M.W. 115.13 g mol−<sup>1</sup> , Sigma-Aldrich, Santiago, Chile).

The content of Glycine Betaine (GB) was determined according to Grieve and Grattan [38] but modified for quinoa. Briefly, tissue with a dry weight of 100 mg was stirred in 4 mL of water for 24 h at 25 ◦C, after which it was filtered and stored at 4 ◦C until analysis. For the determination of quaternary compounds, 50 µL samples were thawed, diluted in 50 µL of 2 N sulfuric acid, and cooled to 0 ◦C for 1 h. Then, 40 µL of KI-I<sup>2</sup> reagent (15.7 g iodine and 20 g KI in 100 mL of water) was added and gently stirred in a vortex. This solution was stored at −4 ◦C for 16 h, after which it was centrifuged at 10,000 rpm for 15 min. The supernatant was carefully removed, and the precipitate was dissolved in 1.6 mL of 1,2-dichloroethane, shaken vigorously by vortexing, and left to stand for 2.5 h at room temperature. GB content was detected at 365 nm. The standard curve was constructed with betaine (M.W. 117.15 g mol−<sup>1</sup> , Sigma-Aldrich, Santiago, Chile).

### *2.4. Determination of CO<sup>2</sup> Assimilation*

#### Gas Exchange

Gas exchange measurements were performed between 09:00 and 15:00 using a gas exchange chamber connected to a portable infrared analyzer (Licor 6200, LI-COR, Lincoln, NE, USA). The relative humidity of the chamber ranged from 40% to 50%, and the leaf temperature was 25 ± 2 ◦C. Measurements were made on fully expanded mature leaves (middle part of the stem). Each measurement was repeated three times on the same leaf

(one leaf per plant and 6 plants per treatment). These measurements were taken when 50% of the plants had grown the fourth or fifth pair of true leaves (70–75 DAS).

#### (i) A/PFD curves

The photon flows (PFDs) used were 0, 20, 90, 120, 150, 300, 500, 700, 1200, 1500, and 2500 µmol photons m−<sup>2</sup> s −1 . The light source was a halogen lamp, and the different PFDs were obtained by placing neutral filters between the lamp and the photosynthetic chamber.

The temperature in the photosynthetic chamber was 25 ± 2 ◦C. During measurements, the CO<sup>2</sup> and the O<sup>2</sup> concentrations in the chamber were maintained at 360 µL L−<sup>1</sup> and 20%, respectively (CO<sup>2</sup> and O2, certified gas, INDURA S.A, Alto Hospicio, Chile).