1. Introduction
Buckwheat is a pseudocereal belonging to the genus Fagopyrum in the family Polygonaceae. The two most commonly cultivated species are common buckwheat (F. esculentum) and Tartary buckwheat (F. tartaricum). In agriculture, its importance as an environmentally friendly crop that contributes to biodiversity is evident in its ability to improve the biological and physical conditions of the soil and enhance the sustainability of agricultural systems. This is achieved through its use as a cover crop and forecrop, and in crop rotations [1]. Additionally, buckwheat reduces soil acidification, increases the availability of soil phosphorous and nitrogen as an efficient green fertilizer, plays a significant role in carbon cycle regulation, and can be utilized in the phytoremediation of lead, aluminum, mercury, and cadmium. It also prevents the spread of root and plant diseases, providing an effective and sustainable method for pest suppression while improving soil health and crop yields [1,2,3,4].
Buckwheat (Fagoppyrum esculentum Moench) is an ancient annual plant with seeds that are used in both human and animal nutrition. During flowering, it produces up to 2000 flowers per plant, each containing a large amount of nectar, making it a honey-bearing plant. This nectar contributes to the increased content of organic acids, phenolic compounds, and flavonoids in honey [5]. Buckwheat yield and biodiversity are unaffected by bee hives, as wild pollinator density remains constant, indicating no resource competition due to the abundance of flowers [6]. Buckwheat seeds are rich in carbohydrates, proteins, lipids, essential fatty acids, minerals, vitamins, and essential amino acids, especially lysine (5.1 g/100 g protein), and are gluten-free [7]. The grains have extremely favorable nutritional components; the proteins contain all the essential amino acids necessary for human health [8]. Buckwheat seeds are also high in rutin, a bioflavonoid formerly known as vitamin P, which contributes to the use of buckwheat flour in food fortification [9]. The application of different levels of nitrogen fertilizer increased the number and surface area of endosperm cells, as well as the content of amylose, amylopectin, and starch [10]. Amylose content is a key factor in seed quality, while amylopectin determines the properties of buckwheat starch. Due to its smaller granule size, buckwheat starch is often used as a thickener, binder, film, or foam in food and other industries [11]. Buckwheat bran, a by-product of processing, contains various natural compounds, primarily fibers, proteins, flavonoids, and phenols (about 40%) [12]. Studies on bran and endosperm have confirmed the presence of natural antioxidants of a phenolic type [13], while numerous investigations have examined phenolic acids and bioflavonoids in buckwheat flour [14].
Currently, buckwheat holds a prominent place in the functional food industry and on the market. Buckwheat products, such as whole seeds, husks, kernels or groats (unroasted or roasted), and flour, are traditionally consumed in breakfast cereals and bakery products or added to enriched and fortified products such as bread, dough, pasta, pancakes, cookies, biscuits, noodles, sprouts, extruded snack food, honey and tea [15,16,17]. Buckwheat seeds and their processed products are gluten-free making them highly valuable for the development of products with eliminated ingredients containing gluten proteins. Additionally, in the modern meat industry, incorporating buckwheat into processed-meat products has gained importance as a means of improving product quality. Researchers have found that adding buckwheat to meat products enhances the physical, chemical, and sensory properties of various meat products, including semi-smoked sausages, frankfurters, horsemeat, chicken patties, and pork meatballs [18,19,20,21]. Furthermore, buckwheat is recognized for its unique characteristics, contributing favorable flavor, texture, and color to food products [22]. As it contains flavonoids, which are pigments, buckwheat can impart color and initiate color–sensory changes in many food end-products [19,23].
The importance of buckwheat and its products is evident in their various health-related benefits, including hypoglycemic, anticancer, hypocholesterolemic, anti-hypertensive, and anti-inflammatory properties. These attributes significantly increase buckwheat’s value in agricultural, industrial, and pharmaceutical applications [17,20,24,25].
Additionally, the widespread use of buckwheat in daily consumer lifestyles is seen in the production of alcoholic beverages and fermented products such as vinegar [22].
In 2022, global buckwheat seed production was estimated at 2,235,193.31 tons per year (yield of 999,400 g/ha), with Russia, China, Ukraine, and Kazakhstan (1,222,381.54, 506,439.75, 147,690, and 89,802.66 tons per year, respectively) as the leading producers (FAOSTAT, 2022). In these countries buckwheat is primarily processed into traditional foods, such as groat porridge, noodles, and pasta. As the global demand for nutritionally justifiable and health-beneficial food continues to rise, ensuring the long-term storage of raw cereals, including the pseudocereal buckwheat, becomes increasingly important for their later application in the food industry. The natural aging process of grain or seed leads to physiological degradative changes in biological, physical, and chemical properties. Storage conditions play a critical role in preserving nutritional characteristics over extended periods and may influence the extent of these changes [26]. Lipid degradation, in particular, negatively impacts the taste and flavor of the grain or seed, reducing its utility and shelf life [27]. While numerous studies have addressed this issue [28,29,30,31], to the best of our knowledge, the Novosadska variety has not yet been studied in this context.
In this context, the experiment aimed to (i) analyze and evaluate the physicochemical, chemical, nutritional, and antioxidative properties, as well as the phenolic acid and bioflavonoid profiles of stored Novosadska variety buckwheat, by investigating potential losses and changes during long-term storage (3, 6 and 9 months); and (ii) propose the optimal storage time for Novosadska variety buckwheat seeds to preserve the most nutritious components, leading to more technologically justified and health-beneficial end-products. This study on Novosadska variety buckwheat as an alternative crop was conducted with a focus on sustainable agricultural utilization and the potential for extended cultivation of biodiversity-friendly crops in farming fields. Only a few published studies have examined both the sustainable utilization and the storage impact on buckwheat, emphasizing its value in sustainable agriculture and food technology [32,33]. However, to the best of our knowledge, no study has specifically investigated the Novosadska variety in this manner.
2. Materials and Methods
2.1. Materials and Storage Conditions
The buckwheat (Novosadska variety) used in this investigation, a locally significant variety of alternative crops and pseudocereal, was harvested in 2022 at the seed’s technological maturity. The raw grains were then cleaned of impurities and damaged grains. Sampling was performed at 30-day intervals and in accordance with the ISO method [34]. A sample of approximately 4 kg of freshly harvested buckwheat seeds was brought to the laboratory. A detailed description of the sample storage procedure has been previously reported [35]. The experimental conditions and workflow were summarized and presented in graphical form (Figure 1).
For each parameter, analyses were conducted on three separately prepared samples from sub-samples, each in triplicate.
2.2. Chemicals, Reagents, and Standards
All chemicals, reagents, and standard solutions used in the experiments were of analytical grade, and the enzyme solutions had the recommended enzyme activity. The Folin–Ciocalteu reagent was supplied by Reagecon Diagnostics Ltd. (Shannon, Ireland), and 2,4,6-tripyridyl-s-triazine (TPTZ, ≥99%) was obtained from Fluka (Honeywell, Charlotte, NC, USA). Additionally, 1,1-diphenyl-2-picrylhydrazyl radical (DPPH, >97.0%) and 6-hydroxy-2,5,7,8-tetramethylchroman-2-carboxylic acid (Trolox, 97%) were purchased from Sigma-Aldrich Chemie GmbH (Merck KGaA, Darmstadt, Germany). Analytical standards of phenolic acids (gallic acid, dihydrocaffeic acid, phloretic acid, trans-cinnamic acid, chlorogenic acid, caffeic acid, p-coumaric acid, ferulic acid, hesperetic acid, ≥98.0%, w/w) and bioflavonoids (catechin, epicatechin, naringin, daidzein, naringenin, quercetin, ≥98.0%, w/w) were also provided by Sigma-Aldrich Chemie GmbH (Merck KGaA, Darmstadt, Germany). For chromatographic analysis, gradient grade HPLC methanol CHROMASOLV™ (≥99.9%) and HPLC grade water CHROMASOLV Plus were obtained from Riedel-de Haën (Honeywell, Charlotte, NC, USA).
2.3. Determination of Chemical and Nutritional Properties
The physicochemical, chemical, and nutritional properties were analyzed through the examination of proximate composition (moisture, total protein, total lipid, total ash, and total carbohydrate content), as well as starch, β-glucan (dietary fiber), energy value, pH value, and alcoholic acidity.
Moisture content was determined using the gravimetric method according to ISO 712 [36] with a Digitheat-TFT electric oven (J.P. Selecta, Barcelona, Spain). Before adding the samples, the metal dish with a lid was dried at a temperature between 130 °C and 133 °C. The closed dish was then cooled in a desiccator to room temperature, and its weight was recorded. Five grams of undried flour was weighed into a metal dish, and the mass of the sealed dish containing the sample was measured. An open metal dish with flour and a lid was dried for 90 min at a temperature between 130 °C and 133 °C. The closed dish with the flour sample was cooled in a desiccator for 45 min, and then the mass of the closed metal dish with flour was measured. The drying and cooling process was repeated until a constant mass was obtained. Moisture (in %) was calculated as the mass difference of the dish with lid plus undried flour and the dish with lid plus oven-dried flour divided by the mass difference of the dish with lid plus undried flour and the empty dish with lid.
Total lipid content was determined using the extraction-gravimetric method according to NMKL 160 [37] with a Soxhlet-type extraction apparatus, DK-2000-IIIL water bath (ChemLand, Stargard, Poland), and a Digitheat-TFT electric oven (J.P. Selecta, Barcelona, Spain). A 2 g sample of flour was placed in an Erlenmeyer flask, then boiled and hydrolyzed for 1 h with 50 mL of 4 mol/L hydrochloric acid to liberate bound fats and transform fatty acids salts into free acids. The mixture was cooled to room temperature, filtered through filter paper, and washed three times with warm distilled water. The residue on the filter paper was dried in an electric oven for 90 min at 102 ± 2 °C. The dried filter paper with residue was placed into an extraction thimble and put into the Soxhlet-type extraction apparatus, where the extraction was carried out for about 6 h. Meanwhile, a receiving round-bottom flask was filled two-thirds with extraction solvent (petroleum ether). Approximately 10 mL of petroleum ether was condensed per minute during continuous extraction. After extraction, the solvent was distilled off, and the receiving round-bottom flask with the fat residue was dried in the oven for 2 h at 102 ± 2 °C, then cooled in a desiccator for 30 min before measuring its mass. The drying and cooling procedure was repeated until a constant mass was obtained. Total lipids (in %DM) were calculated as the mass difference between the receiving round-bottom flask with fat residue after extraction and the empty receiving round-bottom flask before extraction, then recalculated to 100 g of the sample’s dry matter.
Total ash content was determined using the gravimetric method according to ISO 2171 [38] with a ME 520 electrically heated muffle furnace (Prederi Vittorio & Figli s.n.c., Milan, Italy). Before adding the sample, the cleaned platinum ashing dish was placed in the furnace and heated to 900 ± 25 °C for 5 min, then cooled in a desiccator to room temperature, and the mass of the empty ashing dish was measured. About 2 g of flour was placed in the platinum ashing dish and transferred to the muffle furnace heated to 900 ± 25 °C. The sample was first ignited, and then incineration was continued for at least 1 h. After incineration, the platinum ashing dish with the residue was cooled for 20 min in a desiccator to room temperature, and the mass of the ashing dish containing the ignited residue was measured. The drying and cooling procedure was repeated until a constant mass was obtained. Total ash content (in %DM) was calculated as the difference in mass before and after incineration, then recalculated to 100 g of the sample’s dry matter.
ISO 20483 [39] was used for the Kjeldahl volumetric determination of total nitrogen content, employing the K-424 digestion system unit (Buchi, Flawil, Switzerland), K-350 distillation system unit (Buchi, Flawil, Switzerland), and a Titrette® digital burette (Brand GmbH & Co KG, Wertheim, Germany). A 2.5 g sample of flour was placed in a digestion flask (Kjeldahl tube), and one Kjeldahl catalyst tablet (a mixture of 10 g of potassium sulfate, 0.3 g of copper sulfate pentahydrate, and 0.3 g of titanium dioxide) was added, followed by 20 mL of sulfuric acid. The flask was swirled until the contents were completely wetted. The digestion flask with the mixture was then placed in the digestion system unit, which was heated to 420 ± 10 °C, and the digestion process was allowed to continue for at least 2 h until the digestion mixture became clear and foaming and vaporization ceased. After digestion, the flask with the content was cooled, and 50 mL of water was added and stirred slowly. The flask and its contents were then transferred to the distillation system unit. Simultaneously, in the part below the condenser of the distillation system unit, a 250 mL Erlenmeyer flask filled with 50 mL of boric acid solution (40 g/L) was placed so that the end of the condenser hose was immersed in the solution. Then, 10 drops of indicator (0.2 g bromocresol green and 0.2 g methyl red dissolved in 100 mL ethanol, mixed in a ratio of 5:1) were added. Steam distillation was initiated after introducing 50 mL of sodium hydroxide (32%) into the distillation column. The distillation rate was maintained at approximately 25 mL/min (distillation time 5 min), and the process was stopped when 125 mL of distillate was obtained. The distillate was titrated with a standard sulfuric acid volumetric solution (0.05 mol/L) until a color change at the endpoint was observed. The same procedure was performed with a blank sample. The consumption of sulfuric acid solution in the flour sample and the blank sample was recorded. Total protein (in %DM) was calculated from the nitrogen content multiplied by 6.25 and then recalculated to 100 g of the sample’s dry matter.
Total carbohydrate content was determined by difference, subtracting the sum of the mass percentage of constituents from the mass percentage of total solids according to AOAC 986.25 [40] using the following Equation (1):
(1)
Total carbohydrates (in %DM) were expressed as a percentage of mass and then recalculated to 100 g of the sample’s dry matter.
Energy value was calculated according to the Codex Alimentarius Commission [41], using conversion factors (cf 17 kJ/g for protein and carbohydrates, cf 37 kJ/g for lipids) in the standard Equation (2):
(2)
The energy value was expressed in kJ/100 g of sample and reported to the nearest whole number.
Starch content was determined using the Ewers polarimetric method according to ISO 10520 [42] with a Polartronic M TOUCH high-performance circle polarimeter equipped with 200 mm measuring tubes (Schmidt + Haensch GmbH & Co., Berlin, Germany). The method involved two steps of polarimetric determination. For the total optical rotation, approximately 2.5 g of flour was weighed into a 100 mL volumetric flask. To this, 25 mL of dilute hydrochloric acid (0.309 mol/L) was added and shaken vigorously. An additional 25 mL of the same dilute hydrochloric acid was then added. The flask with its content was immersed in a boiling water bath for 15 min ± 5 s, then removed, filled with 30 mL of cold water, and cooled immediately to a temperature of 20 ± 2 °C. Next, 5 mL of Carrez solution I (10.6 g of potassium hexacyanoferrate(ll) trihydrate dissolved in 100 mL water) was added and shaken for 1 min, followed by 5 mL of Carrez solution II (21.9 g of zinc acetate dihydrate and 3 g of glacial acetic acid dissolved and diluted to 100 mL with water) also shaken for 1 min. The solution was clarified, diluted to the mark with water, mixed, and filtered. The optical rotation of the filtrate was measured at 20 °C using a polarimetric cell of 200 mm optical path length at a wavelength of 589.3 nm. To determine the optical rotation of substances soluble in 40% ethanol, about 5 g of flour was weighed into a 100 mL volumetric flask. To this, 80 mL of ethanol solution (40%, v/v) was added, and the content was left at room temperature for 1 h, with vigorous shaking, repeated six times during this period. The solution was diluted to the mark with the same ethanol solution, mixed, and filtered. Then, 50 mL of the filtrate was pipetted into a 100 mL volumetric flask, 2.1 mL of dilute hydrochloric acid (7.7 mol/L) was added, and the flask was fitted with a reflux condenser and immersed in a boiling water bath for 15 min ± 5 s. After cooling to 20 ± 2 °C, the previous clarification procedure was repeated, including the addition of 5 mL of Carrez solutions I and II. The solution was then diluted to the mark with water, mixed, and filtered. The optical rotation was measured as described previously. Starch (in %DM) was calculated as the difference between these two measurements, multiplied by a factor, and then recalculated to 100 g of the sample’s dry matter.
β-glucan content was determined using the enzyme-spectrophotometric method according to AOAC 995.16 [43] utilizing a DK-2000-IIIL water bath (ChemLand, Stargard, Poland), an EBA 280 centrifuge (Hettich, Tuttlingen, Germany), and a UV-VIS 2100 spectrophotometer (Shimadzu Corporation, Kyoto, Japan). The necessary reagents and solutions were prepared according to the procedures described in the standard. Approximately 0.1 g of homogenized flour sample was weighed into a centrifuge tube (17 mL capacity). Next, 0.2 mL of ethanol solution (50%, v/v) and 4.0 mL of sodium phosphate buffer (20 mmol/L, pH 6.5) were added. The centrifuge tube was incubated in a boiling water bath for 1 min, vortex mixed, and then returned to incubate for an additional 2 min. Afterward, the incubation was continued at 50 °C for 5 min, followed by the addition of 0.2 mL of lichenase solution. The centrifuge tube was capped and incubated for another 1 h at 50 °C. Next, the procedure required the addition of 5.0 mL of sodium acetate buffer (200 mmol/L, pH 4.0) to the centrifuge tube and the equilibration at room temperature for 5 min, followed by centrifugation at 2800 rpm for 10 min. Into each of the three centrifuge tubes (12 mL capacity), 0.1 mL of supernatant was pipetted. To two of these tubes, 0.1 mL of β-glucosidase solution was added (reaction solution), while to the third tube, 0.1 mL of sodium acetate buffer (50 mmol/L, pH 4.0) (reaction blank) was added instead. All tubes were then incubated at 50 °C for 10 min. Subsequently, 3 mL of glucose oxidase-peroxidase-buffer mixture was added to each tube, and the incubation in the water bath continued at 50 °C for 20 min. For each set of determinations, a reagent blank (0.2 mL of 50 mmol/L sodium acetate buffer and 3.0 mL glucose oxidase-peroxidase-buffer mixture) and a D-glucose standard working solution for quality control (0.1 mL of 50 mmol/L sodium acetate buffer, 0.1 mL D-glucose standard of 1 mg/mL and 3.0 mL glucose oxidase-peroxidase-buffer mixture) were included. Absorbance was measured at λ = 510 nm against the reagent blank within 1 h. β-glucan (in %DM) was calculated according to the equation specified in the standard and then recalculated to 100 g of sample dry matter
The pH was determined following the direct potentiometric method according to AOAC 943.02 [44] using a HI2020 Edge pH-meter (HANNA Instruments Inc., Woonsocket, RI, USA) with a glass electrode. Standard buffer solutions of pH 4 and 9 were used for calibration. Approximately 10 g of flour was placed in an Erlenmeyer flask, and 100 mL of boiled water was added. The suspension was digested for 30 min and then allowed to settle for 10 min. Afterward, the supernatant was decanted into a 250 mL beaker, and the pH was measured.
Alcoholic acidity was determined following the volumetric acid-alkaline titration method according to IS 12711 [45] using a Titrette® digital burette (Brand GmbH & Co KG, Wertheim, Germany). Approximately 5 g of flour was transferred into a stoppered conical flask, and 50 mL of neutral ethyl alcohol (90%, v/v) was added. The suspension was shaken and allowed to settle for 24 h. The alcoholic extract was then filtered, and the filtrate was titrated with standard volumetric sodium hydroxide solution (0.05 mol/L) using phenolphthalein as an indicator until a permanent pink color appeared. Alcoholic acidity (in %DM) was expressed as sulfuric acid and calculated according to Equation (3), then recalculated to 100 g of the sample’s dry matter.
(3)
where A is the volume of consumed standard volumetric sodium hydroxide solution, N is the normality of standard volumetric sodium hydroxide solution, and M is the weight of the sample.2.4. Extraction Process
To determine the antioxidant activity and the profiles of phenolic acids and bioflavonoids, an extraction process was carried out. The extraction procedure, with some modifications, used a suitable extraction solvent system composed of methanol and 10% hydrochloric acid in an 85:15 (v/v) ratio, as per previously established methods [46,47]. The powdered samples (0.5 g) were extracted with 10 mL of solvent in 15 mL conical-bottomed plastic centrifuge tubes. The tubes were first allowed to settle for 15 min at room temperature, then extracted in a digital ultrasonic cleaner-bath model DU-45 (Argolab, Carpi, Italy) at an ultrasound frequency of 40 kHz for 30 min. After extraction, the samples were centrifuged at 6000 rpm for 15 min using an EBA 280 centrifuge (Hettich, Tuttlingen, Germany). The residues were extracted again with 10 mL of solvent following the same steps. The supernatants were merged and evaporated to dryness at 45 °C under vacuum. Finally, the residues were diluted to 10 mL with solvent and filtered through 0.45 μm pore size PTFE membrane syringe filters to obtain the ready-to-use acidified methanol extracts, which were then stored for 1–2 weeks at 4 °C before analysis.
2.5. Determination of Antioxidant Activity
Antioxidant activity was analyzed by conducting total phenol content (TPC), ferric reducing/antioxidant power (FRAP), and radical-scavenging activity (DPPH) assays.
The total phenol content (TPC) was determined spectrophotometrically using the Folin–Ciocalteu method according to the reported procedure by Dang et al., with some modifications [46]. Prior to use, Folin–Ciocalteu reagent was diluted with deionized water in a ratio of 1:1 (v/v). For each sample, 50 µL of the extract was placed into a glass test tube and mixed with 3 mL of deionized water, 250 µL of the diluted Folin–Ciocalteu reagent, and 750 µL of 20% Na2CO3 solution. The mixture was allowed to react for 8 min at room temperature. Subsequently, 950 µL of deionized water was added to each glass test tube to bring the final volume to 5 mL. The obtained test sample solutions were kept away from light and incubated at room temperature for 2 h. A stock solution of gallic acid (1000 mg/L) was prepared in methanol, and working standard solutions (0, 25, 50, 75, 100, 150, and 200 mg/L) were obtained by further dilution in methanol. Working standard solutions and blank samples (methanol instead of extract) were subjected to the same procedure steps as the test samples. The absorbances of working standard and test sample solutions were measured against the blank sample at a wavelength of λ = 765 nm using a UV-VIS 2100 spectrophotometer (Shimadzu Corporation, Kyoto, Japan). A calibration curve was constructed, and the equation of the regression line was performed. The total phenol content (in mg GAE/g DM) was calculated using gallic acid (GA) as the standard and then recalculated to 1 g of the sample’s dry matter.
The ferric reducing/antioxidant power (FRAP) was determined spectrophotometrically using the previously described method by Benzie and Strain, with some modifications [48]. The FRAP reagent was prepared by mixing 2.5 mL of 10 mmol/L TPTZ in 40 mmol/L HCl solution, 2.5 mL of 20 mmol/L FeCl3 × 6H2O solution, and 25 mL of 300 mmol/L pH 3.6 sodium acetate buffer solution, in a simplified ratio of 1:1:10 (v/v/v). The sample extracts were previously diluted 10-fold with methanol, and 150 µL of each diluted extract was taken, placed in glass test tubes, then mixed with 4.5 mL of FRAP reagent, and thoroughly shaken. The obtained test sample solutions were incubated at 37 °C for 30 min to react. A stock solution of FeSO4 × 7H2O (1 mmol/L) was prepared in deionized water, and working standard solutions (0, 25, 50, 100, 250, 500, and 750 μmol/L) were prepared by further dilution in deionized water. Working standard solutions and blank samples (methanol instead of diluted extract) were subjected to the same procedure steps (except for 10-fold dilution) as the test samples. Based on the reduction of Fe3+ ion to Fe2+ ion and the formation of the blue complex Fe2+-TPTZ, the absorbances of working standard and test sample solutions were measured against the blank sample at the wavelength of λ = 593 nm using a UV-VIS 2100 spectrophotometer (Shimadzu Corporation, Kyoto, Japan). A calibration curve was constructed, and the equation of the regression line was performed. Ferric reducing/antioxidant power (in μmol Fe2+/g DM) was calculated using FeSO4 × 7H2O as the standard and then recalculated to 1 g of the sample’s dry matter.
Radical-scavenging activity (DPPH) was determined spectrophotometrically using the previously described method by Bakar et al., with some modifications [49]. The DPPH reagent was prepared in the form of 0.1 mmol/L DPPH–methanol solution by dissolving 3.9 mg of DPPH in methanol up to a final volume of 100 mL in a volumetric flask. The volumetric flask was then wrapped in aluminum foil and placed in a digital ultrasonic cleaner-bath model DU-45 (Argolab, Carpi, Italy) for 30 min. The sample extracts were previously diluted 10-fold with methanol; 200 µL of each extract was then transferred to glass test tubes, mixed with 3.8 mL of DPPH–methanol solution, and thoroughly shaken. The obtained test sample solutions were kept in the dark and incubated at room temperature for 30 min to develop color. A stock solution of Trolox (1 mmol/L) was prepared in methanol, and working standard solutions (0, 20, 50, 100, 250, 500, and 750 μmol/L) were prepared by further dilution in methanol. Working standard solutions were subjected to the same procedure steps (except for 10-fold dilution) as the test samples. Based on the reduction of the free radical DPPH, the absorbances of working standard and test sample solutions were measured against a blank sample (methanol) at the wavelength of λ = 517 nm using a UV-VIS 2100 spectrophotometer (Shimadzu Corporation, Kyoto, Japan). A calibration curve was constructed, and the equation of the regression line was performed. Radical-scavenging activity (in μmol TE/g DM) was calculated using Trolox (Trolox equivalents, TE) as the standard and then expressed per gram of the sample’s dry matter.
2.6. Determination of Phenolic Acids and Bioflavonoids Profiles—HPLC Analysis
The profiles of phenolic acids and bioflavonoids were determined by reverse-phase high-performance liquid chromatography (RP-HPLC) according to the reported method, with some modifications [46]. The compounds were separated, detected, identified and quantified using a Nexera HPLC System (Shimadzu Corporation, Kyoto, Japan) consisting of a quaternary pump (Nexera XR LC-20AD XR; Shimadzu Corporation, Kyoto, Japan), degassing unit (DGU-20A SR; Shimadzu Corporation, Kyoto, Japan), auto-sampler (Nexera XR SIL-20AC XR; Shimadzu Corporation, Kyoto, Japan), column oven (Prominence CTO-20AC; Shimadzu Corporation, Kyoto, Japan), and photodiode array detector (Prominence SPD-M20A; Shimadzu Corporation, Kyoto, Japan). The system was equipped with a reversed-phase column Zorbax SB C18 (250 × 4.6 mm, I.D. 5 μm; Agilent Technologies Inc., Santa Clara, CA, USA). The binary gradient elution system used two solvents as the mobile phase: (A) water containing 0.1% formic acid and (B) methanol. Gradient programming of the solvent system (in v/v) was as follows: 0 min 5% B, 25 min 30% B, 35 min 40% B, 40 min 48% B, 50 min 70% B, 55 min 100% B, 65 min 5% B, re-equilibration time 10 min, with a flow rate of 1 mL/min. Therefore, the total run time for analysis was 75 min. A total of 10 μL of acidified methanol extract of the sample and the standard solution was injected from vials (1.5 mL capacity), and the column was thermostatically controlled at 25 °C. Dual wavelengths of λ = 280 nm and 325 nm were used to detect the eluted phytochemicals. The obtained chromatograms of the standards and samples were overlayed, and the unknown phytometabolites from the samples were identified by comparing the retention time of the peaks. Quantification was performed using a calibration curve made for each compound based on the peak areas of a standard mixture with known concentrations (0, 25, 50, 100, 150, and 200 mg/L). Stock solutions were prepared in a methanol-water mixture (75:25, v/v) at a concentration of 1000 mg/L. The calibration standard mix solutions were prepared by combining the stock solutions and diluting them with the methanol-water mixture to appropriate concentrations. Method quality parameters (linearity of calibration curves (R2), repeatability (RSD), the limit of repeatability (r = 2.8∙Sr), bias (%), the limit of detection (LOD = 3∙Srblank), the limit of quantification (LOQ = 10∙Srblank), measurement uncertainty (Uexpanded = 2∙combined MU), and instrument performance parameters (flow, carry-over, linearity, etc.) were ensured as shown in Table 1 and Table 2. HPLC analysis at 280 nm was used to identify and quantify the peaks of gallic acid, dihydrocaffeic acid, phloretic acid, trans-cinnamic acid, catechin, epicatechin, naringin, daidzein, naringenin, while at 325 nm for chlorogenic acid, caffeic acid, p-coumaric acid, ferulic acid, hesperetic acid, and quercetin. The content of phenolic acids and bioflavonoids (in μg/g DM) was calculated using a standard mixture and then recalculated to 1 g of the sample’s dry matter.
2.7. Statistical Analysis
To interpret obtained results of the Novosadska variety buckwheat experiments, the data were presented as mean ± standard deviation (M ± SD) and statistically processed by one-factor analysis of variance (one-way ANOVA, p < 0.05) to evaluate the effect of storage time, and Tukey’s HSD (Honestly Significant Difference, p < 0.05) post hoc test to determine the differences between means, using Statistica 12.5 software (StatSoft, Inc., Tulsa, OK, USA).
3. Results
3.1. Physicochemical, Chemical, and Nutritional Properties
The results of proximate composition, nutritional properties, pH values, and alcoholic acidity of the Novosadska variety buckwheat (flour) used in this research during the 9-month storage are presented in Table 3. During the storage period, the trends of a significant (p < 0.05) successive decrease in moisture, total protein, and total lipids content were observed, in contrast to total carbohydrate content, where the trend of a significant (p < 0.05) successive increase was recorded.
During the 9-month storage period, no significant changes (p > 0.05) were noted in total ash content. There was a statistically significant difference (p < 0.05) in moisture content between all treatments. However, for total protein, total lipid, and total carbohydrate content, a statistically significant difference (p < 0.05) was observed at 6 months, but later without significant (p > 0.05) changes between treatments. Compared to freshly harvested seeds, which had the highest moisture (10.21%), total protein (13.46% DM), and total lipid (3.44% DM) content, during storage, the lowest moisture (6.02%) and total protein (11.57% DM) content was recorded at 9 months, and total lipid content (2.64% DM) at 6 months, resulting in a decrease of 41%, 14% and 23%, respectively. On the other hand, the highest total carbohydrate content (83.48% DM) was observed at 9 months, and the lowest amount (81.13% DM) in freshly harvested seeds, which corresponded to an increase of 3%.
Furthermore, progressive changes with increasing storage time were observed for all examined nutritional parameters of buckwheat flour. A significant decrease (p < 0.05) in starch content was observed during storage, while significant increases (p < 0.05) in β-glucan content and energy value were recorded. There was a statistically significant difference (p < 0.05) in β-glucan content between all treatments. However, for starch content and energy value, statistically significant differences (p < 0.05) between treatments were observed at 3 months, i.e., at 6 months, respectively, and continued thereafter. Compared to freshly harvested seeds, which had the highest starch content (62.55% DM), during storage, the lowest amount (47.67% DM) was recorded at 6 months, which corresponded to a decrease of 24%. On the other hand, the highest β-glucan content (0.120% DM) and energy value (1623 kJ/100 g) were observed at 9 months, while the lowest β-glucan content (0.030% DM) and energy value (1558 kJ/100 g) were noted at 3 months, i.e., in freshly harvested seeds, increasing by 300% (for freshly harvested seeds it was 140%) and 4%, respectively.
During the storage period, a significant (p < 0.05) successive decrease in pH value and alcoholic acidity was observed. Statistically significant (p < 0.05) changes in pH value and alcoholic acidity were noted between treatments at 3 months and beyond, but without significant (p > 0.05) changes in pH value among treatments at 6 months and later. Compared to freshly harvested seed, which had the highest pH value (6.74), and alcoholic acidity (0.86% DM), during storage, the lowest pH value (6.19) and alcoholic acidity (0.53% DM) were recorded at 9 months, resulting in a decrease of 0.55 pH units, and 0.33% DM, respectively.
3.2. Potential of Antioxidant Activity and Capacity
The results of the antioxidant activity of the Novosadska variety buckwheat (flour) used in this research during the 9-month storage presented via total phenol content (TPC), ferric reducing/antioxidant power (FRAP), and radical-scavenging activity (DPPH) assays are shown in Table 4.
During the storage period, significant successive decreases (p < 0.05) in total phenol content, FRAP, and DPPH were observed. There was a statistically significant difference (p < 0.05) in FRAP value between all treatments. However, for total phenol content and DPPH, a statistically significant difference (p < 0.05) among treatments was observed at 6 months and beyond. Compared to freshly harvested seeds, which had the highest total phenol content (7.28 mg GAE/g DM) and FRAP (63.45 μmol Fe2+/g DM), during storage, the lowest TPC (5.57 mg GAE/g DM) and FRAP (22.20 μmol Fe2+/g DM) values were recorded at 9 months, resulting in a decrease of 1.3 and 2.9 times, respectively.
On the other hand, the highest DPPH (19.66 μmol TE/g DM) was observed at 3 months, and the lowest value (8.12 μmol TE/g DM) at 9 months, which corresponded to a decrease of 2.4 times.
3.3. Phenolic Acids and Bioflavonoids Profiles
In the present research, 15 phytochemical compounds were studied to be identified and quantified in Novosadska variety buckwheat (flour) during 9 months of storage. Nine of them were phenolic acids: gallic acid, dihydrocaffeic acid, phloretic acid, trans-cinnamic acid, chlorogenic acid, caffeic acid, p-coumaric acid, ferulic acid, hesperetic acid, and six belonged to bioflavonoids: catechin, epicatechin, naringin, daidzein, naringenin, quercetin. The obtained results are presented in Table 5.
Eight phenolic acids ranged as follows: gallic acid (91.0–94.1 μg/g DM), dihydrocaffeic acid (11.5–19.1 μg/g DM), chlorogenic acid (7.1–14.4 μg/g DM), caffeic acid (37.6–39.2 μg/g DM), phloretic acid (25.8–46.5 μg/g DM), p-coumaric acid (95.9–102.4 μg/g DM), ferulic acid (56.5–57.4 μg/g DM), and trans-cinnamic acid (87.9–90.0 μg/g DM). Hesperetic acid, which belongs to the class of phenolic acids, was not detected in any section (0—freshly harvested seeds, 3, 6, and 9 months). Additionally, six bioflavonoids ranged as follows: catechin (49.1–52.2 μg/g DM), epicatechin (59.8–95.7 μg/g DM), daidzein (56.1–69.3 μg/g DM), quercetin (9.8–47.1 μg/g DM), naringin (nd—54.9 μg/g DM), and naringenin (37.8–38.7 μg/g DM).
During the storage period, a significant (p < 0.05) successive decrease in chlorogenic acid content was observed among phenolic acids. Although the changes in the content of dihydrocaffeic and phloretic acid were also significant (p < 0.05), irregular and mutually opposite patterns were found. During the storage period of 9 months, no significant changes (p > 0.05) were recorded in the content of the following phenolic acids: gallic acid, caffeic acid, p-coumaric acid, ferulic acid, and trans-cinnamic acid. Regarding bioflavonoids during storage, a significant (p < 0.05) successive increase in daidzein, quercetin, and naringin content was observed, while a significant (p < 0.05) successive decrease was recorded in epicatechin content. No significant changes (p > 0.05) were noted in the content of catechin or naringenin during the observed 9-month storage period.
Statistically significant (p < 0.05) changes in chlorogenic acid content were observed between treatments at 3 months and beyond; however, no significant (p > 0.05) changes were found among treatments at 6 and 9 months. The changes in the content of dihydrocaffeic and phloretic acid between treatments were significant (p < 0.05) only at 3 months but without significant (p > 0.05) variations among other treatments, as can also be seen in Figure 2a.
Although no statistically significant differences (p > 0.05) in the content of gallic acid, caffeic acid, ferulic acid, p-coumaric acid, and trans-cinnamic acid were observed between treatments (Table 5), there was an obvious tendency that reflected in a small increasing increment of gallic acid content, and a small decreasing increment of p-coumaric and trans-cinnamic acid content, or these changes in the content between treatments were without a definite pattern as in caffeic and ferulic acid. Regarding bioflavonoids, a statistically significant (p < 0.05) alteration in epicatechin content among treatments was observed during storage at 3 months and beyond, as shown in Figure 2b. However, for daidzein, quercetin, and naringin contents, statistically significant differences (p < 0.05) between treatments were observed at 9 months, 6 months, and beyond, as well as at 3 months and beyond, respectively, but for naringin content, there were no significant (p > 0.05) changes among treatments at 6 and 9 months. On the other hand, it was observed across all treatments that the individual contents of catechin, as well as naringenin, were at a very similar quantitative level, with invariability (p > 0.05) and constancy in content.
Regarding phenolic acids, compared to freshly harvested seeds, which contained dihydrocaffeic acid and phloretic acid of 18.7 μg/g DM and 25.8 μg/g DM during storage, the lowest dihydrocaffeic acid content (11.5 μg/g DM) and the highest phloretic acid content (46.5 μg/g DM) were recorded at 3 months, resulting in a decrease of 1.6 times, and an increase of 1.8 times, respectively (as shown in Figure 2a). The highest content of chlorogenic acid (14.4 μg/g DM) was recorded at the beginning in the freshly harvested seeds, but it decreased over time to the lowest value (7.1 μg/g DM) at 9 months, which corresponded to a 2-fold decrease. In the case of bioflavonoids, the highest content of epicatechin (95.7 μg/g DM) was in the freshly harvested seeds, which was 1.6 times higher than the lowest content (59.8 μg/g DM) at 9 months. On the other hand, after nine months of storage, the amount of daidzein (69.3 μg/g DM) and quercetin (47.1 μg/g DM) was 1.2 and 4.8 times higher than in the freshly harvested grains (57.0 μg/g DM, 9.8 μg/g DM). Similarly, at the beginning of the storage period, naringin was not detected in the freshly harvested seeds, but over time, its content increased significantly to 54.9 μg/g DM (6 months) and 51.9 μg/g DM (9 months), as shown in Figure 2b.
The chromatographically determined profiles of phenolic acids and bioflavonoids, as an example of separation and detection at 3 months of storage, are introduced in Figure 3.
4. Discussion
The results of these studies indicate that Novosadska variety buckwheat seeds are a good source of energy and essential building and functional components for the daily diet. At the same time, great efforts are being made to promote fewer consumed cereals and pseudocereals, such as millet, sorghum, buckwheat, quinoa, etc. This involves combining industrial processing techniques to increase the content of macro- and micronutrients, the levels of bioactive polyphenols, bioflavonoids, and other compounds in the seeds of these crops, therefore enabling the development of functional foods with high nutritional quality [32,33].
4.1. Physicochemical, Chemical, and Nutritional Properties
Chemical changes caused by the natural aging process, and especially under the conditions to which the samples in this study were exposed, led to changes that did not drastically affect the quality. Similar to our initial findings on the proximate composition findings of Novosadska variety buckwheat whole-seed flour (Table 3), Dapčević et al. [50] reported moisture, total protein, total lipid, and total ash mass fractions of raw whole-seed buckwheat flour as 9.76, 13.40, 3.08 and 1.97% DM, respectively. In another study, the total carbohydrate mass fraction was slightly lower and reported to be 72.4% DM [51]. The decrease in moisture content during storage is attributed to thermal action at 40 ± 2 °C and the gradual loss of water due to the seed’s porous capillary structure [52]. The deterioration of cultivation conditions and standards and uncontrolled post-harvest treatment mentioned in the previous research [52] could be the main reasons for discrepancies in moisture content compared to the results of our research. Our results in the overall decreasing trends of total protein and lipid content remain within the ranges for buckwheat reported by Nalinkumar and Singh for proteins 11–14% DM and lipids 1.5–3.7% DM [53]. However, the slight decrease is likely due to biochemical processes such as protein and lipid hydrolysis, leading to the release of amino and fatty acids and lipid oxidation [52]. It is important to mention that the biological value of buckwheat protein is higher compared to wheat, oats, rye, and barley, while buckwheat lipids mainly consist of 95% of saturated palmitic and unsaturated oleic and linoleic fatty acids [53]. Although the increase in total carbohydrate content over time is statistically significant, it could be attributed to the calculation methodology of the determination itself and the possible inhomogeneity of the sample. Nalinkumar and Singh [53] reported a value of total ash content (common buckwheat 2.1%, and Tartary buckwheat 1.8%) similar to the value we obtained at the beginning of the research. During long-term storage, there is little effect on the mineral content [54], which is consistent with our results.
At the beginning of this research, the starch content in raw whole-seed buckwheat flour was slightly lower than the 67.40% DM reported by Dapčević et al. [50]. Over time, the changes in starch content led to a partial overlap with the starch content of buckwheat, ranging from 59% to 70%, as reported by Nalinkumar and Singh [53]. The decrease in starch content may be due to enzymatic and chemical (acid) hydrolysis, which efficiently converts starch into monosaccharides. This process is facilitated by pH (5.0–7.0) and thermal conditions (45 °C) [52,55], which are consistent with the storage conditions in our experiment, where the pH ranged from 6.19 to 6.74, and the temperature was 40 ± 2 °C. In addition, the differences in starch content reported in the literature vary due to the different extraction methods applied and cultivar varieties [53,56].
β-glucan content and structure in crops have implications for enhanced nutritional value. The findings reported by Singla et al. [57] somewhat support our findings, as they stated that buckwheat is poor in β-glucan content compared to cereals (oat, barley). However, our results were inconsistent with the findings of Hozová et al. [58], who found a multiple times higher amount of β-glucan in buckwheat seed cv. Spacinska (A and B) (more than 20%). A possible explanation for this huge difference in the β-glucan content is the influence of various factors, such as the species and cultivar, the quality of the agricultural soil where the plant was grown, and climatic conditions during the year of cultivation [58]. On the other hand, during storage, the growth trend could be explained by the gradual and easier release of this polysaccharide from the cell wall and the aleurone layer of the endosperm of the damaged seed, making it more readily detected [46].
In this study, the energy values measured both at the beginning and after 9 months were very similar to the results previously reported by Silav-Tuzlu and Tacer-Caba [51], who found an energy value of 320.0 kcal/100 g (1338 kJ/100 g). The minor increase in energy values during storage could be attributed to variations in component quantities and measurement uncertainty inherent in the determination process.
Changes in acidity and pH can lead to deteriorated quality, especially in terms of nutritional and sensory properties, and these changes indicate the existence and development of proteolytic and lipolytic processes, which make the seeds less suitable for human consumption and usage alone or as an ingredient in functional food. Controlled conditions with lower temperature and humidity can extend the shelf life of the seed and long-storage while preserving its best quality. Our initial pH value and alcoholic acidity were similar to those reported by Jara et al. [59], who examined fine whole-seed buckwheat flour and found a pH of 6.36 and an average titratable acidity value of 6.2% (recalculated to alcoholic acidity of 0.62%), and Mousavi et al. [60], who determined a pH value of buckwheat flour as 6.25. In our research, both pH and acidity values decreased over storage time, which aligns with the declining pH trend found by Silav-Tuzlu and Tacer-Caba [51] of 5.57 to 5.27 in biscuits with buckwheat over 45 days, and the reduced acidity found by Mgaya-Kilima et al. [61] due to acidic hydrolysis of polysaccharides where acid converts non-reducing sugars into reducing sugars (mono- and disaccharides). These decreases in pH value may result from the activity of certain microorganisms, the release of fatty acids due to lipid hydrolysis, or from sugar conversion to small amounts of alcohols and acids [52,62].
4.2. Potential of Antioxidant Activity and Capacity
Buckwheat has a higher antioxidative activity compared to cereal grains, and frequent consumption of buckwheat products could protect the human body from oxidative damage caused by free radicals [22].
The amount of total phenolics (Table 4), attributed to the presence of polyphenols, significantly contributes to the overall antioxidant activity, which is reflected, among other things, in the inhibition of lipid peroxidation [63]. However, not all polyphenols have functional and nutritional benefits. For example, tannins can bind to proteins, carbohydrates, some vitamins, and minerals, reducing their intestinal absorption, which makes them a group of polyphenols with antinutrient properties ranging from 15 to 41 mg/100 g [2]. Our initial total phenolic content was lower than that reported by Djordjevic et al. [64], who found a TPC of 50.7 mg GAE/g dry extract in buckwheat (Fagopyrum esculentum), which had the highest content among tested plant materials. It can be assumed that this apparently large difference in TPC values between the studies arose from the way the results were expressed, as our study measured per gram of dry matter in flour, while theirs measured per gram of dry extract. In addition, differences could originate from different commercially available cultivars and factors such as natural chemical composition, harvest maturity, soil state, conditions of post-harvest storage, as well as the loss of antioxidants or the formation of compounds with pro-oxidant action. Examining the seeds of six buckwheat varieties, Zhu et al. [65] found TPC values ranging from 5.81 to 14.40 mg GAE/g DM, with 7.31 mg GAE/g DM in black common buckwheat, identical to our value for freshly harvested seeds at month zero. Our observed 23% decrease in TPC during 9 months of storage is consistent with the findings of Starowicz and Zieliński [66], who found a 24% decrease in the content of total phenolic compounds in rye-buckwheat cakes over 18 months, but for a period twice as long as ours and at a temperature almost twice as low (23 °C). The decrease in TPC during storage could be explained by the participation in strong antioxidative activity inhibiting lipid oxidation but also partly by the compound degradation entering various reactions forming Maillard reaction compounds or favorable polyphenol-sugar adducts, which subsequently rearrange to pigments [66].
Furthermore, antioxidant ability was assessed using ferric ion-reducing antioxidant power (FRAP). Flour made from freshly harvested Novosadska variety buckwheat seeds (0 months) exhibited 1.3 times higher activity of electron-donating substances compared to the findings of Djordjevic et al. [64], who discovered ferric reducing antioxidant power of 49.43 nmol Fe2+/mg dry extract (recalculated to 49.43 μmol Fe2+/g dry extract). Furthermore, our results showed a 1.8-fold higher antioxidant power capacity compared to Estivi et al. [67], which was 34.68 mmol TE/kg DM (recalculated to 34.68 μmol Fe2+/g DM). Furthermore, our results for FRAP activity agreed with the findings of Zhu et al. [65], who measured FRAP values for six buckwheat varieties in the range of 31.81–87.67 μmol Fe2+/g DM, with Novosadska variety buckwheat demonstrating higher antioxidant power capacity. The decrease in FRAP values over the 9-month storage period can be attributed to strong antioxidative activity, which is based on a much less selective reduction that resulted in the consumption of electron-donating substances present in flour [64].
Generally, buckwheat seeds exhibit very high DPPH· activity due to their high polyphenolic content, which allows OH-groups to donate H-atoms to free radicals or peroxide [63]. Zhu et al. [65] in their study found DPPH· antioxidant activities in the range of 25.23–119.56 μmol TE/g DM, which were 1.3–6.1 times higher compared to our initial results (0 months). Moreover, a possible explanation for these differences in DPPH· radical-scavenging activity lies in inequality in the phytochemical composition of seeds that quantitatively and qualitatively depends on genotypes and environmental factors [65]. Observing a decreasing tendency in activity over time, Starowicz and Zieliński [66] found a reduction of 7% over 18 months, which was significantly less than the obtained reduction of 58% in DPPH· activity over 9 months in our study. This can be explained by different storage conditions. Starowicz and Zieliński [66] conducted the experiment at a lower temperature of 23 °C, which did not lead to thermally aggressive degradation of the antioxidant compound and thus to a decrease of scavenging ability, although the type and amount of used flour and the structure and interaction among the antioxidants are also important [2,64]. The reduction in DPPH·radicals reveals that examined materials contain radical inhibitors or scavengers with the possibility to act as primary antioxidants [63].
4.3. Phenolic Acids and Bioflavonoids Profiles
Buckwheat seeds contain many bioactive compounds (about 180) that, alongside essential nutrients, contribute to positive health benefits.
In addition to one phenyl group, phenolic acids have one carboxylic group and one or more hydroxyl groups and are classified as hydroxycinnamic acids (derived from cinnamic acid) and hydroxybenzoic acids (derived from benzoic acid) [68]. The roles of phenolic acids in plants are numerous and various, as these specialized compounds serve as signaling molecules in plant-microbe interactions, attract pollinators and seed dispersers, provide UV protection, or regulate growth and development processes, survival, and adaptation to environment [2,68]. Mostly, they influence or are involved in biological, biochemical, and chemical processes, such as the induction of oxidative stress, the modification of cell division and permeability or the alteration of photosynthesis, respiration, and transpiration, as well as the modulation of gene expression, protein biosynthesis, phytohormone, and enzyme activities [68]. Observed through the ability of antioxidant action, phenolic acids scavenge free radicals and other reactive oxygen species, helping with the protection of cells from oxidative damage. Likewise, phenolic acids contribute to sensory characteristics, primarily color and flavor [68]. In our research, gallic, caffeic, p-coumaric, ferulic, and trans-cinnamic acids were initially found in high quantities (0 months) (Table 5), which agreed with the findings of other researchers [14,68,69] who found a predominant content of these acids in buckwheat seeds and flour. Compared to our findings, Zhu et al. [65] discovered in all examined samples of buckwheat flour a similar average content of gallic acid, with two exceptions of lower and higher content (59.79 vs. 135.67 μg/g DM). In contrast, our results for Novosadska variety buckwheat were not in agreement with the findings of Beitāne et al. [14] and Škrobot et al. [70], who found 827 and 24 times lower total content of gallic acid in buckwheat flour, respectively. Similarly, for ferulic acid content, the researchers [14,65,70] found significantly lower amounts 209, 13, and 12 times, respectively. For caffeic acid content, our results were comparable with the findings of Škrobot et al. [70], who found a slightly higher total value (recalculated for comparison to μg/g DM) of 48.08 μg/g DM (free 42.73 and bound 5.35 μg/g DM), but these values were not in agreement with the findings of Beitāne et al. [14] who determined 1.72 μg/g DM (recalculated) in buckwheat flour, which was 22 times lower. Our findings for p-coumaric acid content disagreed with the results of Beitāne et al. [14] and Škrobot et al. [70], who found 4.46 μg/g DM and a total of 2.34 μg/g DM (free 1.49 and bound 0.85 μg/g DM), respectively, which were 23 and 44 times lower. The content of trans-cinnamic acid found in our research was 391 times higher than that (0.23 μg/g DM) examined by Škrobot et al. [70] and 11.5 times higher than that (7.86 μg/g DM) determined by Zieliński et al. [69] in biscuits made with whole buckwheat flour. The chlorogenic acid content examined in our research was multiple times higher (22.5- and 8-fold) than the content (0.64 μg/g DM and 1.86 μg/g DM) reported by other researchers [14,70]. All differences in the phenolic acid content between studies could be explained by the different degree of release from the food matrix or pH-dependent transformations, the interactions between them and food components, as well as by the existence of differences in genotypes and species, the influence of environmental factors and climate changes, storage conditions, etc. [2,65,69]. Dihydrocaffeic and phloretic acids are products of the reduction of hydroxycinnamic acids. Dihydrocaffeic acid is known as a metabolite of caffeic acid. It is based on catechol structure and shows powerful antioxidant activity via an antiradical effect, even higher than that of α-tocopherol [71]. Previous studies were mainly focused on its monitoring in various plant species, especially in the flowers of the rainforest tree Polyscias murrayi, where the highest amount of 352.32 mg/kg was recorded, as well as on the observation of the presence in human plasma [72,73]. On the other hand, De Pasquale et al. [71] conducted a study on semolina-pasta fortified with fermented black chickpea flour, where dihydrocaffeic acid was not detected in unfermented black chickpea doughs. In contrast, our study identified this acid in Novosadska variety buckwheat flour at a concentration of 18.7 μg/g DM. This finding adds new data to the list of well-studied phenolic acids that contribute to the overall antioxidant potential of buckwheat seeds. Observed phloretic acid belongs to the class of organic compounds named phenylpropanoic acids. In the same study [71], phloretic acid also was not detected, which is contrary to our findings of 25.8 μg/g DM. It could also be added to the list of phenolic acids that contribute to the overall antioxidant potential of buckwheat flour. Hesperetic acid, one of the hesperetin metabolites, was detected in buckwheat leaves by Sytar et al. [74] at approximately 0.12 mg/g DM. However, in our study, this acid was not detected, suggesting that hesperetic acid may accumulate more in the leaves rather than in the seeds (or flour). Over a 12-month storage period, Škrobot et al. [70] found a significant increase in the total content of gallic, caffeic, and p-coumaric acids, which was in contrast with our findings where no significant changes were observed during 9 months of storage, though the values of gallic and p-coumaric acids were noticeably higher, by about 6 and 26 times, respectively. Conversely, in our research, the content of caffeic acid was slightly lower, 1.4 times lower than in the findings of mentioned researchers [70]. Considering the trends of total content in a chlorogenic acid increase and a ferulic acid decrease, which were determined by Škrobot et al. [70] in buckwheat flour, there was a discrepancy with our findings regarding the declining chlorogenic acid and unchanging ferulic acid content, although both were present in higher quantities. Storage conditions, such as high temperature that easily oxidizes and degrades chlorogenic acid, and biochemical processes based on its interactions with carbohydrates, amino acids, and protein side chains, as well as scavenging and inhibition of free radicals in the process of lipid oxidation, can reduce the content. The non-significant variability of the trans-cinnamic acid content observed in our study during storage agreed with the findings of the previous study [70], in which about 284 times lower content was found. No appropriate comparative studies on buckwheat flour were found for the discussion on the content changes in dihydrocaffeic, phloretic, and hesperetic acids during storage. All the compared content changes did not follow the same pattern and could be explained by enzyme activity, oxidation processes caused by storage conditions or conversion of one compound into another [70]. It is important to note that slight variations in the content of trans-cinnamic acid, p-coumaric acid, caffeic acid, and chlorogenic acid in sections 0, 3, 6, and 9 months in buckwheat flour, as presented in Table 5, among others, may occur following the free phenolics inter-transformation pathway: trans-cinnamic acid via p-coumaric acid and caffeic acid to chlorogenic acid [75]. In addition, the enzyme phenylalanine ammonia-lyase (PAL), which plays a key role in phenolic biosynthesis, along with enzymes capable of oxidizing phenolic acids, such as polyphenol oxidases (PPOs) and peroxidases (PODs) and laccases, may contribute to the reduction of phenolic acid content during storage. This is particularly true for phenolic acids with catechol groups, as both types can oxidize o-diphenols into o-quinones in the presence of molecular oxygen [76].
Buckwheat seeds are particularly known for their high flavonoid content, which consists of secondary metabolites and naturally occurring antioxidants, with the main structure that includes two phenyl rings connected by a heterocyclic pyran ring [2,53,68]. Regarding the flavonoid content in Novosadska variety buckwheat flour, the results showed that the examined compounds (Table 5) were present in relatively high quantities in both fresh and stored samples, with naringin going undetected at the beginning. Quercetin, a precursor of rutin, is characterized as a bitter compound [53,68]. The initial results for quercetin demonstrated a partial agreement with a quote by Zamaratskaia et al. [2] about Tartary buckwheat flour due to the relatively high concentrations found. However, our results diverge from those of another study [70], which reported quercetin in common buckwheat whole-seed flour at a concentration four times lower (2.39 μg/g DM) or not detected at all [77]. Catechin is also a very valuable buckwheat constituent, featuring a high antioxidant capacity. In the research of Škrobot et al. [70], the initial total catechin content of 76.76 μg/g DM (free 68.99 and bound 7.77 μg/g DM) was almost two times higher than in our investigation. Novosadska variety buckwheat flour contained 4 and 5 times higher epicatechin content than that found in previous studies [14,70], respectively, and reaffirmed by the mentioned quote of researchers [2] where it was identified as dominant in three varieties of buckwheat (flour). There is limited data on daidzein, naringin, and naringenin as bioactive compounds in buckwheat seeds (flour) or generally in the buckwheat plant. To perform the promotion of the Novosadska variety buckwheat in a certain way, some comparison of daidzein with other similar pseudocereals, such as quinoa, was made. Daidzein is the respective aglycone of glycoside daidzin. Novosadska variety buckwheat seeds initially showed a high concentration of daidzein, almost 3-fold higher than in quinoa seeds (Chenopodium quinoa Willd.), where it varied between 7.0 and 20.5 μg/g (recalculated for comparison to μg/g DM) [78]. Naringin and its aglycone form, naringenin, could be found in the leaves of buckwheat. Dihydrokaempferol and dihydroquercetin are formed by enzymatic catalysis from naringenin, which is then further converted into kaempferol and quercetin [79]. In a previous study [46], the authors reported concentration ranges for naringin and naringenin in highland barley grains as 2.37–6.63 μg/g and 5.32–9.08 μg/g, respectively. These values are generally several times lower (at least 6 and 4 times, respectively) compared to those found in Novosadska variety buckwheat seeds, except for freshly harvested seeds where naringin was not detected. Regarding storage time, our results align with the findings of Škrobot et al. [70], who observed an increasing trend in quercetin content. However, our study reported a 13-fold higher value in Novosadska variety flour at 9 months compared to the levels reported in their study after one year of storage. The same researchers [70] found an increasing trend in total epicatechin and total catechin content, which is in contrast with our findings on decreasing epicatechin and unchanging catechin content. In the same study, the total epicatechin was twice as low, and the total catechin content was twice as high compared to our results at the 9-month storage mark. Regarding the content of daidzein, naringin, and naringenin during the nine-month storage in relation to the stated amounts in the papers [78,79], at least 3.4 times more daidzein was found than in quinoa seeds, i.e., at least 7.8 and 4.2 times more naringin and naringenin than in highland barley grains. All differences and changes in the quantities of mentioned bioflavonoids, including disagreements with the results of other researchers, directly depend on factors such as the difference in growth factors of the seed, size and shape of the seed, the color of the flower, and sowing time, buckwheat varieties, soil quality and location, environmental fluctuations, climate changes, growth stages, area of collection, period and storage conditions, transformations and interactions between compounds, etc. [2,53]. Enzyme-mediated biochemical reactions and environmental stimuli determine and affect the transformation of glucose into three molecules of malonyl-CoA, which are used to synthesize ring A. Meanwhile, the shikimate pathway produces phenylpropanoids through the production of the amino acid phenylalanine, which generates 4-coumaroyl-CoA molecules for ring B. These rings merge through a linking chain called the C ring. Another factor might be the conversion or release of glucosides or the heating time required to break down cellular constituents and release individual bioflavonoids. Additionally, it is important to note that to the best of our knowledge, this is the first report on the contents of daidzein, naringin, and naringenin in buckwheat seeds (flour).
The preserved approximate and nutritional properties of Novosadska variety buckwheat, as well as the antioxidant activity and capacity during storage, provide the possibility of its equal use in the bakery industry (flour) mixed in a certain ratio, but taking into account that the absence of gluten-forming proteins may change the finished product. Similarly, the proven quality and health benefits of buckwheat promote and facilitate its application in the development of the functional food industry, bearing in mind the inhibition of oxidation processes, which contributes to extending the shelf life of end-products. Based on our research, a similarity with studies [32,33] was observed in the potential of the underutilized buckwheat crop to be included in a widely accepted diet. Buckwheat fully meets the needs in terms of quality and health-promoting properties as a sustainable and biodiversity-friendly crop. Further research is needed to improve the properties of buckwheat polyphenols and to increase the content of macro- and micronutrients in seeds based on new approaches through procedures, genetics, and industrial processing.
Regarding the proximate and nutritional composition, Novosadska variety buckwheat seeds (flour) showed no significant changes even after 9 months of storage at 40 ± 2 °C. A considerable amount of phenolic acids, such as gallic, ferulic, p-coumaric, trans-cinnamic, and chlorogenic, conjugated polyphenols and bioflavonoids contribute to the high potential of antioxidant activity and capacity demonstrated via at least 1.8-fold higher ferric ion-reducing antioxidant power (FRAP) after 9-month storage. Bioflavonoids quercetin and epicatechin were present in amounts 4–5 times higher than in the well-known common buckwheat whole seeds and other flours. Dihydrocaffeic and phloretic acids, daidzein, naringin, and naringenin, were quantified in buckwheat flour for the first time.
5. Conclusions
This research provides insight and emphasizes the paramount importance of storage in maintaining sufficient seeds across all seasons. Our study highlights the potential of Novosadska variety buckwheat as a sustainable and biodiversity-friendly crop, emphasizing its value for extended cultivation in agricultural practices. The results showed that Novosadska variety buckwheat seeds maintained high quality and health-promoting properties, including significant phenolic acids and bioflavonoids, without significant loss of antioxidant activity after 9 months of storage at 40 ± 2 °C. These seeds compare favorably with other commercial buckwheat varieties and cereals such as wheat and barley. However, the content of dihydrocaffeic, chlorogenic, and phloretic acids, as well as epicatechin, daidzein, quercetin, and naringin, showed significant impacts due to the storage period. It is important to note that, to the authors’ knowledge, dihydrocaffeic and phloretic acids have been quantified in buckwheat flour for the first time for the Novosadska variety, along with the report on the contents of daidzein, naringin, and naringenin.
Due to its adaptability to extreme environmental and climate conditions, ease of application in agroecological farming systems, ability to attract a variety of pollinators and provide shelter and food for beneficial insects, and as a speedy short-season plant which can give a relatively quick response to the growing demand for food of expanding population, it is necessary to promote and unlock the full potential of buckwheat (locally, Novosadska variety). The acceptance of value-added buckwheat end-products by consumers, an essential factor in the consumption process, opens opportunities for improving sustainability capabilities. As a pseudocereal with many nutritional and health benefits and all the aforementioned properties, abilities, and roles, buckwheat stands out as a sustainable and biodiversity-friendly crop.
Conceptualization, B.P. and R.R.; methodology, B.P., S.Đ., Z.Ž.S. and G.K.; software, Z.Ž.S. and T.M.; validation, B.P., S.Đ. and G.K.; formal analysis, B.P., S.Đ., R.R. and G.K.; investigation, R.R. and B.P.; resources, B.P. and S.R.; data curation, B.P.; writing—original draft preparation, B.P.; writing—review and editing, S.Đ., Z.Ž.S. and S.R.; visualization, T.M.; supervision, B.P. and T.M.; project administration, T.M.; funding acquisition, all authors. All authors have read and agreed to the published version of the manuscript.
All relevant data are included in the manuscript. All other data are available from the corresponding author upon reasonable request.
The authors declare no conflicts of interest. The funders had no role in the design of the study, in the collection, analyses, or interpretation of data, in the writing of the manuscript, or in the decision to publish the results.
Footnotes
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Enlarge this image.Figure 1. A flowchart of sampling, storage conditions, sample preparation, and analysis.
Enlarge this image.Figure 1. A flowchart of sampling, storage conditions, sample preparation, and analysis.
Enlarge this image.Figure 2. Surface plot of significant (a) phenolic acids and (b) bioflavonoids concentration in function with storage time (months) and temperature, °C.
Enlarge this image.Figure 3. Chromatogram of phenolic acids and bioflavonoids of one Novosadska variety buckwheat flour sample at 3 months of storage (the sequence of peaks (a) at 280 nm: 1—Gallic acid, 2—Catechin, 3—Dihydrocaffeic acid, 4—Epicatechin, 5—Phloretic acid, 6—Naringin, 7—Daidzein, 8—trans-Cinnamic acid, 9—Naringenin; (b) at 325 nm: 10—Chlorogenic acid, 11—Caffeic acid, 12—p-Coumaric acid, 13—Ferulic acid, 14—Quercetin, Hesperetic acid not detected).
The summarized method quality parameters.
| Parameters | |||||||
|---|---|---|---|---|---|---|---|
| Compound | Linearity, | Repeatability (RSD, %) * | Limit of Repeatability (μg/g DM) * | Bias, | LOD (μg/g DM) * | LOQ (μg/g DM) * | MU, Uexp |
| Phenolic acids | |||||||
| Gallic acid | P = 30,578.9 × C − 162,570, 0.9997 | 8.0 | 20.87 | 8.8 | 0.34 | 1.04 | 16.4 |
| Dihydrocaffeic acid | P = 9498.89 × C − 56,038.4, 0.9997 | 10.1 | 4.59 | 1.0 | 0.25 | 0.76 | 20.4 |
| Chlorogenic acid | P = 35,960.3 × C − 197,214, 0.9998 | 9.7 | 2.70 | −5.0 | 0.34 | 1.03 | 19.7 |
| Caffeic acid | P = 63,744.5 × C − 354,944, 0.9997 | 10.1 | 10.82 | 1.9 | 0.35 | 1.07 | 20.4 |
| Phloretic acid | P = 5931.32 × C − 30,952.6, 0.9998 | 9.3 | 8.57 | −0.1 | 0.33 | 0.98 | 18.8 |
| p-Coumaric acid | P = 59,917.0 × C − 321,082, 0.9998 | 7.0 | 19.69 | 9.8 | 0.33 | 0.98 | 14.4 |
| Ferulic acid | P = 61,344.4 × C − 338,195, 0.9998 | 6.7 | 10.61 | 4.9 | 0.33 | 0.99 | 13.8 |
| Hesperetic acid | P = 66,213.6 × C − 362,089, 0.9998 | 8.0 | 5.6 | −2.0 | 0.32 | 0.98 | 16.3 |
| trans-Cinnamic acid | P = 110,419 × C − 463,141, 0.9987 | 5.1 | 12.71 | 3.0 | 0.79 | 2.40 | 10.7 |
| Bioflavonoids | |||||||
| Catechin | P = 8304.41 × C − 46,342.0, 0.9997 | 8.4 | 11.70 | −7.7 | 0.38 | 1.15 | 17.1 |
| Epicatechin | P = 8107.73 × C − 52,057.6, 0.9998 | 8.3 | 18.00 | −6.4 | 0.33 | 1.01 | 16.9 |
| Daidzein | P = 28,430.8 × C + 11,326.8, 0.9996 | 7.2 | 12.12 | 3.9 | 0.39 | 1.18 | 14.8 |
| Quercetin | P = 14,806.9 × C + 51,060.0, 0.9935 | 9.6 | 5.49 | −5.0 | 0.41 | 1.24 | 19.5 |
| Naringin | P = 21,336.8 × C − 121,518, 0.9998 | 7.2 | 10.11 | −3.1 | 0.34 | 1.04 | 14.8 |
| Naringenin | P = 44,072.3 × C − 404,501, 0.9986 | 6.3 | 6.78 | 6.4 | 0.82 | 2.48 | 13.0 |
*—average values; R2—correlation coefficient; RSD—relative standard deviation; P—peak area; C—concentration of compound, ml/L; LOD—limit of detection; LOQ—limit of quantification; MU—measurement uncertainty, Uexp—expanded measurement uncertainty; DM—dry matter.
The summarized instrument performance parameters.
| Item | ||
|---|---|---|
| Instrument | Parameters | Results |
| Pump | Flow precision, %RSD | 0.18 |
| Flow accuracy (bias), % | 1 | |
| Gradient accuracy (bias) AB/CD port, % | −2.3–1.1 | |
| Injector | Injected volume precision, %RSD | 0.08 |
| Injected volume linearity | R2 = 1.0000; 0.78%RSD | |
| Carry-over, % | 0.000349 | |
| Column | Temperature precision, %RSD | −0.75 |
| Temperature accuracy (bias), % | −0.8 | |
| Temperature stability, Δt | 0.2 °C | |
| Detector | Linearity response | R2 = 0.9999; 1.22%RSD |
| Wavelength accuracy, % | 100 | |
| Noise, AU | <5.0 × 10−5 | |
| Drift, AU/h | <2.0 × 10−3 | |
Influence of storage time (S) on proximate composition, nutritional properties, pH values, and alcoholic acidity of Novosadska variety buckwheat flour.
| n = 9 | Storage Time (S) | |||
|---|---|---|---|---|
| Parameters | Freshly Harvested Seed | 3 Months | 6 Months | 9 Months |
| Moisture, % | 10.21 ± 0.006 d | 9.09 ± 0.020 c | 7.89 ± 0.066 b | 6.02 ± 0.021 a |
| Total protein, % DM | 13.46 ± 0.344 b | 12.62 ± 0.590 ab | 12.25 ± 0.336 a | 11.57 ± 0.448 a |
| Total lipids, % DM | 3.44 ± 0.152 b | 2.99 ± 0.303 ab | 2.64 ± 0.235 a | 2.99 ± 0.050 ab |
| Total ash, % DM | 1.97 ± 0.020 | 2.01 ± 0.020 | 1.94 ± 0.010 | 1.96 ± 0.050 |
| Total carbohydrates, % DM | 81.13 ± 0.510 a | 82.38 ± 0.676 ab | 83.17 ± 0.465 b | 83.48 ± 0.370 b |
| Starch, % DM | 62.55 ± 0.105 c | 49.12 ± 0.315 a | 47.67 ± 0.737 a | 52.12 ± 1.342 b |
| β-glucan, % DM | 0.050 ± 0.0013 b | 0.030 ± 0.0012 a | 0.100 ± 0.0028 c | 0.120 ± 0.0027 d |
| Energy value, kJ/100 g | 1558 ± 2.5 a | 1569 ± 6.1 a | 1584 ± 5.1 b | 1623 ± 1.2 c |
| pH value | 6.74 ± 0.059 c | 6.63 ± 0.046 b | 6.22 ± 0.025 a | 6.19 ± 0.021 a |
| Alcoholic acidity, % DM (as H2SO4) | 0.86 ± 0.028 c | 0.65 ± 0.035 b | 0.67 ± 0.025 b | 0.53 ± 0.031 a |
a, b, c, d Means within the same row with different superscripts differ significantly (p < 0.05); DM—dry matter.
Influence of storage time (S) on antioxidant activity and capacity of Novosadska variety buckwheat flour.
| n = 9 | Storage Time (S) | |||
|---|---|---|---|---|
| Parameters | Freshly Harvested Seed | 3 Months | 6 Months | 9 Months |
| Total phenol, mg GAE/g DM | 7.28 ± 0.194 c | 6.97 ± 0.115 c | 6.04 ± 0.121 b | 5.57 ± 0.122 a |
| FRAP, μmol Fe2+/g DM | 63.45 ± 1.520 d | 28.68 ± 0.476 c | 25.32 ± 0.445 b | 22.20 ± 0.440 a |
| DPPH, μmol TE/g DM | 19.47 ± 0.907 c | 19.66 ± 0.333 c | 14.02 ± 0.490 b | 8.12 ± 0.344 a |
a, b, c, d Means within the same row with different superscripts differ significantly (p < 0.05); DM—dry matter.
Influence of storage time (S) on phenolic acids and bioflavonoids profile of Novosadska variety buckwheat flour.
| n = 9 | Storage Time (S) | |||
|---|---|---|---|---|
| Parameters | Freshly Harvested Seed | 3 Months | 6 Months | 9 Months |
| Phenolic acids | ||||
| Gallic acid, μg/g DM | 91.0 ± 8.44 | 92.7 ± 6.97 | 94.1 ± 6.88 | 93.6 ± 7.53 |
| Dihydrocaffeic acid, μg/g DM | 18.7 ± 1.93 b | 11.5 ± 1.16 a | 19.1 ± 1.95 b | 15.5 ± 1.52 ab |
| Chlorogenic acid, μg/g DM | 14.4 ± 1.29 c | 11.1 ± 1.10 b | 7.6 ± 0.74 a | 7.1 ± 0.72 a |
| Caffeic acid, μg/g DM | 38.2 ± 3.83 | 37.6 ± 3.97 | 39.2 ± 3.78 | 38.3 ± 3.87 |
| Phloretic acid, μg/g DM | 25.8 ± 2.44 a | 46.5 ± 4.11 b | 31.4 ± 3.04 a | 29.1 ± 2.65 a |
| p-Coumaric acid, μg/g DM | 102.4 ± 7.52 | 101.9 ± 7.57 | 97.8 ± 6.14 | 95.9 ± 6.90 |
| Ferulic acid, μg/g DM | 56.5 ± 3.79 | 57.0 ± 3.56 | 57.4 ± 4.00 | 56.5 ± 3.80 |
| Hesperetic acid, μg/g DM | nd | nd | nd | nd |
| trans-Cinnamic acid, μg/g DM | 90.0 ± 4.76 | 89.9 ± 4.53 | 89.5 ± 4.27 | 87.9 ± 4.60 |
| Bioflavonoids | ||||
| Catechin, μg/g DM | 49.1 ± 4.12 | 52.2 ± 4.28 | 49.9 ± 4.12 | 49.3 ± 4.20 |
| Epicatechin, μg/g DM | 95.7 ± 7.87 c | 78.5 ± 6.31 b | 77.0 ± 6.46 b | 59.8 ± 5.08 a |
| Daidzein, μg/g DM | 57.0 ± 4.25 a | 59.3 ± 4.25 ab | 56.1 ± 3.84 a | 69.3 ± 4.98 b |
| Quercetin, μg/g DM | 9.8 ± 0.93 a | 12.5 ± 1.27 ab | 16.6 ± 1.69 b | 47.1 ± 3.95 c |
| Naringin, μg/g DM | nd | 42.7 ± 2.91 a | 54.9 ± 4.22 b | 51.9 ± 3.70 b |
| Naringenin, μg/g DM | 38.4 ± 2.78 | 37.8 ± 2.33 | 38.7 ± 2.24 | 38.3 ± 2.33 |
a, b, c Means within the same row with different superscripts differ significantly (p < 0.05); nd—not detected; DM—dry matter.
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; Rakić, Radojica 2 ; Rakić, Sveto 2 ; Sekulić, Zoran Ž 3
; Milićević, Tijana 4
; Kulić, Gordana 2 ; Đurović, Sanja 1
1 Institute for Plant Protection and Environment, Teodora Drajzera 9, 11040 Belgrade, Serbia;
2 Faculty of Agriculture, University of Belgrade, Nemanjina 8, 11080 Belgrade, Serbia;
3 Institute of Public Health of Belgrade, Bulevar Despota Stefana 54a, 11108 Belgrade, Serbia;
4 Institute of Physics Belgrade, National Institute of the Republic of Serbia, University of Belgrade, Pregrevica 118, 11080 Belgrade, Serbia;