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. 2026 May 4;68(9):3260–3271. doi: 10.1111/jipb.70282

Genome editing generates high oleic soybean and eliminates beany flavors

Hongtao Xie 1,2,3,4, † , Leping Geng 1,2,3, † , Ziyi Hu 1,2,3, Feiyang Chen 1,2,3, Mingfang Tan 1,2,3, Dagang Wang 5, Zhiping Huang 5, Weijun Ye 5, Peng Chen 6, Jianhua Zhu 1,2,3,✉
PMCID: PMC13545845  PMID: 42081667

ABSTRACT

Soybeans serve as excellent sources of vegetable oil, protein, and other valuable nutrients for human consumption, materials for diverse industries, including the cosmetics and medical industries, and feedstocks for animals. Nevertheless, some people do not favor soy oil or other various food products derived from soybeans, due to inadequate levels of oleic acid in the oil and the presence of undesirable grassy and beany flavors associated with oxidation products of polyunsaturated fatty acids in the seeds. In this study, we developed soybean cultivars with very high levels of oleic acid in the seeds, and without grassy and beany flavors. We achieved this by using CRISPR‐Cas‐SF01 to edit genes in the elite cultivar Xudou 18 (XD18), targeting two microsomal Δ‐12 fatty acid desaturase 2 (GmFAD2‐1A and GmFAD2‐1B) and three lipoxygenase (GmLOX1, GmLOX2, and GmLOX3) genes. Our findings showed that fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants performed similarly to XD18 plants in the field, indicating no obvious growth penalties. Overall, this research has demonstrated that the development of soybean germplasms with high levels of oleic acid and without undesirable beany flavors through gene‐editing of multiple relevant genes is effective, and this endeavor can contribute to the health of a broader global consumer population.

Keywords: Cas‐SF01, grassy and beany flavors, lipoxygenase, oleic acid, soybean, Δ‐12 fatty acid desaturase 2

Cas‐SF01‐mediated editing of two Δ‐12 fatty acid desaturase and three lipoxygenase genes in the elite soybean germplasm XD18 increased oleic acid to 80% and eliminated beany flavor, yielding improved soybeans that support broader consumer health without compromising field performance.

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INTRODUCTION

Soybean (Glycine max L. Merrill) is a crucial economic crop, contributing approximately 29% of vegetable oil and 71% of protein meals globally. Soybean oil is extracted from seeds via diverse pressing methodologies. Composed predominantly of triacylglycerol, which is synthesized through the esterification of three fatty acids to a glycerol backbone, soybean oil exhibits remarkable antioxidant characteristics due to the presence of α‐, δ‐, and γ‐tocopherols, isoflavones, and their derivatives. The nutritional and utilization values of soybean oil are ultimately contingent upon the fatty acid composition within the seeds.

Commercial soybean oil typically encompasses five primary fatty acids: 10%–11% palmitic acid (16:0), 4% stearic acid (18:0), 18%–25% oleic acid (18:1), 52%–55% linoleic acid (18:2), and 8%–13% linolenic acid (18:3) (Fehr, 2007; Clemente and Cahoon, 2009). The relatively high content of linoleic acid (a ω‐6 fatty acid) in soybean oil may potentially have a negative nutritional impact, as diets abundant in linoleic acid tend to lead to a reduction. It is widely acknowledged that augmenting the dietary intake of the ω‐6 polyunsaturated fatty acid arachidonic acid (ARA) or its precursor linoleic acid will heighten inflammation. This is because ARA serves as a precursor to a series of potent pro‐inflammatory mediators, such as the well‐documented prostaglandins and leukotrienes (Innes and Calder, 2018). The nutritionally beneficial effects of health‐promoting ω‐3 fatty acids, such as stearidonic acid (SDA; 18:4Δ6,9,12,15), eicosapentaenoic acid (EPA; 20:5Δ5,8,11,14,17), and docosahexaenoic acid (DHA; 22:6Δ4,7,10,13,16,19) in plant tissues have been reported (Clark et al., 1992; Friesen and Innis, 2010; Blasbalg et al. 2011). Additionally, the high concentrations of linoleic acid and linolenic acid are unfavorable for food production due to their low oxidative stability, which causes foods to develop rancidity and off‐flavors rapidly (Warner and Gupta 2005). It has been documented that the dietary consumption of plant‐based oils abundant in oleic acid (a monounsaturated ω‐9 fatty acid), such as olive oils, can reduce total cholesterol, low‐density lipoprotein (LDL), the ratio of total cholesterol to high‐density lipoprotein (HDL), and blood pressure, while simultaneously inhibiting platelet aggregation (Terés et al., 2008; Carrillo et al., 2012; Yubero‐Serrano et al., 2019). A growing body of evidence indicates that the anti‐inflammatory and antioxidant properties of oleic acid may be associated with the actions of oleoyethanolamide, a derivative of oleic acid (Santa‐Maria et al., 2023). Additionally, polyphenols (a constituent of the plant‐based oil) act in synergy with oleic acid to postpone postprandial lipemia, thus decreasing the risk of cardiovascular diseases (Yubero‐Serrano et al., 2019; Chrysant and Chrysant, 2024). Furthermore, oleic acid itself may potentially impede the risk of cancer by suppressing the oncogene HER2 or by restoring the impaired antitumor immunity of γδ‐T cells induced by palmitic acid (Colomer and Menéndez, 2006; Zhang et al., 2025). Therefore, to enhance the quality of soybean oil and processed food products, chemical hydrogenation has been utilized to elevate the oleic acid content to approximately 50% and decrease the quantity of polyunsaturated fatty acids like linolenic acid (Yusem and Pintauro 1992). Oil hydrogenation produces trans‐fatty acids, which have been associated with cardiovascular diseases, elevated levels of “bad” cholesterol (low‐density lipoproteins), and a higher risk of developing type 2 diabetes (Mozaffarian et al., 2006; Danaei et al., 2009). Another effective strategy to enhance the functionality of soybean oil without hydrogenation is to genetically augment the oleic acid content in soybean seeds while reducing the linoleic and linolenic acid contents. In the lipid biosynthetic pathway, the Δ‐12 fatty acid desaturase 2 (GmFAD2) is accountable for the transformation of oleic acid to linoleic acid (Heppard et al., 1996; Schlueter et al., 2007). In developing soybean seeds, among the identified FAD2 genes, the two microsomal GmFAD2‐1 desaturases encoded by GmFAD2‐1A (Glyma.10g42470) and GmFAD2‐1B (Glyma.20g24530) exhibit the highest expression levels (Tang et al., 2005). Therefore, GmFAD2‐1A and GmFAD2‐1B are regarded as playing a crucial role in regulating the oleic acid level in developing soybean seeds and were employed as targets or candidate genes to produce high oleic acid soybeans. Two methods were utilized to generate soybeans with an oleic acid content exceeding 80% among the total fatty acids. Methods such as genetic engineering and candidate gene‐based molecular breeding have been employed (Buhr et al., 2002). Nevertheless, in the commercial market, there is a scarcity of high oleic acid soybean oil derived from germplasms generated via induced mutations in GmFAD2 genes through X‐ray irradiation or EMS mutagenesis. This is primarily attributable to the yield reductions associated with these germplasms (Bachlava et al., 2008; Scherder and Fehr, 2008).

Lipoxygenases are enzymes that belong to a group of non‐heme iron‐containing proteins, which are extensively distributed in plants, fungi, and animals (Brash, 1999). In plants, lipoxygenases can be detected in all organs, accounting for up to 2% of soybean seed proteins (Loiseau et al., 2001). Lipoxygenases catalyze the oxidation of polyunsaturated fatty acids, such as linoleic acid (18:2) and α‐linolenic acid (18:3), to generate unsaturated fatty acid hydroperoxides (Song et al. 1990; Brash, 1999; Liavonchanka and Feussner, 2006). The products of the lipoxygenase pathway have been demonstrated to play a role in a variety of plant processes, including vegetative growth, responses to wounding, herbivore attacks, and pathogen invasions, and are regulated by various factors (Porta and Rocha‐Sosa, 2002). In germinating soybean seeds, it has been determined that lipoxygenases do not participate in lipid mobilization (Wang et al., 1999). The limited oxygenation of polyunsaturated fatty acids during the germination process provides evidence suggesting that soybean seed lipoxygenases may have evolved to serve as storage proteins, despite retaining their intact yet non‐functional enzymatic capabilities (Siedow, 1991; Wang et al., 1999). Irrespective of the specific physiological role of lipoxygenases in soybean seeds, it has been shown that lipoxygenase‐free seeds can develop into normal, defect‐free plants (Hajika et al., 1992; Wang et al., 1999).

Seed lipoxygenases hold considerable significance in the food industry, and there is a market demand for lipoxygenase‐free soybean varieties. The oxidation products generated by seed lipoxygenase activity have been linked to the development of unpleasant grassy and beany flavors in products containing protein derived from soybean and other leguminous seeds (Sessa, 1979; Robinson et al.,1995; Wilson, 1996; Gerde and White, 2008). Mature soybean seeds predominantly contain three lipoxygenases, namely GmLOX1 (Glyma.13g347600), GmLOX2 (Glyma.13g347500), and GmLOX3 (Glyma.15g026300) (Axelrod et al., 1981).

In this investigation, the effects of the loss‐of‐function of two GmFAD2‐1 genes (GmFAD2‐1A and GmFAD2‐1B) and three lipoxygenase genes (GmLOX1, GmLOX2, and GmLOX3) were systematically examined. The objective was to develop a soybean variety with seeds characterized by high levels of oleic acid and devoid of grassy and beany flavors. The CRISPR‐Cas‐SF01 genome editing tool was utilized to generate fad2‐1a/b and fad2‐1a/b/lox1/2/3 mutants in the Xudou 18 (XD18) cultivar. When these mutant plants were cultivated in the laboratory or in the field, their performance was essentially comparable to that of XD18. The oleic acid content in the mature seeds of the fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants reached up to 80% of the total fatty acids, and the seeds of the fad2‐1a/b/lox1/2/3 plants were free of grassy and beany flavors. Our findings indicate that, through the powerful genome‐editing tool, it is viable to develop soybean varieties with elevated levels of health‐beneficial oleic acid and devoid of beany flavor in the seeds. This would be of great significance in offering an alternative food option for individuals who seek the health advantages of high oleic acid but are averse to the grassy and beany flavors of soybean products.

RESULTS

Development of soybean plants with high oleic acid content and devoid of grassy and beany flavors

The objective of this study was to generate soybean plants with high oleic acid levels and devoid of grassy and beany flavors using the Cas‐SF01 genome editing tool. The superior soybean variety Xudou 18 (XD18) from the Huang‐Huai‐Hai Region was selected as the genetic background. Two Δ‐12 fatty acid desaturase genes (GmFAD2‐1A and GmFAD2‐1B) and three lipoxygenase genes (GmLOX1, GmLOX2, and GmLOX3) were targeted (Figures 1A, B, S1; Nelson and Cox, 2017). Homozygous fad2‐1a/b plants were obtained in the E3 generation, with a 10‐bp deletion in the GmFAD2‐1A locus and a 12‐bp deletion in the GmFAD2‐1B locus, respectively (Figures 1C, S2). T‐DNA‐free fad2‐1a/b plants were identified in the E4 generation, and these plants, along with their progeny, were utilized in subsequent experiments. During the soybean genetic transformation process, two independent and homozygous lines of fad2‐1a/b/lox1/2/3 plants were successfully obtained in the E6 generation (Figure 1C). The fad2‐1a/b/lox1/2/3 #1 plants exhibited multiple mutations (two deletions and one substitution) in GmFAD2‐1A, a 9‐bp deletion in GmFAD2‐1B, a 7‐bp deletion in GmLOX1, a 129‐bp deletion in GmLOX2, and a 21‐bp deletion in GmLOX3, respectively (Figures 1C, S2). Meanwhile, the fad2‐1a/b/lox1/2/3 #2 plants had multiple mutations (two deletions and one substitution) in GmFAD2‐1A, a 12‐bp deletion in GmFAD2‐1B, a 10‐bp deletion in GmLOX1, two deletions (11‐bp and 13‐bp, reflecting the gene editing effects of two separate crRNAs for this locus) in GmLOX2, and a 21‐bp deletion in GmLOX3, respectively (Figures 1C, S2). T‐DNA free fad2‐1a/b/lox1/2/3 #1 and fad2‐1a/b/lox1/2/3 #2 plants were obtained in the E7 generation, and these plants and their progeny were employed in subsequent experiments.

Figure 1.

Figure 1

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Generation of the fad2‐1a/b and fad2‐1a/b/lox1/2/3 mutants by targeting two Δ‐12 fatty acid desaturases (FAD2‐1A and FAD2‐1B) and three lipoxygenases (GmLOX1, GmLOX2, and GmLOX3) genes through the CRISPR‐Cas‐SF01 genome editing tool

(A) CRISPR RNA (crRNA) target sites for the GmFAD2‐1A, GmFAD2‐1B, GmLOX1, GmLOX2, and GmLOX3 genes. (B) Illustration of the T‐DNA portion of the binary vector used for soybean genetic transformation. N, nuclear localization signal; SpecR, coding region of a gene whose product confers resistance to spectinomycin. The section presenting a detailed description of the crRNA expression cassettes beneath the vector map was not drawn to scale. (C) Genotypes of the homozygous fad2‐1a/b and fad2‐1a/b/lox1/2/3 mutants were created in the genetic background of XD18.

Accumulation of metabolites in seeds of the fad2‐1a/1b and fad2‐1a/1b/lox1/2/3 plants

We utilized a Near‐infrared Grain Analyzer to determine total fatty acid levels to observe the effects of the fad2‐1a and fad2‐1b mutations on fatty acid accumulation in harvested soybean grains. We found that the total fatty acid levels in the seeds of the fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants were marginally higher than those in the seeds of the XD18, ZH301, and WD37 plants (Figure 2A). To elucidate the effects of fad2‐1a and fad2‐1b mutations on the fatty acid composition, the accumulation of saturated and unsaturated fatty acids in the seeds of the XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, fad2‐1a/b/lox1/2/3 #2, ZH301, and WD37 plants was determined. As anticipated, the linoleic acid levels in the seeds of the fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants were significantly lower than those in the seeds of the XD18, ZH301, and WD37 plants (Figure 2B). Consequently, the seeds of the fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants produced slightly lower amounts of linolenic acid compared to the seeds of the XD18, ZH301, and WD37 plants (Figure 2B). The seeds of the fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants contained up to 80% oleic acid, in contrast to the seeds of the XD18, ZH301, and WD37 plants (Figure 2B). Moreover, the seeds of the fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants produced slightly lower amounts of two saturated fatty acids (stearic acid and palmitic acid) than the seeds of the XD18, ZH301, and WD37 plants (Figure 2B). Furthermore, we determined the total protein levels in harvested seeds with the Near‐infrared Grain Analyzer, and found that the protein levels in the seeds of the fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, fad2‐1a/b/lox1/2/3 #2, and ZH301 plants were higher than those in the seeds of the XD18 and WD37 plants (Figure 2C).

Figure 2.

Figure 2

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Contents of total fatty acids and protein, as well as the composition of fatty acids, in the seeds of the fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants

(A) Total fatty acids in the seeds of the XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3, ZH301, and WD37 plants. (B) Compositions of saturated fatty acids in the seeds of the XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3, ZH301, and WD37 plants. (C) Protein contents in the seeds of the XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3, ZH301, and WD37 plants. Data are means ± SD (n = 5). The significant difference was calculated using one‐way ANOVA with the Tukey multiple comparison test. Different letters over the columns indicate significant differences (P < 0.05), and the same letters indicate no significant difference.

Total proteins were extracted from the seeds of the XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants, and the enzymatic activities of LOX1, LOX2, and LOX3 were determined. In comparison with the seeds of the XD18 and fad2‐1a/b plants, the activities of LOX1, LOX2, and LOX3 were substantially diminished in the seeds of the fad2‐1a/b/lox1/2/3 #1 and fad2‐1a/b/lox1/2/3 #2 plants (Figure 3A–F). Subsequently, we ascertained the accumulation of products derived from the oxidation reactions of unsaturated fatty acids and their derivatives, which frequently generate grassy and beany flavors (Figure S1; Nelson and Cox, 2017; Wang et al., 2021). The seeds of the fad2‐1a/b/lox1/2/3 #1 plants exhibited significantly lower levels of hexanal, (E)‐2‐hexenal, nonanal, 1‐hexanol, 1‐octen‐3‐ol, and nonanal compared to those of the XD18 plants (Figure 3G). The seeds of the fad2‐1a/b/lox1/2/3 #2 plants showed significantly lower levels of 1‐octen‐3‐ol and had no detectable levels of hexanal, (E)‐2‐hexenal, and 1‐hexanol, in contrast to the seeds of the XD18 plants (Figure 3G). The seeds of the fad2‐1a/b/lox1/2/3 #1 and fad2‐1a/b/lox1/2/3 #2 plants had no detectable levels of trans‐2‐octenal and trans‐2‐nonenal (Figure 3G). These findings imply that the grassy and beany flavors emitted from the seeds of the fad2‐1a/b/lox1/2/3 #1 and fad2‐1a/b/lox1/2/3 #2 plants were effectively diminished to a minimal level via the combination of the dysfunction of two Δ‐12 fatty acid desaturases and three lipoxygenases through gene editing.

Figure 3.

Figure 3

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Measurements of lipoxygenase activity and determination of grassy and beany flavors in the seeds of the XD18, fad2‐1a/b, and fad2‐1a/b/lox1/2/3 plants

(A–C) Colorimetric assays to determine the activity of LOX1, LOX2, and LOX3, respectively. (D–F) Quantification of LOX1 (as indicated by the absorption at 600 nm wavelength), LOX2 (as indicated by the absorption at 600 nm wavelength), and LOX3 (as indicated by the absorption at 452 nm wavelength) activities, respectively. (G) Measurements of volatile compounds in the seeds of the XD18, fad2‐1a/b, and fad2‐1a/b/lox1/2/3 plants, and these compounds are related to grassy and beany flavors. ND, not detectable. Data are means ± SD. n = 12 in (D–F); n = 3 in (G). The significant difference was determined using one‐way ANOVA with the Tukey multiple comparison test. Different letters over the columns indicate significant differences (P < 0.05), and the same letters indicate no significant difference.

Agronomic traits of the fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants

During the amplification of the seeds from the fad2‐1a/b/lox1/2/3 #1 and fad2‐1a/b/lox1/2/3 #2 in natural fields in Hainan Province, an analysis was conducted on the performance of these plants at maturity. The XD18, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants reached the same height (Figure S3A, B), and the fad2‐1a/b/lox1/2/3 #1 plants exhibited a slightly greater number of branches compared to the other genotypes (Figure S3C). The XD18, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants produced an equivalent number of nodes on the main stem, effective pods per plant, and had comparable 100‐seed weights (Figure S3D, E, G). The fad2‐1a/b/lox1/2/3 #1plants had seeds with a slightly greater weight per plant than the other genotypes (Figure S3F). The yields of the XD18, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants were identical (Figure S3H).

The performance of the fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants was evaluated in two provinces within the Huang‐Huai‐Hai Region: Shandong and Anhui. When cultivated in Shandong in 2024, the fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants demonstrated essentially the same performance as XD18 (Figure 4A–E). The fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants flowered at approximately the same time as XD18, and the fad2‐1a/b/lox1/2/3 plants required slightly less time to reach maturity than XD18 and fad2‐1a/b (Figure 4F, G). Additionally, the fad2‐1a/b/lox1/2/3 plants were slightly shorter and produced a slightly lower number of nodes on the main stem than XD18 and fad2‐1a/b (Figure 4H, J). Conversely, the fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants showed similar performance to XD18 in terms of the number of branches, number of effective pods, number of seeds per plant, and weight of seeds per plant (Figure 4I, K–M). Moreover, the 100‐seed weight of the fad2‐1a/b/lox1/2/3 plants was slightly lower than that of fad2‐1a/b (Figure 4N), and the yield of the fad2‐1a/b plants was marginally higher than that of XD18 and fad2‐1a/b/lox1/2/3 plants (Figure 4O).

Figure 4.

Figure 4

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Phenotypic analysis of the fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants in Shandong in 2024

(A–E) Morphology of the XD18, fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants at the flowering stage (A, B) and mature stage (C–E). (F–O) Quantification of the flowering time (F), time to maturation (G), plant height (H), branch number (I), node number of main stem (J), effective pods per plant (K), number of seeds per plant (L), weight of seeds per plant (M), 100‐seed weight (N), and yield per 666.67 m2 (O) of the XD18, fad2‐1a/b, and fad2‐1a/b/lox1/2/3 plants. Bars = 5 cm. Data are means ± SD. n = 10 in (F–H), n = 20 in (I–M); n = 3 in (N, O). The significant difference was calculated using one‐way ANOVA with the Tukey multiple comparison test. Different lowercase letters on top of the columns indicate significant differences (P < 0.05), and the same letters indicate no significant differences.

In 2024, the XD18, fad2‐1a/b, and fad2‐1a/b/lox1/2/3 plants performed slightly better in Anhui Province as they produced a greater number of effective pods per plant than in Shandong (4, 5). At the maturation stage, plants of all tested genotypes grew to nearly the same height and produced a similar number of nodes on the main stem, except for ZH301, which was the tallest genotype and had the largest number of nodes (Figure 5B, C). Among all genotypes, the fad2‐1a/b/lox1/2/3 #1 plants produced the greatest number of branches (Figure 5D). All plants had more than 100 effective pods per plant (Figure 5E). It seemed that the fad2‐1a/1b and WD37 plants had slightly larger seeds than the other genotypes (Figure 5F). The fad2‐1a/b/lox1/2/3 #1 plants produced the greatest number of seeds per plant, and the fad2‐1a/b and WD37 plants had a higher 100‐seed weight (Figure 5G, H). The fad2‐1a/b and fad2‐1a/b/lox1/2/3 #1 plants had a greater weight of seeds per plant than the other genotypes (Figure 5I). The yields of all genotypes were approximately the same, except that the yield of ZH301 was the highest (Figure 5J).

Figure 5.

Figure 5

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Phenotypic analysis of the fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants in Anhui in 2024

(A) Morphology of the XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3, ZH301, and WD37 plants at the mature stage. (B–J) Quantification of the plant height (B), node number of main stem (C), branch number (D), effective pods per plant (E), seed area (F), number of seeds per plant (G), 100‐seed weight (H), weight of seeds per plant (I), and yield per plot area (J) of the XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3, ZH301, and WD37 plants. Bars = 10 cm. Data are means ± SD. n = 40 in (B–I); n = 3 in (J). The significant difference was determined using one‐way ANOVA with the Tukey multiple comparison test. Different letters on top of the columns indicate significant differences (P < 0.05), and the same letters indicate no significant difference.

DISCUSSION

The fatty acid composition of soybean oil is crucial for its quality and nutritional value. By leveraging the precision of gene editing technology, we targeted the two Δ‐12 fatty acid desaturase 2 genes (GmFAD2‐1A and GmFAD2‐1B) and developed soybean plants with a significantly elevated oleic acid content, which accounts for over 80% of the total fatty acids in the seeds. We subsequently obtained soybean plants in which the two Δ‐12 fatty acid desaturase 2 genes (GmFAD2‐1A and GmFAD2‐1B) and three lipoxygenase genes (GmLOX1, GmLOX2, and GmLOX3) were edited. The objective was to produce soybean plants with high oleic acid content and seeds devoid of grassy and beany flavors. Our results demonstrated that the seeds of two independent fad2‐1a/b/lox1/2/3 #1 and fad2‐1a/b/lox1/2/3 #2 plants contained up to 80% oleic acid relative to the total fatty acids, and there was a significant reduction or absence of products associated with grassy and beany flavors. Field tests indicated that the fad2‐1a/b and fad2‐1a/b/lox1/2/3 plants exhibited normal growth performance comparable to the XD18 plants. The high oleic acid soybean oil produced from these genetically edited soybeans is expected to have better oxidative stability and a longer shelf‐life compared to traditional soybean oils. This is because oleic acid is a monounsaturated fatty acid that is less prone to oxidation than polyunsaturated fatty acids.

Collectively, these results revealed that our genome editing strategy, which synergistically targeted both the two FAD2‐1 and three LOX genes, generated ideal soybean varieties. The seeds of these varieties can serve as excellent materials for the production of healthy soybean oil and other grassy‐ and beany‐flavor‐free food products, such as soymilk and yogurt, for consumers, particularly those who previously disliked soybean products due to the grassy and beany flavors. This could lead to an increase in the market share of soybean‐based food products and contribute to a more sustainable food system by reducing the reliance on animal‐based dairy products.

Previous studies, through the selection of natural or induced mutations in FAD2‐1 genes or genome editing targeting these two genes, have shown that certain germplasms have the potential to accumulate high levels of oleic acid (Bachlava et al., 2008; Scherder and Fehr, 2008; Haun et al., 2014; Do et al., 2019). Additionally, reports on lipoxygenase‐free soybean varieties are available (Lee et al., 2014; Wang et al., 2020). However, there is no report on existing soybean varieties with seeds that contain high oleic acid and are free of grassy and beany flavors. One possible explanation for the scarcity of high oleic acid soybean oil or lipoxygenase‐free soybean food products is the severe growth impairment of soybean plants with mutations in the FAD2‐1 or LOX genes. Our study not only enhanced the oleic acid levels in seeds but also significantly reduced the substrate levels for LOX enzymes. Therefore, a combinatorial design for the dysfunction of two Δ‐12 fatty acid desaturases and three lipoxygenases is a practical and effective approach to achieving the goal of creating a soybean variety with high oleic acid content and seeds free of grassy and beany flavors.

Our work can provide a framework for the genetic improvement of other soybean germplasms to enhance their nutritional value. By targeting similar genes involved in fatty acid metabolism and flavor formation, it may be possible to develop other oil‐rich crops with enhanced nutritional value and improved sensory qualities. This would have a significant impact on global food security and human health, as oils are an important source of energy and essential fatty acids in the human diet. Overall, the results of this study represent a major breakthrough in soybean genetic engineering and have the potential to revolutionize the soybean industry. The development of high oleic acid soybean varieties with seeds free of grassy and beany flavors not only meets the growing demand for healthy and flavorful food products but also provides a model for the genetic improvement of other crops.

MATERIALS AND METHODS

Plant materials and growth conditions

In this study, the soybean cultivar Xudou 18 (XD18) was employed. Soybean plants were cultivated in fields in Sanya (Hainan Province, China) from January to May 2022, as well as in Fuyang (Anhui Province, China) and Jinan (Shandong Province, China) during the normal soybean‐growing seasons (typically from June to October) in 2024. When required, the soybean plants were also grown in a growth chamber under a photoperiod of 12 h of light (approximately 800 μmol·m−2·s−1) followed by 12 h of darkness at a temperature of 25°C.

Vector construction and soybean genetic transformation

The crRNAs were designed through the online software CRISPR‐GE (Xie et al., 2017). The CRISPR‐Cas‐SF01 binary vector contains the expression cassettes as illustrated in Figure 1. The binary vector was introduced into the Agrobacterium tumefaciens (strain EHA105). The soybean fad2‐1a/b and fad2‐1a/b/lox1/2/3 mutants were generated by the Agrobacterium tumefaciens‐mediated cotyledon node transformation (Paz et al., 2006; Jing et al., 2026), and homozygous fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 mutants were identified by genotyping PCR reactions and subsequent sequencing of the relevant PCR products.

Determination of oil and protein contents in soybean seeds

Measurements of oil and protein contents were conducted at the Crop Institute, Anhui Academy of Agricultural Sciences (Hefei, Anhui, China). Uniform dry seeds of XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 were collected from Fuyang (Anhui Province, China) and assayed using the Near‐infrared Grain Analyzer, DA7250 (Perten, PerkinElmer, Shelton, CT, USA) to determine the oil and protein contents in accordance with the standard procedures described by the manufacturer. During spectral acquisition, the uniform dry seeds were uniformly distributed in the sample container to ensure the absence of spaces, which could lead to interference from objective factors such as spectral scattering.

To determine the composition of oil components, we analyzed the free fatty acids and related metabolites in uniform dry seeds of XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 following standard procedures at MetwareBio (Wuhan, China). The prepared samples were analyzed using a gas chromatography (Agilent 8890) coupled to triple quadruple mass spectrometry (Agilent 7000E) (GC‐MS). The GC analytical conditions were as follows, the separation was conducted on a DB‐5MS capillary column (30 m × 0.25 mm × 0.25 μm, Agilent) with high purity helium (purity > 99.999%) as carrier gas. There were five biological replicates per genotype.

Assays of GmLOX enzymatic activities

A colorimetric assay for the determination of lipoxygenase activity was performed as previously described with minor modifications (Suda et al., 1995; Narvel et al., 2000). Dry seed samples were ground into fine powder and stored at −80°C freezer prior to use. To detect LOX1 activity, 1.5 mL of 0.2 M sodium borate buffer (pH 9.0) was added to 15 mg ground sample, mixed, and incubated for 5 min on a rotating shaker. The homogenized mixture was centrifuged at 12,000 g for 5 min at 4°C, and the supernatant was collected in a 2‐mL microcentrifuge tube. Subsequently, 1.0 mL reaction buffer (0.125 M sodium borate, 12.5 μM methylene blue, 1.375 mM sodium linoleate (the substrate), pH 9.0) was added to 0.5 mL of the supernatant, mixed, and incubated for 15 min. The color of the incubated sample/reaction buffer mixture was recorded, and absorbance at 660 nm was measured with the modular multimode microplate reader, BioTek Synergy H1 (Agilent, Santa Clara, CA, USA).

To detect LOX2 activity, 1.5 mL of 0.2 M sodium phosphate buffer (pH 6.0) was added to 15 mg ground sample, mixed, and incubated for 5 min on a rotating shaker. The homogenized mixture was centrifuged at 12,000 g for 5 min at 4°C, and the supernatant was collected in a 2‐mL microcentrifuge tube. Subsequently, 1.0 mL reaction buffer (0.125 M sodium phosphate, 12.5 μM methylene blue, 1.375 mM sodium linoleate (the substrate), 25 mM DL‐dithiothreitol (DTT), 12.5% acetone, pH 6.0) was added to 0.5 mL of the supernatant, mixed, and incubated for 15 min. The color of the incubated sample/reaction buffer mixture was recorded, and absorbance at 660 nm was measured with the modular multimode microplate reader, BioTek Synergy H1 (Agilent, Santa Clara, CA, USA).

To detect LOX3 activity, 1.5 mL of 0.2 M sodium phosphate buffer (pH 6.6) was added to 15 mg ground sample, mixed, and incubated for 20 min on a rotating shaker. The homogenized mixture was centrifuged at 12,000 g for 10 min at 4°C, and the supernatant was collected in a 2‐mL microcentrifuge tube. Subsequently, 1.0 mL reaction buffer (0.125 M sodium phosphate, 1.375 mM sodium linoleate (the substrate), 12.5% β‐carotene at 50% saturation, pH 6.6) was added to 0.5 mL of the supernatant, mixed, and incubated for 15 min. The color of the incubated sample/reaction buffer mixture was recorded, and absorbance at 452 nm was measured with the modular multimode microplate reader, BioTek Synergy H1 (Agilent, Santa Clara, CA, USA).

Determination of grassy and beany flavors in soybean seeds

Determination of the grassy and beany flavors in soybean seeds was conducted as described (Chigwedere et al., 2019). The dry seed samples of XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 were ground into fine powder and stored at −80°C freezer prior to use. One‐fifth grams of the ground sample was transferred to a 20 mL amber glass vial (Agilent, Palo Alto, CA, USA), containing 0.2 g NaCl to inhibit any enzyme reaction. The vials were sealed using crimp‐top caps with TFE‐silicone headspace septa (Agilent, Palo Alto, CA, USA). At the time of solid phase microextraction (SPME) analysis, each vial was placed in 60°C for 5 min, then an SPME Arrow (Agilent, Palo Alto, CA, USA) of 120 μm divinylbenzene/carboxen/polydimethylsiloxane (DVB/CAR/PDMS) was exposed to the headspace of the vial containing sample to extract a wide range of volatile compounds (VOCs) for 15 min at 60°C. After sampling, desorption of the VOCs from the SPME Arrow coating was carried out in the injection port of the GC apparatus (Model 8890; Agilent) at 250°C for 5 min. The identification and quantification of VOCs was carried out using an Agilent Model 8890 GC and a 7000D mass spectrometer (Agilent, Palo Alto, CA, USA), equipped with a 30 m × 0.25 mm × 0.25 μm DB‐5MS (5% phenyl‐polymethylsiloxane) capillary column. Helium was used as the carrier gas at a linear velocity of 1.5 mL/min. The injector temperature was kept at 250°C. The GC oven temperature was maintained at 40°C for 3.5 min before the temperature was ramped up to 100°C at 10°C/min and maintained for 1 min after which it was ramped again to 220°C at 6°C/min. After a holding of 2 min, a final ramp at 50°C/min to 250°C was made, and immediately, the oven was cooled back to 40°C. For the MS, the temperatures of the quadruple, which was used at 70 eV, and the ion source were 150°C and 230°C, respectively, while a mass‐to‐charge ratio scanning range of 35–550 at 3.8 scans/s was applied. A standard, being a mixture of n‐alkanes (C8–C22), was analyzed in order to obtain a retention index standard (calibration data). Specifically, 10 μL of the alkane standard was mixed with 5 mL of demineralized water in a 20 mL vial and analyzed as described above.

Measurements of agronomic traits in field‐cultivated plants and plot‐based field experiments

The plant height, the number of nodes on the main stem, the number of branches, and the number of effective pods per plant were manually documented. The seed surface area, the number of seeds per plant, and the weight of seeds per plant were measured using the intelligent grain analysis system, TPKZ‐3 (Top Cloud Agricultural Technology Company Ltd., Hangzhou, China). Fully filled grains were employed to measure the 100‐seed weight with this grain analysis system.

In 2024, the XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants were cultivated in the fields of Sanya under natural conditions. Each plot had an area of 30 m2, and four replicate plots of equal size were established for each genotype. The length of each plot was 10 m, with a row spacing of 40 cm and a plant spacing of 12 cm. Three weeks after sowing, manual thinning of seedlings was conducted to attain a uniform density of 14,000 individual plants per 666.67 m2.

In 2024, the XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants were cultivated in the fields of Jinan under natural conditions. Each plot covered an area of 15 m2, with three replications of equal‐sized plots. Six rows were planted in each plot, and the yield was measured using plants collected from the middle four rows. The length of each plot was 6 m, and the row spacing and plant spacing were 40 cm and 12 cm, respectively. Three weeks after sowing, the seedlings were manually thinned to attain a uniform density of 14,000 individual plants per 666.67 m2. The seeds were sown on June 20, 2024, and the grain harvest was completed between October 10 and 18, 2024.

In 2024, plants of XD18, fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2, along with two regional reference cultivars, Wandou 37 (WD37) and Zhonghuang 301 (ZH301), were cultivated in the fields of Fuyang under natural conditions. Each plot had an area of 20 m2, and for each genotype, there were four plot replications of equal size. The length of each plot was 5 m, with a row spacing of 40 cm and a plant spacing of 12 cm. Three weeks after sowing, the seedlings were manually thinned to attain a uniform density of 14,000 individual plants per 666.67 m2. The seeds were sown on June 20, 2024, and the grain harvest was completed between October 5 and 12, 2024.

CONFLICTS OF INTEREST

The authors declare no conflicts of interest.

AUTHOR CONTRIBUTIONS

H.X. and J.Z. contributed to the conceptualization of the project. H.X., L.G., Z.H., F.C., M.T., and Z.J. performed the research. H.X., L.G., D.W., Z.H., W.Y., P.C., and J.Z. analyzed the data. L.G., H.X., and J.Z. wrote the manuscript. All authors read and approved the contents of this paper.

Supporting information

Additional Supporting Information may be found online in the supporting information tab for this article: http://onlinelibrary.wiley.com/doi/10.1111/jipb.70282/suppinfo

Figure S1. Schematic illustration of the synthesis of certain unsaturated fatty acids and partial pathways for the oxidation of linoleic acid and α‐linolenic acid in soybean seeds

Figure S2. Sanger sequencing data of the fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants

Figure S3. Phenotypic analysis of the XD18 and fad2‐1a/b/lox1/2/3 plants in Hainan in 2024

JIPB-68-3260-s001.docx (1MB, docx)

ACKNOWLEDGEMENTS

This work was supported by funds provided by the Key R&D Program of Shandong Province, China (grant number 2023SFGC0301), the National Natural Science Foundation of China (grant number 32572436), the Agriculture Science and Technology Major Project of China, the Program on Industrial Technology System of National Soybean (CARS‐04), and by Bellagen Biotechnology Co. Ltd.

Biographies

graphic file with name JIPB-68-3260-g001.gif

graphic file with name JIPB-68-3260-g006.gif

Xie, H. , Geng, L. , Hu, Z. , Chen, F. , Tan, M. , Wang, D. , Huang, Z. , Ye, W. , Chen, P. , and Zhu, J. (2026). Genome editing generates high oleic soybean and eliminates beany flavors. J. Integr. Plant Biol. 68: 3260–3271.

Edited by: Lanqin Xia, Institute of Crop Sciences, CAAS, China.

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Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

Additional Supporting Information may be found online in the supporting information tab for this article: http://onlinelibrary.wiley.com/doi/10.1111/jipb.70282/suppinfo

Figure S1. Schematic illustration of the synthesis of certain unsaturated fatty acids and partial pathways for the oxidation of linoleic acid and α‐linolenic acid in soybean seeds

Figure S2. Sanger sequencing data of the fad2‐1a/b, fad2‐1a/b/lox1/2/3 #1, and fad2‐1a/b/lox1/2/3 #2 plants

Figure S3. Phenotypic analysis of the XD18 and fad2‐1a/b/lox1/2/3 plants in Hainan in 2024

JIPB-68-3260-s001.docx (1MB, docx)

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