Comparison of IAA and amino acid profiles of the selected rootstocks and their accumulation in grafted scion of Cucumis sativus L.

Plant material

A total of 57 genotypes from the Cucurbitaceae family were utilized in this study, comprising 38 genotypes of C. maxima, 12 genotypes of C. pepo, five genotypes of C. moschata, and two genotypes of C. ficifolia. The plant material was sourced from the Cucurbitaceae collection maintained by the Kazakh Research Institute of Fruit and Vegetable Growing. This germplasm collection has been systematically developed and expanded over the past two decades through the acquisition of diverse genotypes from international gene banks, including contributions from the World Vegetable Center (https://genebank.worldveg.org) and the N.I. Vavilov All-Russian Institute of Plant Genetic Resources (https://www.vir.nw.ru).

Each genotype was assigned a distinct catalog number to verify its origin and identity. Cucurbit cultivation was carried out in 300 ml containers filled with autoclaved perlite and equipped with expanded clay for drainage. Seeds were sown directly into the perlite substrate and irrigated daily with 100 ml of a mineral nutrient solution per plant. For morphometric assessment, plants were sampled 10 days after emergence. Growth parameters—including shoot and root length, as well as biomass—were quantified following the protocol described by Manda, Prasad & Palmei (2018). Post-germination, seedlings were exposed to LED lighting at an intensity of 5,000 lux for 10 days to facilitate the selection of one representative genotype per species, followed by 10,000 lux for the subsequent six weeks. Morphometric traits measured included shoot and root biomass separately. The composition of mineral salts and trace elements used in the nutrient solution is detailed in Table 1, based on General Hydroponics formulations (https://generalhydroponics.com). For biochemical analyses, seedlings at 2 and 4 weeks of age were selected.

During the initial two weeks of cultivation, the nutrient solution was maintained at a concentration of 500 ppm. This concentration was increased to 1,000 ppm during weeks three and four, and further elevated to 1,500 ppm in weeks five and six to support ongoing vegetative growth. To achieve these target concentrations, 1.3 ml, 2.7 ml, and 4.0 ml of each stock solution were added per liter of water, respectively (Table 1). The pH of the nutrient solution was adjusted to 6.0 using a 1M of KOH solution, and the total dissolved solids were monitored using a TDS meter. Neutralized peat (pH 6.0, Kekkila™, Finland) was employed as a substrate to sustain vegetative development in both grafted and control plants. For the determination of thousand-seed weight, shoot biomass, and root biomass, all biochemical analyses were performed using four biological replicates. For biochemical assays, all fully expanded leaves and corresponding stem segments were harvested from each plant. Leaf and stem tissues were pooled separately by cultivar to generate composite samples for analysis.

HPLC analysis of amino acids

The total amino acid content in plant extracts was determined using high-performance liquid chromatography (HPLC) on a Shimadzu Prominence LC-20 system (Shimadzu, Japan), following acid hydrolysis of the samples. Prior to analysis, amino acids were derivatized with phenyl isothiocyanate (PITC) according to the protocol described below. The HPLC setup included a binary pump (LC-20AD), autosampler (SIL-20AC), degasser (DGU-20A5), and column oven (CTO-20A), all controlled via LCSolution software. Chromatographic separation of amino acid derivatives and standards was carried out using a Thermo Hypersil GOLD C18 column (150 mm × 4.0 mm, 5 µm). Detection was performed by monitoring ultraviolet absorbance at 254 nm. The mobile phase was delivered at a flow rate of 0.8 mL/min, with each sample or standard requiring a total run time of 43 min.

Standard amino acids—L-aspartic acid (Asp), L-glutamic acid (Glu), L-serine (Ser), L-asparagine (Asn), L-histidine (His), L-arginine (Arg), L-threonine (Thr), L-alanine (Ala), L-proline (Pro), L-cysteine (Cys), L-tyrosine (Tyr), L-valine (Val), L-methionine (Met), L-cystine (Cys-Cys), L-isoleucine (Ile), L-leucine (Leu), L-phenylalanine (Phe), and L-lysine (Lys)—were obtained from Titan Biotech Ltd (Delhi, India). Calibration curves were generated using five-point serial dilutions of each standard at concentrations of 1, 10, 25, 50, and 100 µg/mL (Zheng et al., 2017; Hao et al., 2021).

HPLC analysis of IAA

Samples and standards were separated using a Restek Ultra C18 high-performance liquid chromatography (HPLC) column (150 mm × 4 mm, 5 µm; Bellefonte, PA, USA) maintained at 40 °C. Standard solutions were prepared according to the protocols outlined by Campanella et al. (2016) and Keskin et al. (2022). A stock solution of IAA was prepared by dissolving 10 mg of IAA in one mL of 1M NaOH, followed by dilution with nine ml of deionized water in a 10 ml test tube. For sample preparation, one g of plant tissue was weighed and homogenized. Each homogenate was transferred to a 10 mL centrifuge tube and mixed with 10 mL of an acetonitrile:deionized water solution (9:1, v/v) under low-light conditions to prevent photodegradation. Subsequently, 100 µL of formic acid was added, and the mixture was thoroughly shaken. The homogenates were incubated for 2 h and then centrifuged at 12,000× g for 20 min at 4 °C. The resulting supernatants were collected and concentrated using a vacuum evaporator (Biobase, China). The dried residues were reconstituted in two mL of acetonitrile and purified with a C18 solid-phase extraction (SPE) cartridge (200 mg/3 mL; Strata, Phenomenex, Torrance, CA, USA). The SPE cartridge was conditioned by sequential elution with two mL water and two mL ethanol through a vacuum manifold (Biobase, Shandong, China) set to a negative pressure of −0.01 MPa. The flow-through was discarded, and IAA was eluted with two mL of an ethanol–water–formic acid mixture (80:20:0.5%, v/v) into a 10 mL centrifuge tube. The eluates were transferred to two mL Eppendorf tubes, concentrated using a sample concentrator (NDK200-2N, Miulab, Hangzhou, China), and reconstituted in two mL of acetonitrile. Aliquots of 20 µL were injected into the HPLC system. The peaks of pure standard substances, retention times, linear calibration parameters, limit of detection, limit of quantification of amino acids and IAA are provided in Figs. S1 and S2. Also, chromatographic peaks of amino acids and IAA quantification of Cucurbitaceae family are shown in Fig. S3.

Grafting methods

Tongue approach grafting was employed as the grafting technique in this study. Seeds of the scion were sown seven days prior to the rootstock material to ensure appropriate developmental synchronization (Lee et al., 2010). The cucumber hybrid cultivar Gentle F1 (Syngenta, Basel, Switzerland) was used as the scion. Grafting was performed by making longitudinal incisions at an angle of 20–30° on the stems of both the rootstock and scion. The incision on the rootstock was oriented downward, while that on the scion was directed upward. The two components were then joined at the cut surfaces (“tongues”) and secured using grafting clips. Initially, both root systems were retained to support early establishment. Ten days post-grafting, the aerial portion of the rootstock and the root system of the scion were excised to complete the graft union. Following grafting, the plants were transferred to a climate-controlled growth chamber with regulated temperature, humidity, and photoperiod conditions. For the first four days, grafted plants were maintained in complete darkness at 20 °C and 92% relative humidity to promote callus formation and vascular reconnection. Subsequently, plants were exposed to a light intensity of 12,000 lux under an 8/16-hour light/dark cycle for ten days to support acclimatization and growth (Lee et al., 2010; Rymbai et al., 2015). Self-grafted C. sativus plants served as the control.

Cultivation of grafted plants

The grafted C. sativus plants were grown in two greenhouses: (1) Experimental stationary greenhouse of the Kazakh National Agrarian Research University (43°14′31″N 76°57′07″E, Almaty, Kazakhstan). (2) Industrial greenhouse of the Kazakh Scientific Research Institute of Fruit and Vegetable Growing Kainar with controlled climate (43°09′22.4″N 76°26′45.5″E, Kaskelen township, Kazakhstan).

Statistical analyzes

Statistical analysis was performed using the one-way anova-test to assess differences within groups. A p-value of <0.05 was considered statistically significant. *p < 0.05, **p < 0.01, ***p < 0.001, ****p < 0.0001. Further, GraphPad Prism 9.0 (GraphPad Software, San Diego, CA, USA) was used to prepare graphs and design statistical analysis of experimental data. Statistical analyses using JMP PRO 17 (JMP Statistical Discovery LLC) were applied for analyses related to discriminant, cluster (for morphometric characteristics of the Cucurbitaceae family) and component (amino acid composition of the Cucurbitaceae family).

Amino acids and IAA content in leaves and stems of Cucurbitaceae plants

Leaf and stem tissues from the four selected cultivars were harvested and pooled separately to ensure representative sampling. Composite extracts were prepared from the mixed leaf and stem material, and aliquots were collected for biochemical analysis at two and four weeks post-germination. Solid-phase extraction was employed to purify the samples prior to analysis. Following extraction, amino acids and IAA were identified based on their retention times and peak profiles in chromatographic output (Fig. S3). Quantitative data were subsequently processed using Excel and normalized to tissue mass, with final concentrations expressed as µg per gram of leaf or stem tissue.

The two-week stems of C. ficifolia contained the highest levels of arginine (142.3 ± 3.1 µg/g) and threonine (110.7 ± 2.9 µg/g), while its leaves were richest in isoleucine (98.5 ± 2.6 µg/g) (Table 3). At 4 weeks, its stems accumulated the highest alanine content (125.2 ± 3.3 µg/g), and the leaves retained the highest arginine concentration (160.8 ± 4.2 µg/g), although these values were generally lower than those of C. maxima at the same age. The two-week stems of C. moschata showed the highest levels of aspartic acid (211.6 ± 4.4 µg/g), while the leaves exhibited elevated serine (112.2 ± 3.1 µg/g), threonine (109.4 ± 2.5 µg/g), and cysteine (86.1 ± 2.2 µg/g). At 4 weeks, its stems remained high in aspartic acid (222.3 ± 4.1 µg/g, p < 0.05) and leucine (131.4 ± µg/g), and the leaves showed the greatest accumulation of glutamic acid (198.7 ± 3.9 µg/g), proline, tyrosine, and leucine. The two-week stems of C. pepo had the highest content of glutamic acid (224.1 ± 5.0 µg/g), serine, asparagine, histidine, alanine, cysteine, methionine, and cystine. Its leaves at the same stage had the highest alanine (120.5 ± 2.7 µg/g) and methionine (89.3 ± 2.1 µg/g). Four-week-old C. pepo stems were found to have the highest content of glutamic acid, arginine, cysteine, and cystine, while leaves had the highest content of alanine and cysteine. Two-week-old C. maxima stems consistently demonstrated the highest overall amino acid accumulation. At two weeks, its stems had the highest concentrations of proline (148.9 ± 3.8 µg/g), tyrosine (117.2 ± 2.9 µg/g), valine, isoleucine, leucine, and lysine, while the leaves showed elevated levels of aspartic acid (231.4 ± 6.8 µg/g), glutamic acid (212.9 ± 4.3 µg/g), asparagine, histidine, arginine, tyrosine, valine, cystine, and lysine. The highest concentrations of serine (145.2 ± 3.2 µg/g), asparagine (142.7 ± 4.5 µg/g), histidine, threonine, proline, tyrosine, valine, methionine, isoleucine, and phenylalanine were detected in the four-week-old stems of C. maxima. In the leaves, elevated levels of aspartic acid, serine, asparagine, histidine, valine, methionine, cystine, isoleucine, phenylalanine, and lysine were observed. Statistical analysis confirmed that C. maxima at 4 weeks had significantly higher total amino acid content than all other species (1,810 ± 35 µg/g in leaves; p < 0.01), particularly in aspartic acid, glutamic acid, serine, asparagine, and histidine (all p < 0.01 vs. other species). The highest total amino acid content was observed in C. maxima at 4 weeks of age, with aspartic acid, glutamic acid, serine, asparagine, and histidine showing the greatest accumulation. These findings highlight C. maxima as the most suitable rootstock candidate based on its superior biochemical profile.

The results on the amino acid content were then used to conduct a principal component analysis (PCA). Figure 3 shows the distribution of traits in each of the first two components, PC1 and PC2, which explained 73.4% and 9.8% of the variability, respectively, with total variability 83.2%. PCA plot shows Cucurbitaceae samples with different genotypes described by organ (leaf and stem) and time points (2 and 4 weeks). It is evident that each variation was subdivided into a separate group that could be distinguished from the others.

The majority of amino acid components were detected in leaf samples collected at four weeks. Alanine showed a strong contribution to PC 2, whereas most other amino acids—such as aspartic acid, lysine, and glutamic acid—clustered along PC1. Variation in amino acid composition was observed among C. pepo, C. moschata, and C. maxima, primarily reflecting differences in leaf profiles over the four-week period. In contrast, C. ficifolia exhibited variation in stem samples at two weeks, with relatively lower diversity in amino acid composition. Amino acids including glutamic acid, lysine, methionine, valine, histidine, serine, leucine, proline, and asparagine formed a tight cluster, indicating coordinated variation across samples—i.e., elevated levels of one are likely associated with increased levels of the others. Alanine, positioned distinctly along PC2, displayed a unique variation pattern, suggesting it behaves independently from the clustered group. Notably, C. moschata appeared slightly separated from the other species in the PCA plot, implying potential differences in its amino acid profile. Overall, the PCA visualization highlights species-specific biochemical variation within the Cucurbitaceae family.

According to the accumulation of IAA, it is evident from Fig. 4 that the highest values were in the C. maxima and C. pepo samples at the age of 2 and 4 weeks, mainly in the leaves. In addition, at the age of two weeks, the IAA content was not detected in the stems of the C. ficifolia and C. moschata samples. The p-value for 2-week-old C. ficifolia plants is 0.030, and for C. moschata it is 0.032. By the end of the fourth week, the presence of IAA was detected in the stems, with the highest content of 0.1 µg/g observed in C. maxima, and 0.4 ± 0.02 µg/g in the leaves, respectively. C. maxima exhibited the highest IAA level, which was significantly higher than in C. ficifolia (0.18 ± 0.01 µg/g, p = 0.002) and C. moschata (0.21 ± 0.01 µg/g, p = 0.004). The highest IAA content in the leaves was found in both two-week-old (Fig. 4B) and four-week-old leaves (Fig. 4B) of C. maxima plants. Based on the results of morphometric and chromatographic analyses of amino acids and IAA, C. maxima was chosen as a rootstock for further work.

Amino acids, IAA content, and yield performance of grafted C. sativus plants

One-week-old C. sativus seedlings were grafted onto C. maxima rootstocks. The resulting grafted plants are depicted in Fig. 5A. The ligulate grafts rapidly established contact with the supporting rootstock stems, achieving complete healing within one week. Graft union and healing were evaluated visually, and tactile inspection confirmed firm junctions between the stem segments. No wilting symptoms were observed after transplanting the grafted plants to their final greenhouse locations. To maintain experimental consistency, C. sativus plants were also grafted onto themselves as controls. Throughout the experiment, no preferential conditions were applied to either the control group or the C. maxima grafts. Post-grafting, all plants were maintained for four days in a growth chamber under complete darkness at 20 °C and 92% relative humidity. Subsequently, they were transferred to conditions of 12,000 lux illumination with an 8/16-hour light/dark cycle for an additional three days.

Starting from the second week post-grafting, the grafted plants were relocated to a climate-controlled greenhouse. Noticeable plant growth commenced two weeks after grafting, and by the end of the third week, differences in growth rate and development among the grafted plants became apparent. Figure 5B presents the growth outcomes of four-week-old plants. Throughout the grafting period –up to six weeks post-procedure –visual assessments indicated that the grafted plants kept pace with both the non-grafted control plants and the C. sativus specimens grafted onto themselves (Fig. 5C). Under identical environmental conditions, grafted seedlings were also cultivated in a second industrial greenhouse, as shown in Figs. 5D–5F.

At four weeks of age, morphometric analysis was conducted, revealing that the C. sativus plants grafted onto C. maxima exhibited superior performance compared to other grafting samples. By six weeks post-grafting, the survival rate of plants grafted onto C. maxima reached 96%, while the self-grafted C. sativus control group showed a similarly high survival rate of 97%. Figure 6 presents the levels and accumulation patterns of amino acids and IAA in both grafted and control plants. In two-week-old specimens, grafted plants displayed elevated concentrations of lysine and tyrosine relative to the ungrafted controls (Fig. 6A), while no significant differences were observed for the remaining amino acids.

Four-week-old grafted plants exceeded the control in lysine, arginine, asparagine, aspartic acid, threonine, alanine, tyrosine, valine, cystine and isoleucine (Fig. 6B). It can be seen the significant increase of the L-valine, L-threonine, L-alanine, L-tyrosine by 28.8; 38.8; 42.5; 98%, respectively. Two-week-old grafted and control plants did not differ in IAA content (Fig. 6C). However, four-week-old grafted plants significantly exceeded the control values (Fig. 6C). Namely, IAA by 39% in the scions compared to the control cucumber plants were observed at the end of the fourth week after grafting.

Grafted plants outperformed the control group in terms of fruit count per plant (Fig. 7A) and fruit number per unit area (Fig. 7B). However, no significant differences were observed between grafted and control plants regarding fruit weight per plant (Fig. 7C) or per square meter (Fig. 7D), nor in individual fruit weight (Fig. 7E). Grafted plants also exhibited greater height compared to controls (Fig. 7F). The growth and fruiting period of grafted plants was, on average, extended by 10 days relative to the control group. By the end of the vegetative season, grafted cucumber plants demonstrated a 23% increase in yield per plant and per square meter compared to controls.

Additionally, for comparative purposes with the C. maxima rootstock, yield parameters of grafted C. sativus plants using C. pepo and C. ficifolia as rootstocks are presented in Figs. S4 and S5.