1. Introduction
The genus
Castanea belongs to the Fagaceae family and comprises seven species. The chestnut (
Castanea sp.) is native to the temperate regions of the northern hemisphere and is cultivated for its nuts, beauty, shelter, and wood and has been an important part of the human diet [
1]. The four most important species are the American chestnut (
Castanea dentata Borkh. and
Castanea pumila Mill.), Asian chestnuts (
Castanea mollissima Blume,
Castanea henryi Rehder & E.H. Wilson and
Castanea seguinii Dode), Japanese chestnut (
Castanea crenata) and European chestnut (
Castania sativa).
Castanea sativa is native to the Mediterranean countries of southern Europe and south-west Asia. The most important diseases of the tree are caused by some
Phytophthora species and by chestnut blight (
Cryphonectria parasitica M.E. Barr) [
2,
3]. Due to the massive decline in
Castanea sativa in Europe, interspecific hybrids with Asian species have been increasingly bred. Interspecific hybrids are tolerant to American canker and chestnut blight, diseases that were previously responsible for the decline of the species [
4,
5]. Recent studies also report better rooting of interspecific hybrids [
6,
7,
8,
9].
The chestnut is a woody species known for its difficult generative or vegetative propagation [
10]. Vegetative propagation by grafting is time-consuming or often fails due to incompatibility between scion and rootstock [
1]. Propagation by cuttings is the most important and most frequently used method of vegetative propagation in horticulture. In this method, a crucial stage is the formation of adventitious roots [
11]. The ability of plants to form adventitious roots on the basal part of stem cuttings varies greatly between different plant species and cultivars [
12]. Adventitious root formation is often limited by the poor rooting ability of
Castanea species [
1].
Adventitious roots form from non-root tissue and can be the result of normal development or in response to stresses such as nutrient deficiency, flooding, heavy metal exposure, and biotic or abiotic wounding. The basis of cutting propagation is wound-induced adventitious rooting [
13]. The formation of adventitious roots is a heritable quantitative genetic trait that is controlled by the interaction of several endogenous and environmental factors [
14]. Exogenous stimuli that affect this process are light, wounding, temperature and mineral supply. The formation of adventitious roots is also regulated by endogenous factors, of which the phytohormone auxin is the most important plant hormone in root development [
14].
Auxin metabolism genes regulate adventitious root formation by influencing auxin levels. Inhibition of polar auxin transport inhibits the early local increase in auxin, the induction of adventitious roots and the formation of sinks in the root regeneration zone [
15]. Auxin can act synergistically or antagonistically with other hormones and other growth regulators such as cytokinins, brassinosteroids, ethylene, abscisic acid, gibberellins, jasmonic acid, polyamines and strigolactones to trigger cascades of events leading to adventitious root initiation and development [
14,
16]. The effect of IAA in the plant regarding the induction of root development is associated with a very short period immediately after the start of propagation (after cutting from a stock plant) [
17]. Nag et al. [
17] discovered that the presence of auxin in the base of the cutting is significant within 24 h after the start of propagation. An important mechanism for influencing auxin signaling pathways is amplification/inhibition by cofactors/inhibitors. Cofactors of auxin action often include phenolic substances [
9]. It has been reported that substances, such as flavonoids, are associated with rooting [
18]. It has been suggested that phenolics influence rooting by inhibiting IAA decarboxylation, which protects IAA from oxidation [
19,
20]. Increased flavonoid content could increase IAA concentration by regulating auxin transport, which is why easy-to-root genotypes were associated with higher flavonoid and endogenous IAA concentrations than difficult-to-root genotypes [
12,
21]. Previous reports indicate that the concentrations of endogenous phenolic compounds do not change in the days following severance [
8,
9,
22].
Strigolactones, a group of phytohormones reported recently, have been characterized as inhibitors of adventitious root formation [
16]. Strigolactones are produced primarily in roots [
23]. Pacuar et al. [
14] reported an inhibitory effect on adventitious root formation in
Arabidopsis and
Pisum sativum, as adventitious rooting was enhanced in strigolactones-deficient and response mutants. Negative regulation of strigolactones in adventitious root formation has been reported in both herbs and woody plants [
24]. Inhibitory effects of strigolactones on the formation of adventitious roots were also reported by Fan et al. [
24] in apple stem cuttings. Ozbilen et al. [
25] reported on the effects of rac-GR24 as a strigolactone analog and TIS108 as a strigolactone inhibitor on easy and difficult-to-root olive cultivars, with the strigolactone inhibitor TIS108 increasing the rooting ability of cuttings of the difficult-to-root cultivar. The inhibitory effect of strigolactones on adventitious root formation appears to be indirect, mainly by inhibiting the positive role of auxin [
26]. The quantification of specific strigolactones in plants could help to understand the role of this newly labeled group of plant hormones in adventitious root formation.
Castanea sativa is a widespread tree species in Slovenia. It covers more than 230,000 ha, which is 22% of the total forested area in Slovenia [
3]. Hybrids are increasingly used because of their better rooting ability and their tolerance to common diseases, but also because of their interesting fruit for fruit growing. The conservation and propagation of Slovenian genotypes are essential for the preservation of biodiversity and the conservation of plant genetic resources in our area. Slovenian genotypes are adapted to our growing conditions; it is also important to preserve the natural and cultural heritage by preserving the genotypes native to the area.
To obtain a first indication of whether endogenous concentrations of auxin, flavonoids and strigolactones may play a role in the low rooting of native genotypes of Castanea, we compared the rooting rate of native genotype ‘Kozjak’ and hybrid clones ‘Marsol’ and ‘Maraval’. Furthermore, the concentrations of endogenous auxin, flavonoids as cofactors and strigolactones as inhibitors for adventitious roots were analyzed and compared. This result should indicate whether these endogenous substances may limit the rooting of Castanea.
3. Discussion
Many authors reported difficulties in the root formation of
Castanea sativa [
10,
27,
28] under in vivo and in vitro conditions. Our results also showed the difficulty in the rooting of
Castanea sativa (‘Kozjak’) with significantly lower rooting success and higher undesirable traits compared with the Japanese × European chestnut hybrids (
Castanea crenata ×
Castanea sativa) ’Marsol’ and ‘Maraval’. Callus development is usually the result of difficult rooting [
11], which we were also able to confirm with our results. Callus development was observed in all evaluated cuttings of the ‘Kozjak’ genotype, while callus development was significantly lower in ‘Marsol’ and ‘Maraval’. According to Kunc et al. [
11], acrobasal roots are more desirable in horticultural practice because the rooted cuttings are easier to dig up, and the newly formed roots are less damaged during digging and transplanting. The main root number is also a very good indicator of root quality [
7]. Our results are in agreement with those of Kunc et al. [
11], who report a lower proportion of acrobasal roots in cuttings obtained from mature stock material (difficult-to-root) of
Prunus subhirtella Miq. ‘Autumnalis’ and lower rooting results compared with cuttings obtained from rejuvenated stock material (easy-to-root). Eugenia Miranda-Fontaina [
29] also reported more undesirable characteristics of
Castanea sativa compared with hybrid clones. In contrast to our results, Osterc et al. [
8] reported differences in rooting and other evaluation parameters between the ‘Maraval’ and ‘Marsol’ genotypes. According to their results, the ‘Maraval’ genotype rooted better, the root system was longer, and the number of main roots was greater. Osterc et al. [
9] and Stefanic et al. [
30] also reported better rooting and less frequent callus formation in the ‘Maraval’ compared with the ‘Marsol’ genotype. In the results of Song et al. [
6], no differences in rooting success were found between ‘Marsol’ and ‘Maraval’, which is consistent with our results. High endogenous auxin concentration in the induction phase (0–24 h) of adventitious root formation is associated with a high rooting rate in
Sequoia sempervirens Endl. [
31] and
Prunus dulcis Batsch. [
32]. These results are consistent with our results, where the genotype ‘Maraval’ with the highest endogenous IAA concentration in the induction phase also had the highest rooting rate (see
Table 1 and
Table 2 and
Supplementary Material Table S3). Kunc et al. [
11] reported that the uptake of exogenously applied indole-3-butyric-acid (IBA) does not differ between easy and difficult-to-root plant material. With the same addition of IBA, the total auxin content increases equally for easy and difficult-to-root plant material. The present results highlight the importance of endogenous auxin for adventitious root induction in
Castanea.
Even though the strigolactones and flavonoids were measured at the end of the rooting period, their concentrations may indicate the functions of these compounds in the rooting process. Strigolactones are a special class of phytohormones that are categorized as repressors of adventitious root formation. However, the role of specific strigolactones is poorly understood [
33]. The concentration of three strigolactones (strigol, orobanchol and 5-deoxystrigol) in the roots and callus of three genotypes of
Castanea sp. (‘Kozjak’, ‘Marsol’, ‘Maraval’) and their correlation with initial endogenous IAA in the stem base were evaluated (see
Supplementary Material, Table S2). Previous studies [
33,
34,
35,
36] identified several genes of the strigolactone biosynthic pathway and showed that the inhibition of strigolactones promotes adventitious root formation. Our results may indicate that strigol and orobanchol in roots were negatively correlated with rooting parameters, while 5-deoxystrigol has no inhibitory effect on rooting parameters (see
Supplementary Material, Table S4). As can be seen from
Figure S1, 5-deoxystrigol is represented to a greater extent in the genotypes with higher rooting rates, ‘Marsol’ and ‘Maraval’. Our results may indicate that the group of strigolactones does not only have an inhibitory effect on adventitious root development. There are no significant differences in the total strigolactone concentration, although the rooting result is different between genotypes. This may be attributed to the fact that individual strigolactones have different effects on the rooting process. Sun et al. [
37] also reported that strigolactones can positively regulate the growth of adventitious roots. There is no evidence for the role of specific strigolactone in adventitious root formation. Given the lack of research on strigolactones, the identification of the specific strigolactones synthesized in plant roots is crucial. According to previous studies [
16,
17,
18,
19], we expected a higher concentration of strigolactones in the ‘Kozjak’ genotype, which showed the lowest rooting results and, consequently, many undesirable traits. The strigol concentration in the roots of ‘Kozjak’ and ‘Marsol’ was significantly higher than in ‘Maraval’. This result agrees with previous studies [
12,
13,
14] indicating better rooting of ‘Maraval’ compared with ‘Marsol’. This difference could also be explained by the ratio of strigolactones. The ratio of strigolactones in ‘Marsol’ and ‘Maraval’ is similar, with the strigol content being low and the 5-deoxystrigol content being high, which could indicate a positive effect of the latter on the formation of adventitious roots. Orobanchol in roots is negatively correlated with the development of adventitious roots, but there are no differences in concentration between genotypes. It can, therefore, be concluded that orobanchol does not play an important role as an inhibitor of adventitious root formation. A positive moderate correlation between IAA and orobanchol was observed (see
Supplementary Material, Table S2). Previous studies [
16,
37,
38,
39] reported a negative interaction between strigolactones and auxins in relation to root system formation, branching and other functions of strigolactones in the plant. Furthermore, the effects of strigolactone on root growth are likely linked to root responses not only to auxin but also to other plant hormones that influence root architecture. Understanding the role of individual strigolactones in the formation of adventitious roots is the key to the first step in achieving better rooting in difficult-to-root species.
The results of the flavonoid analyses showed differences in the concentration of flavonoid compounds not only between ‘Kozjak’ and ‘Maraval’ but also between ‘Marsol’ and ‘Maraval’. The correlation between the IAA concentration and the flavonoids was also analyzed (see
Supplementary Material, Table S1). Since there were no significant differences in the evaluation of rooting between ‘Marsol’ and ‘Maraval’, it can be concluded that it is not only the concentration of individual flavonoids that is decisive for adventitious root formation but also the total concentration and ratio. These results agree with recent studies [
12,
19,
40], which report that a higher concentration of total flavonoids also had a positive effect on rooting results. Higher concentrations of total flavonoids were observed in easy-to-root genotypes than in difficult-to-root genotypes. Osterc et al. [
8] also reported a highly significant correlation between quercetin concentrations and the rooting process. Zanoni do Prado et al. [
18] reported that substances such as flavonoids are associated with rooting. The concentration of quercetin-3-O-glucoside and quercetin-3-O-rhamnoside was determined in cuttings of
Eucalyptus gunnii Hook.f.. The two identified flavonoids were present in very low amounts in cuttings that could not root and in higher amounts in cuttings that could root, which is consistent with our results.
The significance of the proportion of the individual flavonoids in adventitious root formation has not been investigated yet. Since the ratio of specific flavonoids varied among the genotypes analyzed in our study, a correlation test was performed to investigate the relationship between the specific flavonoids and the rooting process. In ‘Kozjak’, ‘Marsol’ and ‘Maraval’, the main differences were observed in the distribution of isorhamnetin-3-O-glucoside, isorhamnetin-3-O-glucuronide and isorhamnetin-3-O-rutinoside. The percentage of isorhamnetin-3-O-glucoside in ‘Marsol’ was 53.00%, 33.00% in ‘Maraval’ and 17.00% in ‘Kozjak’. The percentage of isorhamnetin-3-O-glucuronide in ‘Marsol’ was 5.00%, 17.00% in ‘Maraval’ and 8.00% in ‘Kozjak’. The percentage of isorhamnetin-3-O-rutinoside was 8.00% in ‘Marsol’, 10.00% in ‘Maraval’ and 30.00% in ‘Kozjak’ (
Figure 1).
No significant differences in isorhamnetin-3-O rutinoside concentration were found between ‘Kozjak’ and ‘Maraval’, but the proportion in ‘Kozjak’ was higher (30.00%) than in ‘Marsol’ and ‘Maraval’ (8.00 and 10.00%). Since isorhamnetin-3-O-rutinoside was moderately and negatively correlated with the length of the root system, a significantly shorter length of the root system in the ‘Kozjak’ genotype could be explained by a higher proportion of isorhamnetin-3-O-rutinoside. The proportion and concentration of isorhamnetin-3-O-glucuronide were similar in ‘Kozjak’ and ‘Marsol’ but significantly higher in ‘Maraval’. Isorhamnetin-3-O-glucuronide correlated moderately with the number of main roots. The number of main roots also correlated with isorhamnetin-3-O-glucoside, but the correlation coefficient was high. The higher number of main roots in ‘Marsol’ can be explained by the high correlation between isorhamnetin-3-O-glucoside and the number of main roots, including a higher proportion of isorhamnetin-3-O-glucoside in ‘Marsol’ than in ‘Kozjak’.
Isorhamnetin-3-O-glucuronide and isorhamnetin-3-O-glucoside correlated highly with the length of newly formed shoots. The ratio of isorhamnetin-3-O-glucuronide in ‘Marsol was 5.00% and 8.00% in ‘Kozjak’, while the isorhamnetin-3-O-glucoside ratio was 53.00% in ‘Marsol’ and 17.00% in ‘Kozjak’, thus explaining the longer newly formed shoots in ‘Marsol’. The concentration of isorhamnetin-3-O-glucoside, which is associated with successful rooting, the number of main roots and the length of newly formed shoots, length of newly formed shoots and acrobasal rooting, is significantly different between all genotypes. A higher proportion of isorhamnetin-3-O-glucoside in ‘Maraval’ and ‘Marson’ is manifested in lower rooting success, the number of main roots and acrobasal root formation in ‘Kozjak’.
4. Material and Methods
4.1. Samples
Leafy cuttings of the two commercial cultivars ‘Marsol’ and ‘Maraval’ (Castanea crenata × Castanea sativa) and native accession ‘Kozjak’ (Castanea sativa) were collected in June from an Experimental Field for Nut Crops in Maribor (Slovenia; 46°34′01″ N; 15°37′51″ E; 280 m a.s.l.).
4.2. Experimental Design
The shoots of the current year of each genotype (n = 15) were used for the auxin analyses on the day the experiment was set up. The shoots were removed from the stock plant, and from the lowest parts, 3 cm of the cuttings were placed in liquid nitrogen, freeze-dried using the lyophilization method and stored for the analysis of endogenous IAA.
The propagation beds in which the cuttings were placed were divided into 15 plots (40 × 40 cm), with five replicates for each genotype and three biological replicates in each plot (
n = 15). After preparation, the leafy cuttings were planted in the plots for 120 days. The quality of the cuttings was evaluated in autumn, at the end of the growing season in October. The samples were washed so that no residues of substrate or IBA mixture were present. Root formation and quality of cuttings were assessed according to the protocol described by Kunc et al. [
11]. Callus, roots, and shoots were separated, placed in liquid nitrogen, freeze-dried using the lyophilization method, and stored at −20 °C for further analyses. In the next steps, the concentration of strigolactones (strigol, orobanchol, 5-deoxystrigol) in the callus and roots was analyzed. The identification and quantification of flavonoid compounds in the shoots of selected genotypes were also carried out. The experiment was set up as a single factorial design with five replicates. The distribution of replicates was based on simple random sampling. Each plot represented a single experimental replicate.
4.3. Preparation of the Cuttings
The shoots of the current year of each genotype were used as plant material for the experiment. The cuttings were shortened to a length of 15 cm, and the lower four leaves were removed. The base of the cuttings was treated with 0.0025 M IBA using the quick dip method (the basal end of 0.5 to 1 cm of the cutting is dipped in the solution for 3 to 5 s) and placed in the substrate. The substrate used was a mixture of peat and sand in a 1:1 ratio and was fertilized with 2.0 g/L 3–4 M Osmocote Exact fertilizer (16 (N)-9 (P)-12 (K) + 2(MgO) + trace elements) (ICL). The experiment was carried out in a greenhouse at the Biotechnical Faculty. The greenhouse was equipped with an automatically controlled fogging system that enabled a humidity of 98–100%. A high-pressure fogging system (Plantog, Fischament, Austria), which enables a high-quality rooting process, was used. The air temperatures in the greenhouse fluctuated greatly throughout the day, up to 50 °C during the day and between 18 and 20 °C at night. The system was in operation from 8 a.m. to 8 p.m. and was switched off at certain intervals. On sunny days, the interval was set to 40 s of fogging and a 1 min pause. On cloudy and rainy days, the intervals were extended to 3 min. The fogging system was always switched off at night. The experiment was conducted for 120 days, from June 2023 to October 2023.
4.4. Indole-3-Acetic Acid (IAA) Analyses
IAA was extracted according to the modified method of Trapp et al. [
41]. A total of 100 mg of dry samples (the lowest parts, 3 cm of the cuttings) were ground to a fine powder using a mortar and pestle in liquid nitrogen. Extraction was performed, adding 1.0 mL of isopropanol into each tube containing dry plant material. Samples were shaken (Unimax 1010 shaker, Heidolph, Schwabach, Germany) for 30 min centrifuged at 11,000×
g and 4° C for 5 min. The supernatant was transferred into a new 2 mL Eppendorf tube and dried in speed vac (Eppendorf). After drying, 200 µL of MeOH was added to each sample, homogenized and centrifuged at 11,000×
g and 4 °C for 10 min.
The supernatant was analyzed using a UHPLC system (Dionex UltiMate 3000; Thermo Scientific, Waltham, MA, USA) coupled with a tandem mass spectrometer (LTQ XL; Thermo Scientific, Waltham, MA, USA). The C18 column (150 × 4.60 mm; 3 µm column; Gemini; Phenomenex, Torrance, CA, USA) was used. The temperature was operated at 25 °C. The solvents used for the analysis were solvent A, consisting of 0.1% formic acid with 3% acetonitrile in double-distilled water (
v/
v/
v), and solvent B, consisting of 0.1% formic acid with 3% double-distilled water in acetonitrile (
v/
v/
v). The elution profile was as follows: 0–10 min, 42–55% B; 10–13 min, 55–100% B; 13–15 min 100% B; 15.1–20 min 42% B; and at 20–23 min, the reconditioning of the column was performed. The mobile phase flow rate was 0.6 mL min
−1. The injection volume was 20 µL. The identification of IAA was achieved by coupling the UHPLC system with a tandem mass spectrometer (LTQ XL; Thermo Scientific, Waltham, MA, USA) using heated electrospray ionization. The mass spectrometer was operated in negative ion mode. The following parameters were used: capillary temperature 200 °C; capillary voltage 8.0 V; sheath temperature 200 °C; sheath gas 30 arb; ion spray voltage 4.5 kV; tube lens −82 V; and auxiliary gas 10 arb. After determining the retention time and identifying typical fragmentation patterns of the obtained quantification standard, Selected Reaction Monitoring (SRM) was used to identify the selected compound. The transitions with the optimized parameters and retention times are shown in
Table 6.
4.5. Strigolactones Analyses
Strigolactones were extracted according to the modified method by López-Ráez et al. [
42] and Floková et al. [
43], in which 100 mg of dry tissue was ground in a mortar with liquid nitrogen. The samples were extracted for 60 min in a 10 mL glass vial with 1.5 mL of cold ethyl acetate. The samples were then shaken in ice-cold water (Unimax 1010 shaker, Heidolph, Schwabach, Germany). The samples were centrifuged at 2000×
g at 4 °C for 15 min. The organic phase was then carefully transferred to a 2 mL Eppendorf tube. The pellets were re-extracted with another 0.5 mL ethyl acetate; then, the combined ethyl acetate fractions were dried using the speed vac (Eppendorf). The dry residue was dissolved in 1 mL of 10% aqueous acetonitrile. The extracts were further pre-concentrated using reversed-phase polymeric Oasis
® HLB columns (150 mg/3 mL, Waters, Milford, MA, USA). The sorbent in the columns was activated with 3 mL of 100% acetonitrile and equilibrated with 10% aqueous acetonitrile. The samples of the tissue extracts were passed through the activated column. A further 3 mL of 10% aqueous acetonitrile was added. The strigolactone analytes were eluted from the column with 1 mL of 80 aqueous acetone. The solution was filtered into the vials, and the samples were stored at −20 °C until further analysis.
Phytohormones were analyzed on a UHPLC system (Dionex UltiMate 3000; Thermo Scientific, Waltham, MA, USA) coupled to a tandem mass spectrometer (LTQ XL; Thermo Scientific, Waltham, MA, USA) following the modified method by Almeida Trapp et al. [
41]. The C18 column (150 × 4.60 mm; 3 µm column; Gemini; Phenomenex, Torrance, CA, USA) was used for the separation of strigol, orobanchol and 5-deoxystrigol. The temperature was set to 50 °C. The solvents used for the analysis were solvent A, comprising 0.1% formic acid with 3% acetonitrile in double distilled water (
v/
v/
v), and solvent B, comprising 0.1% formic acid with 3% double distilled water in acetonitrile (
v/
v/
v). The elution profile was 0–1 min, 0–25% B; 1.1–7.5 min, 25–45% B; 7.6–12 min, 45–70% B; 12–15 min, 70–10% B; and 15–18 min, 10–0% B. The mobile phase flow rate was 0.4 mL min
−1, and the injection volume was 20 µL. The phytohormones were identified by coupling the UHPLC system with a tandem mass spectrometer (LTQ XL; Thermo Scientific, Waltham, MA, USA) using heated electrospray ionization. The mass spectrometer was operated in a positive ion mode to identify the strigolactones. To identify the strigolactones, the following parameters were used: capillary temperature 200 °C, capillary voltage 8.0 V, sheath temperature 280 °C, sheath gas 35 arb, ion spray voltage 4.5 kV, tube lens 45 V and auxiliary gas 5 arb. Selected Reaction Monitoring (SRM) was used to identify these selected compounds after determining the retention time and identifying typical fragmentation patterns of the obtained quantification standard. The transitions with the optimized parameters and retention times are listed in
Table 6.
4.6. Flavonoid Analyses
The phenolic compounds were extracted according to the protocol described by Medic et al. [
44]. A total of 100 mg of the freeze-dried sample was extracted with 2 mL of 80% methanol in double-distilled water. The extraction ratio was 1:20 (
w/
v). The samples were then shaken (TOP-MIX 94500 vortex mixer; Heidolph, Schwabach, Germany), sonicated for 60 min in ice water (Sonis 4 ultrasonic bath; Iskrapio, Slovenia) and centrifuged at 10,000×
g at 4 °C for 10 min. The samples were filtered with a 0.2 μm polyamide filter (Chromafil AO 20/25; Macherey-Nagel, Düren, Germany) and stored at −20 °C for further analysis. An LTQ XL tandem mass spectrometer with heated electrospray ionization in negative mode connected to a Vanquish UHPLC system (Thermo Scientific, Waltham, MA, USA) with a diode array detector at 350 nm was used to identify and quantify the flavonoids. All parameters were used as described by Medic et al. [
44]. The flavonoid compounds identified in
Castanea sp. are listed in
Table 7 and
Figure S2. The quantification of the different flavonoid compounds is given in
Table 2. The individual compounds were separated by HPLC and identified by a comparison with internal standards. All parameters were used as described by Medic et al. [
44].
4.7. Chemicals
The standards used to determine the phenolic compounds in the samples were quercetin-3-O-galactoside, quercetin-3-O-glucoside, quercetin-3-O-rhamnoside and quercetin-3-O-rutinoside from Sigma-Aldrich Co. (St. Louis, MO, USA). Isorhamnetin-3-O-glucoside was purchased from Extrasynthese (Genay, France) and quercetin-3-O-glucuronide, quercetin-3-O-xyloside, isorhamnetin-3-O-rutinoside and isorhamnetin-3-O-glucuronide were purchased from Fluka Chemie (Steinhein, Germany).
The following standards were used to identify and quantify the phytohormones: orobanchol, 5-deoxystrigol and strigol (Sigma-Aldrich Chemie GmbH, Steinheim, Germany). For extraction and further analysis, we used formic acid, ethyl acetate and acetonitrile (Sigma-Aldrich Chemie GmbH, Steinheim, Germany).
Acetonitrile for the mobile phases was HPLC-MS grade (Fluka Chemie GmbH, Buchs, Switzerland). Methanol and formic acid were HPLC grade (Sigma-Aldrich Chemie GmbH, Steinheim, Germany). The bi-distilled water was purified using a water purification system (Milli-Q, Millipore, Bedford, MA, USA).
4.8. Statistical Analysis
The data were collected with Microsoft Excel 2016 and statistically analyzed with R commander (package Rcmdr) version 4.1.0. A one-way analysis of variance (ANOVA) was performed. The “emmeans” package and Duncan tests were used for pairwise post hoc comparisons. A residual plot was used to analyze the normality and homoscedasticity of the data. All data were presented as means ± standard error (SE). Statistical means were calculated at a confidence level of 95% to determine the significance of differences (p < 0.05).