Determination of Phytohormones in Plant Extracts Using In-matrix Ethyl Chloroformate Derivatization and DLLME–GC–MS

Published on: 
, , , ,

LCGC Europe

LCGC Europe, LCGC Europe-06-01-2013, Volume 26, Issue 6
Pages: 310–324

A fast and simple sample preparation method for the simultaneous determination of 11 phytohormones in plants by GC–MS has been developed.

A fast and simple sample preparation method using in-matrix derivatization and dispersive liquid–liquid microextraction (DLLME) for the simultaneous determination of 11 phytohormones in plants by gas chromatography–mass spectrometry (GC–MS) was developed. In this derivatization–extraction procedure, phytohormones in aqueous samples were derivatized with ethyl chloroformate (ECF) and extracted by DLLME simultaneously using ethanol–pyridine (4:1, v:v), both as derivatization catalyst and DLLME dispersant. This proposed rapid and convenient method was also successfully applied for analysing the phytohormones in rice seed callus and cucumber fuits, indicating other wide applications in other plant tissues.

The first phytohormone, auxin, was discovered in 1926, and since then an increasing number of naturally-occurring or synthetic molecules with plant-growth regulation activities have been reported, including abscisic acid (ABA); salicylic acid (SA); gibberellic acid (GA).phenylacetic acid (PAA); 1-naphthylacetic acid (NAA); and 2,4-dichlorophenoxyacetic acid (2,4-D). Numerous aspects of physiological processes in plants — such as seed germination, shoot elongation, and organogenesis — are manipulated delicately by corresponding phytohormones (1,2). Some phytohormones are also involved in the adaptive behaviour of plants in response to environmental and biological stresses (3–6). Many studies have shown evidence that synergistic, as well as antagonistic, actions occur between different phytohormones in plants (7–9). In addition, signalling crosstalks between several phytohormones in regulating plant development are reported instead of their individual effect (10,11). It is therefore necessary to develop reliable methods for the simultaneous monitoring of different phytohormones during physiological processes (12,13).

Many analytical procedures have been developed to determine the importance of phytohormones in plants simultaneously (12–15), however, it is still an analytical challenge. This is because there are low concentrations of phytohormones in plants and the sample matrix is complex. Most analytical methods to determine phytohormones rely heavily on high performance liquid chromatography (HPLC) (14,15); gas chromatography (GC) (16,17); and capillary electrophoresis (CE) (18) for separation. HPLC with tandem mass spectrometry (HPLC–MS–MS) is suitable for phytohormone analysis (14, 19), but requires expensive equipment and is generally expensive. Gas chromatography coupled with mass spectrometry (GC–MS) is preferred because of its cost-effectiveness and improved separation, however, it always requires a derivatization step to improve the volatility and sensitivity of some phytohormones (17, 20).

Despite great advances in instrumentation, most analytical instruments cannot handle sample matrices directly. Sample preparation steps are commonly introduced to transfer analytes into a form that is pre-purified, concentrated, and compatible. Solid-phase extraction (SPE), combining integrated purification and concentration, is most commonly used as the sample pretreatment technique for phytohormones determination (16,19).

However, SPE is laborious, time-consuming, and requires a larger volume of sample (scarce in most plant physiological research projects) because of its low enrichment factor. These problems can be resolved using solid-phase microextraction (SPME) (15) and liquid-phase microextraction (LPME) (13). However, the high cost of SPME fibre and the operational difficulties of LPME mean that they are not widely adopted by other researchers. In addition, SPME and LPME require special conditions and long extraction times for equilibrium — during which the degradation of several labile phytohormones may potentially occur (15).

In recent years, a rapid and simple method termed dispersive liquid–liquid microextraction (DLLME) has been developed by Assadi and co-workers (21). DLLME has now been introduced in the extraction of polybrominated diphenyl ethers (PBDEs), organophosphorus pesticides (OPPs), and other organic pollutants from aqueous samples. It has a high extraction efficiency, as well as being convenient and inexpensive (22,23). With several years of development, in situ derivatization combined with DLLME has also been applied to GC (GC–MS) analysis of polar compounds such as fatty acids, chlorophenols, and anilines (24–26). Compared with post-derivatization that requires special conditions that will introduce extra steps in the sample preparation procedure, DLLME with in situ derivatization is both cost-effective and convenient. In addition, in situ derivatization can reduce the hydrophilicity of polar analytes and thus can enhance extraction efficiency.

Among the derivatizing reagents studied, alkyl chloroformate (ACF) was superior for derivatization of amines, fatty acids, phenoic acids, and amino acids in bio-fluid matrix while leaving sugars and other related compounds unaffected (27). ACF also exhibited superiority when used as an in situ derivatization reagent for DLLME. The organic catalyst (namely alcohol, pyridine, acetonitrile, and other water-miscible solvents) can spontaneously act as dispersant. Simultaneous derivatization and DLLME using ethyl chloroformate (ECF) as the derivatizing agent were first reported by Pusvaskiene for the analysis of fatty acids (24). In a previous study, simultaneous derivatization and DLLME using methyl chloroformate (MCF) as the derivatization reagent was successfully applied for the GC–MS analysis of alkylphenols (APs) in river water samples (28). These two successful applications implied that ACF in combination with DLLME could provide an efficient method for derivatization and extraction of phytohormones that contained carboxylic and hydroxyl groups from plant tissue extracts for GC–MS analysis.

In this study, an in-matrix ECF derivatization and DLLME for the determination of 11 phytohormones in plant tissue extracts is reported for the first time. This study emphasizes the use of an organic catalyst [in this instance ethanol and pyridine at the ratio of 4:1 (v:v)] as the dispersant. Some key parameters — including the amount of catalyst–dispersant, ECF and extraction solvent, pH, and ionic strength — that might affect both derivatization and DLLME were thoroughly investigated and optimized. The established method was validated for an analysis of cucumber fruit extract and was applied to the monitoring of phytohormones in rice seed callus.


Standards and Reagents: Indole-3-acetic acid (IAA); indole-3-butyric acid (IBA), indole-3-propionic acid (IPA); benzoic acid (BA); phenylacetic acid (PAA); p-hydrophenylacetic acid (HPA); salicylic acid (SA); 4-chlorophenoxyacetic acid (4-CPA); 2,4-dichlorophenoxyacetic acid (2,4-D); 1-naphthylacetic acid (NAA); (±) abscisic acid (ABA); and ethyl chloroformate (ECF) were purchased from Sigma-Aldrich. The structures of the phytohormones are shown in Figure 1. HPLC-grade ethanol; acetone; chlorobenzene; chloroform; tetrachloroethylene; carbon tetrachloride; and pyridine were bought from Merck. Sodium hydroxide (NaOH); sodium chloride (NaCl); hydrochloric acid (HCl); and other reagents (all of analytical grade) were supplied by Guangzhou Chemical Reagent Factory. Ultrapure water was produced using Milli-Q Advantage A10 system (Millipore).

Figure 1: Chemical structures of the phytohormones studied.

Stock solutions of individual standard compounds (10 mmol/L in acetone) were prepared by dissolving an appropriate amount of each substance in acetone, making up to 50 mL in a volumetric flask. A mixed stock solution containing 100 μmol/L of each compound was obtained by 100 times dilution of these stock solutions (from 1 mL to 100 mL) with acetone. These solutions were then stored in amber glass vials with Teflon-lined caps at 4 °C. From the mixed stock solution, a working solution containing 1.0 μmol/L of each compound was freshly prepared every week by diluting the mixed stock solution with ultrapure water and storing at 4 °C in the dark until use.

Plant Samples: Samples of cucumber (Cucumis sativus L.) fruits were purchased from a local supermarket. In this study, 100 g of fresh cucumber fruits were homogenized by a food processor, and the resulting fluid centrifuged at 10,000 rpm (Sigma 3K18) to remove cell debris. The supernatant was withdrawn and filtered through a 0.45- μm cellulose acetate membrane, and the resulting supernatant was used for method validation. A sample of 50 mg of rice (Oryza sativa L.) seed calluses (harvested from 30 days' culture in Murashige and Skoog medium) were ground into powder with liquid nitrogen using a porcelain mortar. A 10 mL aliquot of 50 mM potassium phosphate buffer (PB buffer, pH 7) was then added to extract phytohormones from the calluses, followed by centrifugation at 10,000 rpm (Sigma 3K18) for 30 min. The resulting supernatant was withdrawn and diluted with PB buffer. All samples were analysed immediately to prevent any degradation of facile phytohormones.

Simultaneous Derivatization and DLLME Procedure: For simultaneous derivatization and DLLME, a 5-mL aliquot containing 1.0 μmol/L of each compound was placed in a 10 mL screw-cap conical-bottom glass centrifuge tube (height × diameter: 9 cm × 1.5 cm, conical diameter: 0.3 cm; manufactured by Bomex glassware factory [Beijing, China] according to our design), and 1.5 mL ethanol–pyridine mixture (4:1, v:v) was supplemented both as reaction catalyst and dispersant. Then, 100 μL ECF (derivatization reagent) and 55 μL CHCl3 (extraction solvent) were rapidly injected into the aqueous solution with a 100 μL syringe. The tube was tightly capped and shaken vigorously for about 10 s to completely mix the different phases. The tube was then placed in an ultrasonic bath for 5 min of ultrasonication with the cap loosened and CHCl3 was dispersed into the aqueous phase by ultrasonication.

In this step, the phytohormone derivatives were extracted from the aqueous phases into the fine droplets of CHCl3. After centrifuging at 5000 rpm for 5 min, the fine droplets of CHCl3 were sedimented at the bottom of the centrifuge tube. The sedimented CHCl3 (20±1.5 μL) was withdrawn using a 50 μL syringe and subsequently stored in a 2 mL GC vial equipped with a 100 μL insert for automated injection. Finally, 1 μL sample was injected into the GC port for GC–MS analysis. All samples were performed in triplicate. The entire procedure was conducted in a fumehood with gloves and a mask, because of formation of noxious gas during the derivatization step.

In-situ Derivatization and SPE Procedure: An SPE procedure was performed to compare the extraction efficiency with DLLME. Briefly, 5.00 mL sample was supplemented with 1.5 mL ethanol–pyridine mixture (4:1, v/v), and then derivatized with 100 μL ECF. After 5 min of ultrasonication for derivatization and degasification, this solution was diluted to 100 mL with ultrapure water to reduce the organic content of the sample in case it interfered in the SPE procedure. The 100 mL aqueous sample was then passed through a preconditioned C18 cartridge at a flow rate lower than 5 mL/min. After washing the cartridge with 10 mL of ultrapure water, the cartridge was dried for 60 min under a gentle flow of nitrogen, and the analytes were then eluted to 20-mL vials from the sorbents with 10 mL ethyl acetate at a flow rate of 1 mL/min. The extract from SPE was evaporated to dryness under a gentle nitrogen stream. The residue was then dissolved in 50 μL hexane for GC–MS analysis.

Enrichment Factor and Extraction Recovery: The enrichment factor (EF) was defined as the ratio between the analyte concentration in the sedimented phase (Csed) and the initial concentration of analyte (C0) within the sample. The extraction recovery (ER) was defined as the percentage of the total amount of analyte extracted to the sedimented phase. These definitions were in accordance with the published literature (21, 23). The Csed was calculated from the calibration graph obtained by GC–MS determination of the phytohormone derivatives. In brief, 5.0 mL standard solutions with the concentrations ranging from 1 μmol/L to 100 μmol/L in acetone were dried under a gentle nitrogen stream. The dry residues were derivatized by 50 μL ECF in the presence of 100 μL ethanol–pyridine mixture (4:1, v:v) for 5 min and were blown to dryness under a nitrogen stream. The residues were dissolved in 1.0 mL chloroform for GC–MS determination. Five standard solutions were prepared to cover the calibration range of 5–500 μmol/L for each analyte.

GC–MS Analysis: Sample analysis was performed with an Agilent 6890A-5973N GC–MS instrument (Agilent Technologies) equipped with a Gerstel MPS2 multipurpose autosampler (Gerstel). A fused-silica capillary column was used (30 m × 0.25 mm, 0.25 μm DB-5MS) (Agilent Technologies). Helium was used as the carrier gas at a constant flow rate of 1.0 mL/min. The GC injection port was equipped with a 4-mm i.d. glass liner operated at the splitless mode and held isothermally at 280 °C. The oven temperature was programmed initially at 50 °C, which was then raised to 200 °C at 10 °C/min and to 215 °C at 3 °C/min, and was finally raised to 280 °C at 15 °C/min. MS was performed in the electron impact mode (EI) at 70 eV with the interface (transfer line) temperature set at 280 °C. For quantitative determinations, the mass selective detector (MSD) (Agilent Technologies) was operated in selected ion monitoring (SIM) mode (Table 1).

Table 1: Ions for the quantitative and qualitative analysis of target compounds after ethyl chloroformate (ECF) derivatization.

Results and Discussion

Optimization of In-situ Derivatization and DLLME: Chloroformates are known as strong and rapid derivatizing reagents for derivatization of amines, fatty acids, phenols, and amino acids in matrices, with the reaction mechanisms comprehensively described in the literature (27, 29, 30). For phytohormones containing both phenolic hydroxyl and carboxylic groups — exemplified by SA and HPA — the O-ethoxycarbonylation (EOC) of these groups by ECF produces the relevant ethoxycarbonyl ester (mixed anhydride). Unlike the stable phenolic ethoxycarbonyl ester, carboxylic ethoxycarbonyl esters are unstable and undergo further decarboxylation catalyzed by pyridine to yield the relevant carboxylic ethyl ester. The major drawback of ACF for the derivatization of carboxylic groups lies in the by-products formed though ester transformation between the alcohol catalyst and alkoxycarbonyl ester. To resolve this problem, alcohol with the identical molecular structure to the alkyl part of ACF was always used as the catalyst (30). The reaction scheme of SA with ECF in the presence of ethanol and pyridine is illustrated in Figure 2. Figure 3 shows that all phytohormones were successfully derivatized and exhibited satisfactory chromatographic resolution, except ABA which produced two peaks in the chromatogram. The double peak resulted from the cis- and trans- isomerization of ABA-ethyl ester (31). Table 1 reveals the molecular weights and most intense mass spectra fragments of the phytohormone derivatives obtained in the electron impact mode (70 eV) and their corresponding retention time.

Figure 2: Reaction scheme of phytohormones (exemplified by salicylic acid) with ethyl chloroformate in the presence of ethanol and pyridine.

Selection of Extraction Solvent: The extraction solvent is important for DLLME, and should have a higher density than water and low water solubility. The extraction efficiency of four solvents, namely chloroform (CHCl3), chlorobenzene (C6H5Cl), carbon tetrachloride (CCl4), and tetrachloroethylene (C2Cl4) were compared. A series of sample solutions were firstly derivatized with 100 μL ECF in the presence of 1.0 mL ethanol–pyridine mixture (4:1, v/v) by 5 min of ultrasonication. To obtain the same volume of sedimented phase (20± 1.5 μL) for comparing the extraction efficiencies among different extraction solvents, different volumes of CHCl3 (52 μL), CCl4 (33 μL), C6H5Cl (28 μL), and C2Cl4 (27 μL) were added. Table 2 shows that CHCl3 had the highest extraction efficiency of all of the phytohormones studied. Low extraction recoveries obtained for BA (30.03%) and SA (12.63%) could be attributed to their chemical structures, probably because the carboxylic groups that directly attached to the benzene ring could not be derivatized easily in aqueous solution as reported by previous researchers (29). Therefore, CHCl3 was selected as the extraction solvent for the following experiments.


Figure 3: Total ion chromatogram (TIC) of 11 phytohormones in aqueous samples after in situ derivatization dispersive liquid–liquid extraction (DLLME). Sample was spiked with 1 μmol/L of each phytohormone.

Effect of Prior Derivatization and Simultaneous Derivatization and Extraction: Two series of experiments were performed to evaluate the effects of different derivatization procedures, including prior derivatization before extraction and simultaneous derivatization with extraction. For the prior derivatization, analytes at a concentration of 1.0 μmol/L in 5.0 mL working solution were spiked with 1.0 mL ethanol–pyridine mixture (4:1, v:v) as the catalyst–dispersant and derivatized by 100 μL ECF for 5 min under ultrasonication. Following that, 52 μL CHCl3 was added and extracted by ultrasonication for 5 min. For the simultaneous derivatization and extraction, 100 μL ECF and 52 μL CHCl3 were simultaneously injected into the 5.0 mL working solution with 1.0 mL ethanol–pyridine mixture (4:1, v:v), followed by 5 min of ultrasonication. Better sensitivities were obtained when simultaneous derivatization and extraction was performed for most of the phytohormones than the staged procedure, except BA and SA, which showed similar sensitivities between staged and simultaneous procedures. Therefore, simultaneous derivatization and extraction was chosen as the pretreatment procedure because of its high sensitivity and convenience.

Table 2: Efficiency of different solvent for the extraction recovery (ER %) of phytohormones by dispersive liquid–liquid extraction (DLLME) a.

Effect of Organic Catalyst–Dispersant Volume: In DLLME, an appropriate dispersant should be miscible with both extraction solvent and the aqueous sample. Acetone, acetonitrile, methanol, ethanol, and other water soluble organic solvents that met this criterion could be used (21–23). For simultaneous ECF derivatization and DLLME, however, the water-miscible organic catalyst (ethanol–pyridine mixture) can act as DLLME dispersant spontaneously. Since the organic catalyst is essential for ECF derivatization, it should be in ratio to water for the best derivatization efficiency. Ethanol–pyridine mixture (4:1, v:v) was therefore used as the bi-functional catalyst for derivatization and dispersant for DLLME to facilitate the derivatization reaction and to avoid the formation of side products. This ratio of alcohol to pyridine was adopted in most studies for catalysis of the alkyl-O-carboxylation (AOC) reaction (24, 32, 33), and was also found to be more effective for ECF derivatization of the selected phytohormones according to our preliminary experiments (data not shown). When obtaining the optimal volume of ethanol–pyridine mixture (4:1, v:v) both for derivatization and DLLME, different volumes of ethanol—pyridine mixture (4:1, v:v) ranging from 0.5 mL to 2.5 mL at intervals of 0.5 mL were investigated. As variation in the volume of catalyst–dispersant (ethanol–pyridine mixture) could change the volume of the sedimented phase, the volume of CHCl3 was changed accordingly to obtain a constant volume of sedimented phase (20± 1.5 μL). A measurement of 100 μL ECF with 48, 52, 55, 68, and 86 μL CHCl3 was selected, respectively. As illustrated in Figure 4, the enrichment factors of phytohormones increased with the volumes of ethanol–pyridine mixture in the range of 0.5 mL to 1.5 mL, but decreased from 1.5 mL to 2.5 mL. When acting as the catalyst, the organic content in the aqueous phase should be as high as possible to enhance the reaction activities of derivatization (30), however, excessive organic content could cause low extraction efficiency for DLLME. The volume of ethanol–pyridine mixture (4:1, v/v) was therefore set at 1.5 mL as a compromise.

Figure 4: Effect of catalyst–dispersant volume on the derivatization and extraction of phytohormones. Experimental conditions: Sample volume: 5.0 mL; catalyst–dispersant: ethanol–pyridine (4:1, v/v) mixture; extraction solvent: CHCl3; derivatization and extraction time: 5 min.

Effect of CHCl3Volume: To investigate the effect of extraction solvent volume on the enrichment factor and extraction recovery, different volumes of CHCl3 (55 μL, 60 μL, 80 μL, and 100 μL) were examined. The other conditions remained as mentioned previously. It was clear that by increasing the volume of CHCl3, the volume of the sedimented phase increased (from 20 μL to 74 μL). Although the extraction recoveries of phytohormones increased with the increment in the volume of CHCl3 (Table 3), the enrichment factors decreased, probably because of the diluting effect. High enrichment factors were obtained at a low volume of CHCl3 (55 μL) and this volume was selected for subsequent experiments.

Table 3: Efficiency of different volume of CHCl3 for the extraction recovery (ER %) of phytohormones by DLLME a.

Effect of ECF Volume: When used for in situ derivatization, the major use of ACF lies in its hydrolysis in water. The derivatization efficiency is not affected until the volume of ACF used for derivatization is higher than hydrolysis. Different volumes of ECF (50 μL, 75 μL, 100 μL, and 150 μL) were tested to find the appropriate volume for the derivatization of phytohormones. The results revealed that the enrichment factor of analytes increased when the ECF content was raised in the range of 50–100 μL and then approached a plateau; an extra lift in the volume of ECF to 150 μL did not enhance the EF of the analytes (Figure 5). This phenomenon indicated that the derivatization efficiency was not affected by the volume of ECF unless it was more than 100 μL. Therefore, 100 μL ECF was used for the following experiments.

Figure 5: Effect of ethyl chloroformate amount on the derivatization and extraction of phytohormones. Experimental conditions: Sample volume: 5.0 mL; catalyst–dispersant: 1.5 mL ethanol–pyridine (4:1, v/v) mixture; derivatization reagent: 100 μL ECF; extraction solvent: 55 μL CHCl3; derivatization and extraction time: 5 min.

Effect of Derivatization and Extraction Time: In simultaneous derivatization and DLLME, the derivatization and extraction time began with the injection of the mixture of derivatizing agent and extraction solvent and ended at centrifugation. The derivatization and extraction time was examined at 1 min, 3 min, 5 min, and 10 min under ultrasonication. No significant differences were observed for enrichment factors, indicating that 1 min of ultrasonication was enough for derivatization and extraction. Derivatization of hydroxyl and carboxylic groups by ACF in the aqueous sample is a fast reaction that could be completed within several seconds. On the other hand, DLLME equilibrium time is short (cloudy phase formation is considered to be the equilibrium status). However, when derivatization and extraction time was less than 5 min, gas bubbles in the sedimented CHCl3 could affect the withdrawal procedure. A similar result was also found in our previous study (28). These findings suggested that the elimination of air from the solution was essential for the simultaneous ACF derivatization and DLLME. Therefore, the derivatization and extraction time was selected at 5 min.

Effect of pH: Sample pH could potentially affect extraction and derivatization, therefore different pH values of the aqueous solution (pH 3, 7, and 10) adjusted by HCl or NaOH were examined. The other experimental conditions were the same as mentioned previously. Figure 6 shows that the enrichment factors were similar between pH 3 and pH 7 for most phytohormones, except for IAA, IPA, IBA, and AB which had the highest EF at pH 7, probably because of a phytohormone instability under acidic conditions. The enrichment factors were the lowest for all target compounds at pH 10. These findings were different from our previous study which had suggested that both the derivatization and extraction of alkylphenols were not affected by the variation in pH value (28). The differences in the reaction pathway between the phenolic hydroxyl group and carboxylate group could be the reason. It should also be noted that the pH value of each individual experiment decreased remarkably after derivatization because of the production of HCl during the EOC process. To study the effect of pH on the DLLME extraction, the pH value of each individual experiment was adjusted to its original value by 1 M NaOH after derivatization (before DLLME was performed). No statistical differences were found between the two sets of experiments, indicating that variations in pH value affected the derivatization but not the extraction.

Figure 6: Effect of pH value on the derivatization and extraction of phytohormones. Experimental conditions: Sample volume: 5.0 mL; catalyst–dispersant: 1.5 mL ethanol–pyridine (4:1, v/v) mixture; derivatization reagent: 100 μL ECF; extraction solvent: 55 μL CHCl3; derivatization and extraction time: 5 min.

In the case of real sample analysis, the effect of pH on the derivatization could be neglected, as the pH value of most plant extracts is strictly controlled by the relevant extraction buffer (in this instance, PB buffer). Thus the optimized pH value was set at 7 for the following experiments.

Effect of Ionic Strength: The ionic strength was studied by spiking NaCl at concentrations ranging from 0 g/L to 150 g/L, into the sample solution. The sedimented phase was enlarged from 20± 1.5 μL to 58±1.7 μL with increments in ionic strength. Enlargement in the volume of the sedimented phase could cause a decline in the enrichment factor for the phytohormones because of the diluting effect and resultant reduction in the sensitivity. Salt addition could also increase the density of the aqueous sample, thus the sedimented phase could be easily suspended in the aqueous phase and became difficult to withdraw. Since most extracting buffers contain a high salt content (NaCl, K2HPO4, KH2PO4) the effect of ionic strength on DLLME should be considered in real sample analysis. Properly diluting the sample or preparing the calibration graph with the relevant extracting buffer would be necessary.

Method Evaluation and Sample Analysis

Evaluation of the Method: The proposed method was evaluated with respect to the linear range, correlation coefficient (R2), precision (relative stand deviation [RSD] %), limit of detection (LOD), and limit of quantification (LOQ). The LOD and LOQ were calculated based on signal-to-noise ratio (S/N) of 3 and 10, respectively. Ultrapure water (pH 7) spiked with different concentrations of 11 phytohormones was used for the calculation.

Table 4 shows that the calibration ranges of 11 phytohormones using DLLME spanned over four orders of magnitudes (from 1 nmol/L to 10 μmol/L). The RSDs of seven replicates at 1 nmol/L were less than 15% for most phytohormones. With regards to sensitivity, the LOD and LOQ of the analytes were in the range of 0.05 nmol/L to 0.21 nmol/L and 0.18nmol/L to 0.69 nmol/L. Therefore, in matrix ECF derivatization and DLLME provides a simple, convenient, and inexpensive sample preparation method for GC–MS determination of phytohormones.

Table 4: Linear range, enrichment factor, limit of detection (LOD), and limit of quantification (LOQ) in pure water obtained by dispersive liquid–liquid extraction gas chromatography–mass spectrometry (DLLME–GC–MS).

Real Sample Analysis

Cucumber (Cucumis sativus L. fruits): After optimization, the proposed method was applied to the analysis of phytohormones in cucumber fruits. When the fluids of the cucumber fruit were analysed directly, precipitation of water-soluble protein that occurred during the derivatization process was found to interfere with the DLLME extraction. The cucumber fruit fluids were therefore diluted 2× and 10× with ultrapure water to reduce the matrix effect. Recoveries of phytohormones were evaluated in real samples spiked with 0.05 μmol/L of each analyte. The recoveries were calculated by subtracting the results of the non-spiked samples from those of the spiked samples. The matrix effect was also studied by comparing the present optimized method with the conventional SPE procedure. Figure 7 shows the chromatograms of 10× diluted sample and the sample spiked with 0.05 μmol/L of each phytohormone. The concentrations of phytohormones obtained with DLLME matched reasonably well with that of SPE (Table 5). The matrix effect could be successfully handled by 10× dilution of the cucumber fruit extracts. However, matrix effects were observed in extract with 2× dilution, and the extraction recoveries of phytohormones decreased to between 60.3% and 89.8%. This result indicated that when performed with bio-fluids such as plant extracts, the sample matrix could interfere with the DLLME procedure and appropriate dilution of the extracts would be necessary. In addition, extract concentration reduction could also be achieved by reducing the sample size of the plant tissues. This could be a great advantage in some physiological research where the available sample is limited.

Figure 7: Total ion chromatogram (TIC) of phytohormones in (a) cucumber fruit fluid (10× dilution with ultrapure water) before spiking, and (b) spiked with 0.05 μmol/L of each phytohormone using in-matrix derivatization and DLLME method under optimum conditions.

Rice (Oryza sativa L.) Seed Calluses: Plant callus is a superior model for plant physiology research. The induction and differentiation of plant calluses are controlled by endogenous and exogenous phytohormones. In addition, plant transgenic techniques are applied to plant calluses to obtain mutant strains for phytohormone research. Consequently, the determination of phytohormones in plant calluses is important to understand their mechanism in plant physiology. In the present study, rice seed calluses were extracted and analysed using the proposed method mentioned above. A calibration curve was prepared by successive dilution of the mixed stock solution with 50 mM PB buffer (pH 7). Satisfactory recoveries (77.2–108.5%) were obtained from the extracts, as well as the extracts spiked with 0.05 μmol/L of each analyte. Among the phytohormones studied, the concentrations of BA, PAA, SA, NAA, HPA, and IAA in the rice seed callus were 0.019±0.007, 0.218±0.093, 0.004±0.001, 0.007±0.003, 0.010±0.007, and 0.006±0.002 nmol/mg, respectively. Noticeably, NAA, an exogenous phytohormone that is commonly supplemented to the Murashige and Skoog medium, was detected in the calluses. It is also important to note that the sample amount (50 mg) could be afforded by one strain of callus, and the sample volume (25 mL) was sufficient for more than three replicates of analysis.

Table 5: Concentration and recovery of phytohormones in cucumber fruits obtained by DLLME and solid-phase extraction (SPE).


The simultaneous in-matrix ethyl chloroformate derivatization and DLLME coupled with GC–MS technique was successfully established for the determination of 11 phytohormones in plant tissues. The experimental results revealed that this method could provide high enrichment factors and high extraction recoveries within a short time (about 5 min). The procedure was validated and successfully applied to the analysis of phytohormones in cucumber fruits and rice seed calluses. The proposed method was not only precise and sensitive, but also required less organic solvent; derivatization and extraction could be conducted in a single step. Compared with traditional sample preparation techniques which always require more than 1 g of plant tissue, this technique could function with much less sample (0.05 g). This is of great importance for plant physiology research when samples are rare (such as transgenic plants) or hard to obtain (such as microalgae).


Financial support from the National Natural Science Foundation of China (NSFC, No. 21277177), the Fundamental Research Funds (No. 2012-509) by Administration of Ocean and Fisheries of Guangdong Province, and the China Postdoctoral Science Foundation (No. 2013M531855) are gratefully acknowledged.

Shusheng Luo is an engineer at the South University of Science and Technology of China, Shenzhen, China.

Li Lin is an associate professor at the School of Life Sciences, Sun Yat-sen University, Guangzhou, China.

Xiaowei Wang is a lecturer at the School of Marine Sciences, Sun Yat-sen University, Guangzhou, China.

Shichun Zou is a professor at the School of Marine Sciences, Sun Yat-sen University, Guangzhou, China.

Tiangang Luan is a professor at the School of Life Sciences, School of Marine Sciences, Sun Yat-sen University, Guangzhou, China.


(1) S.J. Davis, Plant Cell Environ. 32(9), 1201–1210 (2009).

(2) T. Kai, K. Nimura, H. Yasui, and H. Mizuta, J. Appl. Phycol. 18(1), 95–101 (2006)

(3) O. Atici, G. Agar, and P. Battal, Biol. Plant 49(2), 215–222 (2005).

(4) K.A. Lahey, R.C. Yuan, J.K. Burns, P.P. Ueng, L.W. Timmer, and K.R. Chung, Mol. Plant-Microbe Interact 17(12), 1394–1401 (2004).

(5) K. Urano, K. Maruyama, Y. Ogata, Y. Morishita, M. Takeda, N. Sakurai, H. Suzuki, K. Saito, D. Shibata, M. Kobayashi, K. Yamaguchi-Shinozaki, and K. Shinozaki, Plant J. 57(6), 1065–1078 (2009).

(6) D. Ogawa, N. Nakajima, T. Sano, M. Tamaoki, M. Aono, A. Kubo, M. Kanna, M. Ioki, H. Kamada, and H. Saji, Plant Cell Physiol. 46(7), 1062–1072 (2005).

(7) H. Hansen and K. Grossmann, Plant Physiol. 124(3), 1437–1448 (2000).

(8) D. Wang, K. Pajerowska-Mukhtar, A.H. Culler, and X.N. Dong, Curr Biol. 17(20), 1784–1790 (2007).

(9) C.M.J. Pieterse, A. Leon-Reyes, S. Van der Ent, and S.C.M. Van Wees, Nat. Chem. Biol. 5(5), 308–316 (2009).

(10) M.M.G. Johnstone, D.M. Reinecke, and J.A. Ozga, J. Plant Growth Reg. 24(3), 214–225 (2005).

(11) K. Hartig and E. Beck, Plant Biol. 8(3) 389–396 (2006).

(12) H. Matsuura, A. Aoi, C. Satou, M. Nakaya, C. Masuta, and K. Nabeta, Plant Growth Reg. 57(3), 293–301 (2009).

(13) Y.L. Wu and B. Hu, J. Chromatogr. A. 1216(45), 7657–7663 (2009).

(14) S.D.S. Chiwocha, S.R. Abrams, S.J. Ambrose, A.J. Cutler, M. Loewen, A.R.S. Ross, and A.R. Kermode, Plant J. 35(3), 405–417 (2003).

(15) H.T. Liu, Y.F. Li, T.G. Luan, C.Y. Lan, and W.S. Shu, Chromatographia 66(7–8), 515–520 (2007).

(16) A. Muller, P. Duchting, and E.W. Weiler, Planta 216(1), 44–56 (2002).

(17) E.A. Schmelz, J. Engelberth, J.H. Tumlinson, A. Block, and H.T. Alborn, Plant J. 39, 790–808 (2004)

(18) X. Liu, L. Ma, Y.W. Lin, and Y.T. Lu, J. Chromatogr. A 1021(1–2), 209–213 (2003).

(19) S.J. Hou, J. Zhu, M.Y. Ding, and G.H. Lv, Talanta 76(4), 798–802 (2008).

(20) F.M. Perrine, B.G. Rolfe, M.F. Hynes, and C.H. Hocart, Plant Physiol. Biochem.42(9), 723–729 (2004).

(21) M. Rezaee, Y. Assadi, M.R.M. Hosseinia, E. Aghaee, F. Ahmadi, and S. Berijani, J. Chromatogr. A 1116(1–2), 1–9 (2004).

(22) X.J. Liu, J.W. Li, Z.X. Zha, W. Zhang, K.F. Lin, C.J. Huang, and X.D. Wang, J Chromatogr A 1216(12), 2220–2226 (2009).

(23) S. Berijani, Y. Assadi, M. Anbia, M.R.M. Hosseini, and E. Aghaee, J Chromatogr. A 1123(1), 1–9 (2006).

(24) E. Pusvaskiene, B. Januskevic, A. Prichodko, and V. Vickackaite, Chromatographia 69(3–4), 271–276 (2009).

(25) N. Fattahi, Y. Assadi, M.R.M Hosseini, and E.Z. Jahromi, J. Chromatogr. A 1157(1–2), 23–29 (2007).

(26) J.S. Chiang and S.D. Huang, Talanta 75(1), 70–75 (2008).

(27) P. Hušek, J. Chromatogr. B 717(1–2), 57–91 (1998).

(28) S.S. Luo, L. Fang, X.W. Wang, H.T. Liu, G.F. Ouyang, C.Y. Lan, and T.G. Luan, J. Chromatogr. A 1217, 6762–6768 (2010).

(29) P. Hušek, Chromatographia 34(11), 621–626 (1992).

(30) I. Citova, R. Sladkovsky, and P. Solich, Anal. Chim. Acta. 573, 231–241 (2006).

(31) F.J. Zhang, Y.J. Jin, X.Y. Xu, R.C. Lu, and H.J. Chen, Phytochem Anal 19(6), 560–567 (2008).

(32) Y.C. Fiamegos, C.G. Nanos, and C.D. Stalikas, J. Chromatogr. B 813(1–2), 89–94 (2004).

(33) D.N. Dernroth, A. Rundstrom, and B. Kagedal, J. Chromatogr. A 1216(30), 5730–5739 (2009).