Planta

, Volume 240, Issue 1, pp 55–76 | Cite as

Quo vadis plant hormone analysis?

  • Danuše Tarkowská
  • Ondřej Novák
  • Kristýna Floková
  • Petr Tarkowski
  • Veronika Turečková
  • Jiří Grúz
  • Jakub Rolčík
  • Miroslav Strnad
Review

Abstract

Plant hormones act as chemical messengers in the regulation of myriads of physiological processes that occur in plants. To date, nine groups of plant hormones have been identified and more will probably be discovered. Furthermore, members of each group may participate in the regulation of physiological responses in planta both alone and in concert with members of either the same group or other groups. The ideal way to study biochemical processes involving these signalling molecules is ‘hormone profiling’, i.e. quantification of not only the hormones themselves, but also their biosynthetic precursors and metabolites in plant tissues. However, this is highly challenging since trace amounts of all of these substances are present in highly complex plant matrices. Here, we review advances, current trends and future perspectives in the analysis of all currently known plant hormones and the associated problems of extracting them from plant tissues and separating them from the numerous potentially interfering compounds.

Keywords

Plant hormones Extraction Isolation Mass spectrometry Liquid chromatography Gas chromatography 

Abbreviations

ABA

Abscisic acid

ACC

1-Aminocyclopropane-1-carboxylic acid

BRs

Brassinosteroids

CE

Capillary electrophoresis

CKs

Cytokinins

GAs

Gibberellins

GC-MS

Gas chromatography-mass spectrometry

HPLC

High performance liquid chromatography

IAA

Indole-3-acetic acid

JA

Jasmonic acid

JAs

Jasmonates

LC

Liquid chromatography

MS

Mass spectrometry

SA

Salicylic acid

SLs

Strigolactones

UHPLC-MS/MS

Ultra-high performance liquid chromatography-tandem mass spectrometry

Introduction

Most (if not all) organisms use chemical signals in cell–cell communication. Thus, chemical signalling is extremely ancient. However, the complexity of cell signalling leapt when first prokaryotic and subsequently eukaryotic cells began to associate together in multicellular organisms, putatively several billion and one billion years ago, respectively (Parfrey et al. 2011). Following the emergence of multicellularity, cell specialisation increased as tissues and organs with diverse specific functions evolved. Co-ordination of the growth and development of these cells, tissues and organs, as well as the environmental responses of complex multicellular organisms, required increasingly intricate signalling networks. Many of our current concepts about intercellular communication in plants have been derived from similar studies in animals, in which two main systems evolved: the nervous system and endocrine system. Plants, lacking motility, never developed a nervous system, but they did evolve hormones as chemical messengers. Plant hormones play essential roles (individually and in concert) in the regulation of myriads of physiological processes involved in plants’ growth, development, senescence and responses to environmental stimuli. Until the 1990s, there were just five known types of plant hormone: auxins, cytokinins, gibberellins, ethylene and abscisic acid. However, during the last two decades compelling evidence has emerged that four other classes of substances (brassinosteroids, jasmonates, salicylic acid and most recently strigolactones) act as signalling molecules and probably have growth-regulating activities.

Plant hormones, also known as ‘phytohormones’, are usually present at extremely low concentrations in plant tissues, generally pg/g fresh weight (FW), while substances that interfere with their analysis are present in far greater concentrations. This is the major problem associated with plant hormone analysis. Thus, sound knowledge of the analytical and chemical principles underlying the extraction, purification, identification and quantification of plant hormones is essential for their accurate and precise determination. In this review, we summarise current understanding of these principles, methodologies for plant hormone analysis, factors that complicate their extraction and isolation from the highly complex matrices of plant and other tissues (which contain thousands of substances) and future perspectives.

Extraction and purification

Prior to extraction, plant material must be homogenised to break the cell walls in the tissues (Harrison 2011) and thus allow any hormones present to migrate to an appropriate extraction solvent. This can be done by grinding freeze-dried or fresh plant tissue (gram amounts) in a mortar with a pestle under liquid nitrogen then adding an appropriate solvent to the ground material. Alternatively, very small amounts of plant material (mg) can be ground in 1.5–2.0 ml plastic microtubes with a selected extraction solvent and tungsten carbide or zirconium oxide beads in a homogenizer for an appropriate time at a selected frequency. The most effective devices use multi-directional motions to transmit high kinetic energy to the beads and are capable of grinding tens of samples simultaneously. To avoid enzymatic or chemical degradation of the hormones, the plant material should be kept cold during the entire homogenisation process. The efficiency of extraction of a target hormone from a plant tissue will depend on its polarity, its subcellular localisation and the extent to which it is associated with other compounds in the tissue such as phenolics, lipids, pigments and proteins (Hillman 1978). The solvent used must be capable of extracting the hormone efficiently, while minimising extraction of interfering substances. Methanol, acetonitrile, mixtures of these solvents with aqueous solutions of organic acids (generally formic or acetic) or buffers adjusted to neutral pH are usually used as extraction solvents for isolating plant hormones (Kowalczyk and Sandberg 2001; Nordström et al. 2004; Novák et al. 2008; Kojima et al. 2009; Urbanová et al. 2013). Analyte losses during the sample purification procedure can be accounted for by adding internal standards (usually labelled with stable isotopes) to the plant extracts. This procedure also provides a measure of the percentage recovery of target metabolites throughout the purification procedure. Ideally, recovery markers should be included for every plant hormone metabolite that is being quantified. However, in many studies only a few internal standards have been used, often added at late stages during the extraction process, or even just before quantitative analyses (Witters et al. 1999; van Rhijn et al. 2001). Clearly, all the current methodologies could be further improved by sophisticated internal standardisation of some of the missing labelled standards, mainly to cover the enormous variations in chemical properties of the substances, even within each phytohormone group. Dissimilar chemical nature of endogenous and internal substances subsequently leads to errors in their determination.

The ideal extraction duration depends on the target plant hormone group and (to a lesser degree) the specific target hormones. Generally, it should be long enough to allow quantitative migration of the analytes into the extraction medium and isotopic equilibration between the endogenous compounds and added internal standards. Decomposition of the endogenous hormones during prolonged extractions can be minimised by performing the extraction at low temperature (between −20 and 4 °C) and adding an appropriate antioxidant (for instance diethyldithiocarbamic acid; Pěnčík et al. 2009) to the extraction solvent.

The optimal purification method depends on the chemical nature of the target hormones, the type of analysis to be performed and choice of analytical instrument. In addition, appropriate separation procedures must be applied to reduce levels of interfering compounds in the extracts while maximising recoveries of the hormones in each purification step (Ljung et al. 2004). The first step is often liquid–liquid extraction combined with solid-phase extraction (SPE). SPE columns are packed by the manufacturers with solid sorbents that bind plant hormones (and other compounds, to varying degrees), usually via hydrophobic, polar or ionic interactions (often sorbents with hydrocarbon groups, graphitized carbon-based material and ion-exchange matrices, respectively). Interfering substances are removed by washing the column with a suitable solvent and hormones are then eluted using a solvent that disrupts the bonds formed by the interactions between the hormones and the sorbent in the column. “Mixed-mode” SPE columns, packed with a mixture of two types of sorbent, are also available and have become very popular recently (Nordström et al. 2004; Dobrev et al. 2005; Novák et al. 2008; Kojima et al. 2009; Urbanová et al. 2013) due to their ability to reduce the number of required purification steps (since more than one separation mechanism can be exploited using a single column), while maintaining high sample clean-up efficiency. SPE allows high throughput of samples when combined with automatic systems, SPE robots, which are capable of purifying tens of sample simultaneously (Nordström et al. 2004; Kojima et al. 2009). However, no miniature mixed-mode purification system capable of handling extracts from mg FW samples has been developed yet.

Auxins

Auxins were the first discovered family of plant hormones. In the earliest recorded inference, Charles and Francis Darwin concluded that plant growth is regulated by a signal transported from one part of the plant to another where the physiological growth response occurs (Darwin and Darwin 1880). This “signal” was subsequently called auxin (from the Greek word “auxein” meaning “to grow”) and identified as indole-3-acetic acid (IAA; Kögl and Kostermans 1934; Went and Thimann 1937). IAA (Fig. 1) is the major auxin involved in a plethora of physiological processes in plants. Its activities include induction of cell division and elongation in stems, and regulation of cell differentiation, various tropisms, abscission, apical dominance, senescence and flowering (Woodward and Bartel 2005; Teale et al. 2006). Two major IAA biosynthesis pathways have been postulated in plants: the tryptophan (Trp)-independent and Trp-dependent pathway (Normanly 2010; Mano and Nemoto 2012). After synthesis, IAA may be deactivated by catabolic oxidation (decarboxylative or non-decarboxylative), or conjugation to sugars and amino acids (Normanly 2010; Ljung 2012).
Fig. 1

Structures of auxins

To obtain complete understanding of IAA metabolism in a given biological sample, information on levels of free hormone, its major metabolites and biosynthetic precursors is highly important. Accurate estimation of these substances requires the detection and quantification of minute amounts of analytes in plant extracts containing huge numbers of other substances at far higher concentrations. Therefore, it is essential to use methodology that offers low detection limits and high selectivity, i.e. methods that are minimally sensitive to impurities. Several methods have been described for detecting free IAA, including HPLC with fluorescence detection (Crozier et al. 1980; Sundberg et al. 1986; Mattivi et al. 1999; Dobrev et al. 2005) or chemiluminescence detection (Xi et al. 2009), with or without enhancement by immunoaffinity-based purification techniques (Pengelly et al. 1981; Sandberg et al. 1985; Cohen et al. 1987; Marcussen et al. 1989). However, the most commonly employed method for quantifying IAA in plant tissues seems to be gas chromatography–mass spectrometry (GC–MS) with electron impact ionisation (Chen et al. 1988; Dunlap and Guinn 1989; Edlund et al. 1995; Ribnicky et al. 1998; Perrine et al. 2004; Barkawi et al. 2010). A drawback of this approach is that IAA is not volatile so it must be derivatised (usually by methylation or trimethylsilylation). Several methods for preparing derivatives of IAA precursors for GC–MS analysis have also been developed, including acylation of tryptamine, trimethylsilylation of indole-3-ethanol, and methyl chloroformate derivatisation of tryptophan (Quittenden et al. 2009; Liu et al. 2012a, b, c). Samples can be purified by reversed-phase SPE (Barkawi et al. 2008), mixed-mode SPE (Dobrev et al. 2005) or immunoaffinity extraction (Sundberg et al. 1986; Pěnčík et al. 2009). To avoid preparation of antibodies in animals, selective binding in a polymer matrix with a “molecular imprint” (MIP) of auxin can be used (Zhang et al. 2010). A miniature system for purifying IAA and its biosynthetic precursors using SPE tips has been developed (Liu et al. 2012a, b, c), and the best currently available analytical technology is based on liquid chromatography-tandem mass spectrometry (LC–MS/MS), which is capable of determining both IAA and its amino acid conjugates (Kowalczyk and Sandberg 2001; Pěnčík et al. 2009). However, this requires a much more intricate procedure than measurements of IAA alone, mainly because levels of IAA conjugates in plant extracts are significantly lower. However, all IAA metabolites except indole-3-pyruvic acid (IPyA, Fig. 1) can be analysed without any derivatisation prior to their MS detection in positive or negative electrospray mode (Kai et al. 2007a, b; Sugawara et al. 2009; Mashiguchi et al. 2011; Novák et al. 2012). Recently, IPyA (the most labile auxin precursor) has been identified as an important intermediate in the Trp-dependent IAA biosynthesis pathway in Arabidopsis (Mashiguchi et al. 2011; Stepanova et al. 2011). Tam and Normanly (1998) described a simple, rapid method for its reliable quantification based on derivatisation of the carbonyl group by hydroxylamine to form the oxime. Other methods, such as derivatisation of IPyA by 2,4-dinitrophenylhydrazone (Mashiguchi et al. 2011), cysteamine (Novák et al. 2012) or sodium borodeuteride (Liu et al. 2012a, b, c) have also been developed.

Cytokinins

Cytokinins (CKs) are endogenous N6-substituted adenine derivatives with the well-known primary ability to induce cell division activity in plant callus cultures (Skoog and Miller 1957). However, they also have a very wide spectrum of other physiological effects on various plants and tissues, notably they can delay senescence, inhibit root growth and branching, increase resistance to environmental stresses and initiate seed development (Richmond and Lang 1957; Mok 1994). As shown in Fig. 2, CKs can be divided into two subgroups based on their chemical structure: isoprenoid CKs (ISCKs), which bear an isoprenoid side chain at position N-6 and include zeatin, isopentenyl and dihydrozeatin forms; and aromatic CKs (ARCKs), which bear a side chain of aromatic (benzyl or furfuryl) origin. From a physiological perspective, there are four main types of CK metabolism: interconversion, hydroxylation, conjugation and oxidative degradation. However, the major CK metabolic processes are interconversions of CK bases, nucleosides and nucleotides (Chen 1981), as rates of CK nucleoside and nucleotide conversions to bases (the biologically active forms) reportedly control CK activity in plant cells (Kurakawa et al. 2007). Side chain modifications of ISCKs include stereospecific hydroxylation of the isopentenyl side chain, yielding zeatin (Takei et al. 2004), and reduction of the zeatin side chain, yielding dihydrozeatin (Mok and Martin 1994). Zeatin occurs naturally as two geometric isomers: trans- and cis-zeatin. In general, trans-zeatin (tZ) is considered as a cytokinin with high activity, compared to the little or no active cis-zeatin (cZ) (Kudo et al. 2012). Early investigators postulated that tZ was the predominant form, while the cis-isomer was much less abundant in planta (Schmitz and Skoog 1972; Mok et al. 1978). However, there are growing indications that cZ is the dominant cytokinin species in various plants, such as rice (Takagi et al. 1985), maize (Veach et al. 2003; Vyroubalová et al. 2009), potatoes (Suttle and Banowetz 2000) and several species of legumes (Emery et al. 1998, 2000; Quesnelle and Emery 2007). Interestingly, relative levels of zeatin stereoisomers can also differ substantially during a plant’s lifecycle, cZ-type CKs generally predominate in tissues exposed to various stresses (drought, heat or biotic stress), while tZ-type CKs are often more abundant in unstressed tissues (Havlová et al. 2008; Pertry et al. 2009; Vyroubalová et al. 2009; Dobra et al. 2010). Common modifications of ARCK side chains are regiospecific hydroxylations, leading to formation of either meta- or ortho-derivatives called topolins (Kamínek et al. 1987; Strnad 1997). Meta-position of hydroxyl functional group increases CK activity of the parent compound, while at the ortho-position leads to its decrease (Holub et al. 1998). Glycosylation, leading to the formation of N- or O-glycosides of CKs, also occurs in many plant species (Entsch et al. 1979). N-glycosides lack CK activity in bioassays, indicating that their formation is a form of irreversible inactivation (Laloue 1977). In contrast, O-glycosides are considered inactive storage forms that play important roles in balancing CK levels (McGaw and Burch 1995). Free CK bases and nucleosides with unsaturated N6-side chains may be irreversibly degraded by cleavage of the side chain catalysed by cytokinin oxidase/dehydrogenase, yielding adenine or adenosine and the corresponding side chain aldehyde (Galuszka et al. 2001).
Fig. 2

Structures of cytokinins

CK metabolites have significantly differing chemical properties that must be considered in analyses. Notably, their ionic forms are dependent on pH, which thus strongly influences their behaviour on ion-exchange columns. For instance, at a pH of ca. 2, CK nucleotides are zwitterionic (uncharged), while CK bases and several metabolites (including 9-ribosyl and 3-, 7- and 9-glycosyl metabolites) are cationic. In addition, nucleotides are more polar and thus less hydrophobic than glycosides, which in turn are more polar and less hydrophobic than CK bases and ribosides. Thus, CK metabolites’ chromatographic properties vary widely, which complicates their analysis. In the 1960s, during the GC boom, both GC–MS and GC–ECD techniques were introduced for CK analysis. However, chemical modification of hydrogen-binding functional groups was essential for converting CKs (which are not volatile; Horgan and Scott 1987) into volatile derivatives suitable for GC. Various derivatisation approaches have been published, including trimethylsilylation (TMS, Most et al. 1968), permethylation (Morris 1977) and trifluoroacetylation (TFA, Ludewig et al. 1982). However, these procedures are associated with a number of technical difficulties, such as requirements for extremely water-sensitive reagents, inappropriate and time-consuming preparation, the extreme sensitivity of some derivatives (TMS and TFA) to moisture, and the need for high temperatures to elute permethylated derivatives. To avoid the problems arising from CK derivatisation for GC, attention has focused on LC–MS. The first LC–MS method for CK analysis, involving the separation of underivatised cytokinins using a frit-fast atom bombardment interface, was published by Imbault et al. (1993). The sensitivity of this method was subsequently improved, to low femtomolar detection limits, by derivatising 10 ISCKs using propionyl anhydride to form CK propionyl derivatives (Åstot et al. 1998; Nordström et al. 2004). In addition, atmospheric pressure ionisation (APCI, Yang et al. 1993) and electrospray ionisation (ESI, Prinsen et al. 1995; Witters et al. 1999; Novák et al. 2003) interfaces have been used for CK determination, affording picomolar to low femtomolar detection limits in analyses of 0.1–1 g FW samples of plant tissue. Nowadays, ESI is the only MS interface routinely used for quantitative analysis of CKs that offers sufficient ionisation efficiency not only for CK but also for the majority of plant hormones (Novák et al. 2008; Svačinová et al. 2012; Farrow and Emery 2012; Dewitte et al. 1999). Since CKs strongly absorb UV light (in the 220–300 nm region), several LC–UV methods have been earlier applied for quantitative analysis of CKs (Campell and Town 1991; Chory et al. 1994). UV detection can be further advantageous for analyses of immunoaffinity-purified cytokinin samples (Nicander et al. 1993) and separation of CKs by capillary electrophoresis (CE; Pacáková et al. 1997; Béres et al. 2012). In some cases, CE has been found to have distinct advantages over ultra-high performance liquid chromatography (UHPLC) in terms of separation efficiency, costs and simplicity, while maintaining comparable sensitivity to MS detection (Ge et al. 2006).

As substituted purine derivatives CKs also have typical electroactive properties, so they can be detected by electrochemical reduction or oxidation using appropriate electrodes (Hernández et al. 1995; Hušková et al. 2000; Tarkowská et al. 2003). However, these methods are more useful for screening purposes than routine analysis of endogenous cytokinin levels in plant tissues.

Similarly to other plant hormones, numerous attempts have been made to increase the sensitivity, peak capacity and speed of analyses of the trace quantities of CKs present in small amounts of various plant tissues (e.g. apical roots, stem regions, seeds and buds) or even individual cell organelles. Such requirements can be fulfilled by UHPLC in combination with tandem MS. However, extremely careful attention must also be paid to the efficiency of CK extraction and isolation from plant matrices, which (as mentioned) are very complex and typically contain thousands of substances. SPE followed by a high-throughput batch immunoextraction step and subsequent UHPLC separation has proved to be highly valuable for this, allowing for example the separation of 50 CKs—including bases, ribosides, 9-glycosides, O-glycosides and nucleotides—from several milligrams of poplar (Populus × canadensis Moench, cv Robusta) leaves (Novák et al. 2008).

Recently, miniature sample pretreatments based on hydrophilic interaction liquid chromatography (HILIC) combined with MS/MS have also been used for CK analysis (Liu et al. 2010, 2012a). Further improvements allowing reductions of starting amounts of tissue while maintaining sensitivity have been achieved by miniaturisation of SPE apparatus from polypropylene columns to pipette tips, so-called stop-and-go-microextraction or StageTip purification, which affords attomolar detection limits using 1–5 mg FW of Arabidopsis seedlings (Svačinová et al. 2012). In addition, magnetic solid-phase extraction techniques, involving use of magnetic or magnetizable adsorbents with high adsorption ability and superparamagnetism, have been introduced for effective sample enrichment and purification of CKs prior to HILIC combined with tandem mass spectrometry (Liu et al. 2012b). This approach was applied to analyse CKs in 200 mg FW extracts of rice roots (Oryza sativa) and Arabidopsis thaliana seedlings with pg/mL detection limits. Another approach for improving sample enrichment is to selectively bind CKs from plant extracts using molecularly imprinted polymers (MIPs) prior to LC–MS/MS analysis. This method was developed and applied to estimate levels of two 2 ISCKs and two ARCKs in 5 g FW extracts of tobacco, rape and soybean leaves with pg/mL detection limits (Du et al. 2012).

Gibberellins

Gibberellins (GAs) are a class of diterpenoid carboxylic acids that include biologically active compounds produced by various microbes (fungal and bacterial) and lower as well as higher plants, where they are endogenous growth regulators. To date, 136 naturally occurring GAs from diverse natural sources have been characterised (http://www.plant-hormones.info/gibberellins.htm). The most prominent physiological effects of bioactive GAs (e.g. GA1, GA4, Fig. 3) include for instance induction of flowering and germination, stimulation of stem elongation and delay of senescence in leaves and citrus fruits (Hedden and Thomas 2012).
Fig. 3

Structures of C19 and C20 gibberellins

All naturally occurring GAs possess a tetracyclic ent-gibberellane skeleton consisting of 20 carbon atoms (with rings designated A, B, C and D; Fig. 3), or a 20-nor-ent-gibberellane skeleton (in which carbon-20 is missing, so there are only 19 carbon atoms). Therefore, in terms of carbon numbers, GAs can be divided into two groups: C19-GAs (e.g. GA4, GA1) and C20-GAs (e.g. GA12, GA53). The prefix ent indicates that the skeleton is derived from ent-kaurene, a tetracyclic hydrocarbon that is enantiomeric to the naturally occurring compound kaurene. Like other classes of plant hormones, concentrations of GAs in plant tissues are usually extremely low (generally pg/g FW). Thus, very sensitive analytical methods are required for their detection. However, levels of GAs may vary substantially even within a plant organ. Vegetative tissues (stems, roots and leaves) typically contain several pg/g FW, while reproductive organs (such as seeds and flowers) often have three orders of magnitude higher levels (i.e. ng/g FW). The chemical nature of GAs also varies substantially, notably they cover a broad range of polarities and the only properties they share are that they behave as weak organic acids, with dissociation constants (pKa) around 4.0 (Tidd 1964), and have no spectral characteristics such as fluorescence or UV absorption (except below 220 nm) that could easily distinguish them from other organic acids. The first methods for GA analysis, based on GC–MS determinations of volatile methyl ester trimethylsilyl ether derivatives, were introduced in the 1960s (Pryce et al. 1967; MacMillan and Pryce 1968; Binks et al. 1969). This approach is still used in some laboratories for quantifying and identifying GAs as it is highly sensitive (Mauriat and Moritz 2009; Magome et al. 2013). However, LC–MS is becoming more popular for quantitative analysis of GAs, mainly because it avoids derivatisation requirements. For instance, Varbanova et al. (2007) published a method for analysing 14 GAs in extracts of Arabidopsis mutants within 16 min by LC–MS/MS (after a laborious five-step purification procedure). The quantification procedure involved addition of deuterium-labelled internal standards before purification followed by isotope dilution analysis, as generally recommended for precise quantification (Croker et al. 1994). LC–MS/MS-based analysis has also been successfully used to determine endogenous GAs in Christmas rose (Helleborus niger L.) during flowering and fruit development (Ayele et al. 2010). Most recently, a rapid, sensitive method based on a two-step isolation procedure followed by UHPLC-MS/MS analysis has been published (Urbanová et al. 2013). This methodology is capable of quantifying 20 naturally occurring biosynthetic precursors, bioactive GAs and metabolic products from extracts of 100 mg FW plant tissues with low femtomolar detection limits.

Abscisic acid

Abscisic acid (ABA) is an optically active C15 terpenoid carboxylic acid (Fig. 4) that was discovered during the early 1960s, when it was found to be involved in the control of seed dormancy and organ abscission (Liu and Carns 1961; Ohkuma et al. 1963; Cornforth et al. 1965). Later, it was shown that the role of ABA in regulating abscission is minor and its primary role is in regulating seed dormancy and stomata opening (Patterson 2001). ABA plays important roles in many other numerous physiological processes such as seed maturation, adaptive responses to abiotic stress (Nambara and Marion-Poll 2005), shoot elongation, morphogenesis of submerged plants (Hoffmann-Benning and Kende 1992; Kuwabara et al. 2003), and root growth maintenance (Sharp and LeNoble 2002). It is a non-volatile, relatively hydrophobic substance containing a carboxylic group (Fig. 4). Therefore, commonly applied approaches for its extraction and purification include liquid–liquid extraction (Liu et al. 2002; Schmelz et al. 2003; Durgbanshi et al. 2005), liquid–liquid–liquid microextraction (Wu and Hu 2009; Bai et al. 2012), SPE (Dobrev and Kamínek 2002; Chiwocha et al. 2003; Zhou et al. 2003; Dobrev et al. 2005) and solid-phase microextraction (Liu et al. 2007). Like other phytohormones, it was initially determined by bioassays based on its physiological properties (Sembdner et al. 1988). The naturally occurring form is S-(+)-ABA, and the side chain of ABA is in 2-cis, 4-trans configuration by definition (Addicott et al. 1968). Due to this optical property, ABA was also previously determined by polarimetry (Cornforth et al. 1966). However, specific rotation is often influenced by numerous other substances in plant extracts, thus such determination is very inaccurate. The compound also strongly absorbs ultraviolet (UV) radiation, maximally at about 260 nm (due to the presence of chromophores, chemical groups capable of absorbing light, resulting in the colouration of organic compounds), which allows its detection in HPLC eluates by monitoring their UV absorption (Ciha et al. 1977; Cargile et al. 1979; Mapelli and Rocchi 1983). HPLC has also been used to determine two metabolites of ABA: phaseic acid (PA) and dihydrophaseic acid (DPA) (Durley et al. 1982; Hirai and Koshimizu 1983).
Fig. 4

Structures of stress-related plant hormones

Immunological methods such as radioimmunoassays (RIAs) (Weiler 1979, 1980; Walton et al. 1979; Mertens et al. 1983) and enzyme-linked immunosorbent assays (ELISAs) based on competitive binding between free and alkaline phosphatase-labelled ABA (Daie and Wyse 1982; Weiler 1982) have also been successfully used for estimating ABA levels and are still highly recommended for estimating free ABA levels in plant tissues. A method based on immunoaffinity chromatography (IAC) in combination with LC–MS has also been recently described (Hradecká et al. 2007). Early analytical methods for measuring levels of ABA metabolites employed GC coupled to electron capture detection (GC–ECD; Harrison and Walton 1975; Zeevaart and Milborrow 1976) or flame ionisation detection (FID; Watts et al. 1983) systems. These methods were capable of quantifying PA, DPA, epi-DPA and ABA glucose ester (ABAGE) in plant tissues at levels of about ng/g FW. Boyer and Zeevaart (1982) also developed a method for measuring ABAGE, as its tetraacetate derivative, by GC–ECD. In addition, several methods for quantifying ABA and ABAGE by GC–MS in selected ion monitoring (SIM) mode (Netting et al. 1982; Duffield and Netting 2001) and multiple ion monitoring (MIM) mode (Neill et al. 1983) have been published. However, GC–MS is limited to the analysis of volatile compounds, thus methylation of these analytes with diazomethane prior to the analysis is required. Regarding detection techniques following GC separation, ECD permits quantitative analyses of ABA in much smaller samples of plant material than FID. When GC coupled with MS in SIM mode, even much higher sensitivity is then achieved. Further, methods for determining ABA by CE have been published (Liu et al. 2002, 2003). CE has advantages for analysing ABA (as a trace substance in complex plant extracts), but it suffers from low sensitivity in combination with UV detection. This problem can be overcome using either micellar electrokinetic capillary chromatography (MECC; Liu et al. 2002) or laser-induced fluorescence (LIF) detection, both of which provide high sensitivity, but again require derivatisation because ABA is not fluorescent. Therefore, in the second cited study by Liu et al. (2003), ABA was labelled with 8-aminopyrene-1,3,6-trisulfonate via reductive amination in the presence of acetic acid and sodium cyanoborohydride. The resulting conjugate was quantified, with fmol detection limits, and the method was used to analyse ABA in crude tobacco extracts. Recently, LC coupled to MS with soft ionisation techniques (ESI, APCI) has proved to be very powerful for analysing substances in plant extracts since they are often polar, non-volatile, thermally labile and (hence) inappropriate for GC analysis. Due to its high selectivity and sensitivity, LC–MS in multiple reaction monitoring mode (MRM) has also become increasingly popular for analysing ABA and its metabolites (Gómez-Cadenas et al. 2002; López-Carbonell and Jáuregui 2005; Chiwocha et al. 2007; López-Carbonell et al. 2009). Another technique that has been used for quantifying ABA and ABAGE is LC–MS in SIM mode, either directly (Hogge et al. 1993; Schneider et al. 1997) or following several purification steps (Vilaró et al. 2006). Further improvements in the analysis of ABA metabolites have been obtained through use of a UHPLC-based MS method, which is faster, affords higher throughput and is more sensitive than conventional LC–MS (Turečková et al. 2009). The detection limits of the technique were found to be at low picomolar levels for ABAGE and ABA acids in negative ion mode, and femtomolar levels for ABAGE, ABAaldehyde, ABAalcohol and the methylated acids in positive ion mode.

Ethylene

Ethylene is a flammable unsaturated gaseous hydrocarbon (Fig. 5) with a molecular weight of 28.05 g/mol. It has been indirectly used for thousands of years to ripen fruits, for instance via the ancient Egyptian practice of gashing figs (Galil 1948). It seems to have been first described by Becher (1669, Physica Subterranea), identified as a natural plant product by Gane (1934) and shown to influence plant growth and development by Crocker et al. (1935). It is formed essentially in all cells, but often most abundantly in fruits and wounded tissues, diffuses through tissues and is finally released into the surrounding atmosphere. The levels of ethylene produced by plants are low and of the same order as those of other phytohormones. Thus, sensitive methods are essential for its determination. The first methods for ethylene detection, like those for other plant hormones, were based on certain bioassays, mainly because of the lack of instrumental methods at the time. The first was the ‘triple response’ etiolated pea plant bioassay based on measurement of reductions in stem elongation (Nejlubow 1901) and several others were subsequently developed (Crocker et al. 1932; Addicott 1970; Kang and Rat 1969). However, all the bioassays lack specificity (for instance, propylene, acetylene and butylenes can induce similar responses, albeit at up to a thousand times higher concentrations than ethylene) and thus are rarely used now. The development of chromatographic (especially gas chromatography) techniques allowed the identification (by coupling to MS) and quantification of low molecular weight hydrocarbons including ethylene and its biosynthetic precursors 1-aminocyclopropane-1-carboxylic acid (ACC) and 1-(malonylamino)cyclopropane-1-carboxylic acid (MACC), for structures see Fig. 5. All of the mentioned substances of biological interest can be clearly distinguished from other low molecular weight hydrocarbons with high accuracy at approximately 10−12 m3 dm−3 in a 1 cm3 volume (ppb level) of air. The first GC method for ethylene determination was applied to measure this substance from apples (Burk and Stolwijk 1959; Huelin and Kennett 1959). The major drawback of this approach based on thermal conductivity detection (TCD) was a relatively high detection limit of 10–100 μL/L. The introduction of flame ionisation detection (FID) and the photoionisation detector (PID) in 1980s significantly improved the detection limit of ethylene to tens of nL/L levels (Bassi and Spencer 1985; Bassi and Spencer 1989). At the beginning, the ethylene sampling procedure and its subsequent injection into the GC column have been done manually with a gas-tight syringe, which was filled with gas from the headspace of a closed cuvette, in which the plant was enclosed for a few hours (Abeles et al. 1992). For low reproducibility and high time consumption of this system, it has been later replaced by automatic samplers based on concentric rotary valves (Cristescu et al. 2013). To achieve better sensitivity, GC systems can be equipped with preconcentration devices that enable to store the emitted ethylene (Segal et al. 2000). The plants are placed in closed cuvettes and continuously flushed with air. Ethylene is trapped inside a tube containing an appropriate adsorption material (e.g. carbon molecular sieve) and is then released into a smaller volume by heating the adsorbent. In addition to GC and GC–MS (Smets et al. 2003) approaches, photo acoustic laser spectrophotometry (PALS; Cristescu et al. 2008), LC–MS (Petritis et al. 2003) and CE–LIF (Liu et al. 2004) methods for ethylene (or ACC) determination have been published. PALS offers higher detection sensitivity (ppt level) than GC and is highly selective for particular substances. This is disadvantageous in some respects, as the equipment has much narrower applications than GC. However, before use of the GC–MS and CE–LIF methods, the analytes in plant extracts must be modified by derivatisation, which greatly increases time consumption, and the derivatisation procedure has poor reproducibility when concentrations of ACC are low. Recently, methodology based on in vitro measurement of the activity of two key biosynthetic enzymes, 1-aminocyclopropane-1-carboxylate synthase (ACS) and 1-aminocyclopropane-1-carboxylate oxidase (ACO), as well as ethylene itself, has been reported (Bulens et al. 2011).
Fig. 5

Structure of ethylene and its biosynthetic precursors ACC and MACC

Brassinosteroids

Brassinosteroids (BRs) are relatively young group of naturally occurring triterpenoid plant growth substances with hormonal function (Caño-Delgado et al. 2004). More than 70 BR analogues have been identified so far in nearly 60 plant species (Choe 1999). Their common structural feature is a 5α-cholestane skeleton (Fig. 6) and they are divided into different categories depending on the side chain structure and modifications of the A and B rings. Physiologically, BRs participate with other plant hormones in the regulation of numerous developmental processes, including shoot growth, root growth, vascular differentiation, fertility and seed germination (Fujioka and Sakurai 1997). BRs also have anti-stress effects, i.e. they participate in ameliorative responses to various stresses, such as low and high temperature, drought and infection. Like GAs, they tend to be relatively abundant in reproductive plant tissues, such as pollen, flowers and immature seeds, but their levels are extremely low in vegetative tissues, even compared to those of other plant hormones (fg-pg/g FW).
Fig. 6

Structure of 5α-cholestane and biologically active naturally occurring brassinosteroids

Initially, immunoassays and bioassays were mainly used for detecting BRs (Takatsuo and Yokota 1999). Some of the bioassays have good sensitivity and are still used for testing the biological activity of BRs, particularly the bean second-internode bioassay (Mitchell and Livingston 1968) and rice lamina inclination bioassay (Maeda 1965), which provide 2 × 10−11 mol and 1 × 10−13 mol detection limits, respectively (Thomson et al. 1981; Wada et al. 1984). Immunological methods such as RIAs and ELISAs have also been used for exploring the distribution of BRs in plant tissues (Horgen et al. 1984; Yokota et al. 1990). RIA was found to be useful for detecting the two most common bioactive BRs, castasterone (CS) and brassinolide (BL) (Fig. 6), with approximately 0.3 pmol detection limits. However, ELISA based on a mouse monoclonal antibody against 24-epibrassinolide (epiBL) was shown to respond not only to epiBL but also to other, non-BR phytosterols (sitosterol, ecdysone). So, this method could not be used for analysing BRs. Swaczynová et al. (2007) subsequently improved the ELISA method using selective polyclonal antibodies against 24-epicastasterone (epiCS) and successfully detected this substance in Brassica napus and Arabidopsis tissues. These antibodies cross-reacted with BL and epiBL, but not with non-BR plant sterols. Thus, the method was applied for determining BR levels in extracts of tissues from several plant species. In addition, good agreement was found between results obtained using the ELISA method and a simultaneously developed HPLC–MS approach.

Several hyphenated (GC–MS and LC–MS) techniques were also gradually introduced. Since BRs are not volatile they must be derivatised prior to GC–MS analysis (Takatsuo et al. 1982). The standard derivatisation procedure is based on formation of bis-methaneboronates (BMBs) of BRs with vicinal diol groups (e.g. BL and CS). Thus, a disadvantage of this approach is that it cannot be used to analyse BRs lacking this conformation (e.g. teasterone and typhasterol). The detection limits of BMB derivatives are at the sub-ng level. GC–MS has also been used to elucidate structures of new BRs and BR biosynthesis pathways (Fujioka and Sakurai 1997). Liquid chromatography is generally suitable for non-volatile compounds, therefore, BRs can be advantageously analysed using this technique without derivatisation. However, although several LC methods have been published, only one can be used for direct determination of free BRs (Swaczynová et al. 2007). The others still require derivatisation. Gamoh et al. (1996) developed a method based on preparation of naphthaleneboronates, which is also applicable to teasterone and typhasterol (unlike BMB derivatisation). It has a reported detection limit of 2 ng and was applied to analyse BRs in Cannabis sativa seeds. Another LC method provides 125 attomole detection limits for dansyl-3-aminophenylboronate derivatives of BRs in highly laboriously purified extracts (57 g) of 24-day-old Arabidopsis plants grown in vitro. (Svatoš et al. 2004). Recently, two other LC–MS methods for preparing and analysing boronate derivatives of BRs have been reported (Huo et al. 2012; Ding et al. 2013), but the starting amount of tissue (Arabidopsis) used in the cited studies was still extremely high: 1 or 2 g FW.

Jasmonates

Jasmonic acid (JA) and its metabolites, collectively called jasmonates (JAs), are cyclopentanone compounds (Fig. 4) that share remarkable structural and functional properties with prostaglandins found in animals (Wasternack and Kombrink 2010). In the 1990s, JAs were proposed to be stress-related compounds (Farmer and Ryan 1990; Parthier 1991) that accumulate in plants in response to various stresses, such as wounding or pathogen attack (Creelman et al. 1992), in plant tissues or cell cultures treated with fungal elicitors (Müller et al. 1993), and tissues subjected to abiotic stressors such as UV radiation, low and high temperatures, osmotic stress and ozone exposure (Parthier et al. 1992). JAs seem to occur in most organs of most plant species (Wasternack and Hause 2013). Their in planta concentrations, which can be determined by various methods, are comparable to those of other plant hormones, ranging from ng to μg/g FW, depending on the plant tissue, species, developmental stage and both environmental and physiological conditions (Wilbert et al. 1998). The major physiologically active jasmonates are reportedly (−)-JA, methyl jasmonate (MeJA), and conjugates of (−)-JA with the amino acids isoleucine (JA-Ile), valine (JA-Val), and leucine (JA-Leu) (Sembdner and Parthier 1993). JA-amino acid conjugates are constitutively produced in plant tissues and their levels increase upon osmotic stress (Kramell et al. 1995). In plant–herbivore interactions, JA-amino acid conjugation is necessary for JA activation, and (−)-JA-Ile is the bioactive form of the hormone (Staswick and Tiryaki 2004; Fonseca et al. 2009). MeJA plays important roles as a fragrant volatile compound, particularly in plant–plant interactions, in which it acts, in concert with other volatile substances emitted from the plants, as an aerial signal for communication with their environment (Pichersky and Gershenzon 2002). Its synthesis is induced by various external stimuli, such as adverse weather conditions, and attacks by herbivores or pathogens (Paré and Tumlinson 1999). Another JA metabolite, which is highly active in plant–insect interactions, is cis-jasmone (Birkett et al. 2000; Bruce et al. 2008).

Key considerations in the analysis of jasmonates are that JA and its conjugates are non-volatile, while MeJA and cis-jasmone are volatile. GC–MS is the most frequently used approach for quantifying JA, but as JA is not volatile it must first be derivatised, for instance by preparation of pentafluorobenzyl ester derivatives (Müller and Brodschelm 1994). This provides high sensitivity, but requires an elaborate preconcentration procedure. Another GC–MS-based technique for JA quantification has been described (Engelberth et al. 2003), in which the only purification step is collection of derivatised JA on a polymeric adsorbent. Nevertheless, these time-consuming steps still seriously limit the number of samples that can be processed per day. MeJA can be successfully quantified directly by GC with FID or MS detection after concentration by solid-phase microextraction (SPME) on fused silica fibre coated with a polymeric sorbent (Meyer et al. 1984). The reported detection limit of this method is 1 ng/injection, sufficiently low for detecting MeJA in plant tissues at levels between 10 and 100 ng/g DW (Müller and Brodschelm 1994; Wilbert et al. 1998).

Due to the polarity and non-volatility of JA most researchers use LC-based methods for its analysis. Anderson (1985) described an HPLC assay for the simultaneous determination of ABA and JA in plant extracts, following derivatisation (of both growth regulators) with a fluorescent hydrazide to obtain stable fluorescent products—dansyl hydrazones. This procedure allows detection of both hormones at low pmol levels. The method was demonstrated using extracts of several different tissues of soybean (Glycine max), snap beans (Phaseolus vulgaris), lima beans (Phaseolus lunatus) and broccoli (Brassica oleracea). However, only 20 % of the JA was converted to the corresponding ester during the derivatisation procedure. Fluorescent labelling usually affords great sensitivity for detecting the resulting derivatives (approx. 10−17 mol) and thus prompted other researchers to optimise this kind of derivatisation to introduce fluorophores into the chemical structure of JA to monitor it after separation by either HPLC (Kristl et al. 2005; Xiong et al. 2012) or CE (Zhang et al. 2005). However, the highest selectivity and sensitivity for JA determination can be currently achieved using MS/MS in MRM mode (Tamogami and Kodama 1998; Wilbert et al. 1998; Segarra et al. 2006). Another of JA’s key physicochemical properties is amenability to oxidation, which was recently exploited for its electrochemical detection following LC (Xie et al. 2012). The method was successfully applied to analyse endogenous JA in wintersweet flowers and rice florets with a detection limit of 10−8 mol/L. To study physiological process in plants under various stresses, many researchers also monitor levels of precursors in the JA biosynthetic pathway (especially 12-oxo-phytodienoic acid, OPDA) and JA metabolites. For these studies, LC–MS/MS is the method of choice (Radhika et al. 2012; VanDoorn et al. 2011). Similarly, stress resistance investigators (profiling the main stress response actors such as JAs, ABA and salicylic acid) generally use LC–MS/MS methods, which have been accelerated by coupling UHPLC, rather than conventional HPLC, systems to tandem mass spectrometers (Flors et al. 2008; López-Ráez et al. 2010; Balcke et al. 2012). This strategy also increases sensitivity, allowing successful quantification of target stress hormones in milligram quantities of plant tissue samples.

Salicylic acid

Salicylic acid (SA; Fig. 4) is an endogenous signalling molecule that is predominantly active in plant immune responses to avirulent pathogens, but like other phytohormones it is also involved in the regulation of several developmental processes, especially flowering (Singh et al. 2013). In most plants, pathogen attack, insect feeding and other kinds of physical wounding trigger both local and systemic resistance, mediated by the accumulation of defence-related proteins at sites of infection/damage and healthy tissues, respectively (Hammond-Kosack and Jones 2000). On the molecular level, accumulation of SA in cells leads to the release of NPR1 protein, activation of TGA1 and TGA2 transcription factors and expression of pathogenesis-related proteins (Pieterse and Van Loon 2004). To elucidate their signalling roles, SA and its metabolites (including methylsalicylate, salicylic acid glucoside and salicylic acid glucosylester) must be precisely quantified in plant tissues by robust, sensitive analytical methods. HPLC with fluorescence detection has been successfully used for quantifying SA, following a complex purification procedure (Meuwly and Metraux 1993), in cucumber (Cucumis sativus L.) seedlings infected by Pseudomonas lachrymans on the first leaf. Free SA contents increased locally in the infected leaf and systemically in the second leaf to 33-fold and 4.2-fold higher than detection limits, respectively, while remaining undetectable in controls. Recently, another HPLC method with fluorescence detection has been reported for quantitative analysis of SA in tobacco leaf tissues (Verberne et al. 2002). This methodology increased SA extraction recovery from plant tissues by reducing SA sublimation during purification via the addition of a small amount of HPLC eluent, resulting in recoveries in the range of 71–91 % for free SA and 65–79 % for acid-hydrolysed SA.

However, fluorometric analysis cannot fully distinguish SA and its metabolites from other plant components, particularly simple phenolics and phenylpropanoids, which might be present in infected plants and participate in disease resistance. Partly for this reason, methods allowing their precise, accurate determination based on analyses of their molecular masses and specific daughter fragments (tandem MS) or other analyte-specific approaches have been developed. Initially, a method based on electrospray tandem MS coupled to capillary LC was introduced to detect SA (together with JA and MeJA) in extracts of fresh poplar leaves (Wilbert et al. 1998). In addition, a bacterial biosensor that is highly specific for SA, methyl-SA and the synthetic SA derivative acetylsalicylic acid was recently shown to be suitable for quantifying SA in crude plant extracts (Huang et al. 2005). Following increases in its throughput, this approach was successfully applied for genetic screenings for SA metabolic mutants and characterising enzymes involved in SA metabolism (Defraia et al. 2008; Marek et al. 2010). Another LC–MS/MS approach has been applied to study SA and JA levels in cucumber cotyledons under biotic stress induced by the necrotrophic pathogen Rhizoctonia solani (Segarra et al. 2006). An LC–MS method has also been compared to a capillary electrophoresis (CE) technique, and used to study SA and related phenolics in wild-type Arabidopsis plants and two lines with mutations affecting SA accumulation in response to two avirulent bacterial strains (Shapiro and Gutsche 2003). Furthermore, in efforts to elucidate SA metabolism Pastor et al. (2012) developed an LC–MS/MS method that enabled them to identify two conjugates: salicylic glucosyl ester (SGE) and glucosyl salicylate (SAG). Their results also revealed that SA and its main glucosyl conjugates accumulate in Arabidopsis thaliana in a time-dependent manner, in accordance with the up-regulation of SA-dependent defences following Pseudomonassyringae infection. In addition to SA signalling, Belles et al. (1999) found that gentisic acid, a product of SA hydroxylation, is a complementary pathogen-inducible signal that is essential for accumulation of several antifungal pathogenesis-related proteins in tomato. Both gentisic and salicylic acids were quantified in SA-treated chamomile by a rapid UPLC-MS/MS method originally developed for analysing hydroxybenzoates and hydroxycinnamates in beverages (Gruz et al. 2008; Kovacik et al. 2009). SA was accurately quantified using deuterium-labelled internal standards of salicylic (3,4,5,6-[2H4]) and 4-hydroxybenzoic (2,3,5,6-[2H4]) acids to account for ESI–MS signal trends, matrix effects and potential extraction losses.

GC–MS has also been used to quantify SA, after derivatisation (to enhance volatility and sensitivity) of the carboxylic acid with diazomethane to form SA methylester (Scott and Yamamoto 1994). The disadvantage of this method is that it requires elaborate sample preparation, from ca. 1 g FW of tissue, including anion exchange and preparative HPLC. In 2003, Engelberth and co-workers introduced a method for SA and JA analysis based on collecting derivatised and volatilised compounds on a polymeric adsorbent and GC-positive ion chemical ionisation-MS using only milligrams of plant tissue. This method was later used to explore changes in the metabolic profiles of SA, cinnamic acid, JA, IAA, ABA, unsaturated C18 fatty acids, 12-oxo-phytodienoic acid, and the phytotoxin coronatine in Arabidopsis infected by P. syringae (Schmelz et al. 2004). However, this approach also requires both purification and derivatisation steps.

Strigolactones

Strigolactones (SLs) are the most recently described class of plant hormones. They were originally regarded as a family of carotenoid-derived plant secondary metabolites (Xie et al. 2010) with roles in signalling between organisms (allelochemicals). Initially, they were of interest due to their action as seed germination stimulants of the root parasitic allelopathic weeds Orobanche and Striga (Cook et al. 1966, 1972). Seeds of these parasites germinate only when they perceive chemical signals produced by and released from the roots of other (host or nonhost) plants. This germination-inducing activity of SLs was the basis of a sensitive SL bioassay using Striga and Orobanche seeds (Joel et al. 1995), and very recently a fast convenient method for determining the germination rate of parasitic weeds seeds has been reported (Pouvreau et al. 2013). In 2008, SLs were classified as a new group of plant signalling molecules with endogenous hormonal activity due to involvement in the inhibition of shoot branching (Gomez-Roldan et al. 2008; Umehara et al. 2008). Earlier it was also found that they are able to stimulate hyphal branching in mycorrhizal fungi (Akiyama et al. 2005). To date, more than 20 compounds in this family of sesquiterpene lactones have been identified from root exudates of various plant species (Tsuchiya and McCourt 2009; Xie et al. 2010). The most well-known SLs are strigol and orobanchol (Fig. 7). Structurally, all natural SLs contain a tricyclic lactone skeleton (cycles A, B and C) bound to a butenolide moiety (ring D) via an enol ether bond. The A and B rings bear various substituents (Zwanenburg et al. 2009).
Fig. 7

Natural strigolactones—basic structure and the examples

A pioneering GC–MS approach for analysing SLs in plant samples was introduced by Yokota et al. (1998). However, this method required time-consuming purification of extracts and TMS derivatisation. Furthermore, some SLs partially decomposed during the procedures. An approach for analysing strigol heptafluorobutyrate derivatives obtained from extracts of Striga asiatica host plants (maize and proso millet) was subsequently developed (Siame et al. 1993), based on electron impact (EI) and both positive and negative chemical ionisation (CI) mass analyses of HPLC fractionated samples. This method avoided some of the previous problems, but the purification procedure was still extremely tedious and labour intensive. Another two approaches for analysing SLs, based on LC–MS/MS, have been published, with detection limits (in MRM mode) ranging between 0.1 and 1 pg/μL of SLs (Sato et al. 2003, 2005). Most recently, Xie et al. (2013) reported an LC–MS/MS procedure allowing structural elucidation of 11 SLs in root exudates of tobacco and detection of four SLs in rice. Notably, no SL profiling has been reported to date.

Hormone profiling

Since it is becoming increasingly evident that hormones do not act separately, but have highly interactive physiological effects and mutually affect each other’s biosynthesis and metabolism, there is increasing interest in ‘hormone profiling’, i.e. analysing hormones of several classes (together with their precursors and metabolites) in the same plant tissue simultaneously. This greatly increases analytical complexity as it requires methodology capable of quantitatively detecting chemically and structurally diverse substances rather than a single targeted group of plant hormones. Since many plant hormones are acidic, published methods have often focused on these classes of compounds (Müller et al. 2002; Schmelz et al. 2003; Durgbanshi et al. 2005). Hormone profiling was first successfully applied, by Chiwocha et al. (2003), in a study of thermodormancy where four CKs and 10 acidic plant hormones (IAA, ABA, ABAGE, 7′OH-ABA, PA, DPA and four GAs) in 50–100 mg DW extracts of lettuce seeds were all measured by LC–MS, using a single purification step and a 40 min chromatographic gradient. Pan et al. (2008) subsequently developed an LC–MS/MS technique (requiring no purification or derivatisation) for simultaneous quantification of 17 plant hormones including auxins, CKs, ABA, GAs, JAs, salicylates and corresponding methyl esters in crude extracts of samples (50–100 mg) of Arabidopsis plants that had been mechanically wounded or challenged with the fungal pathogen Botrytis cinerea. Limits of quantification reportedly ranged from 0.01 to 10 pg/g FW. In addition, a fast LC–MS/MS method combined with an automatic liquid handling system for SPE was recently used for simultaneous analysis of 43 plant hormone substances, including CKs, auxins, ABA and GAs in the rice GA-signalling mutants gid1-3, gid2-1 and slr1 to study relationships between changes in gene expression and hormone metabolism (Kojima et al. 2009). To enhance sensitivity, a nanoflow-LC–MS/MS approach has also been used to detect 14 plant hormones, following a two-step purification procedure, in extracts of Arabidopsis and tobacco seedlings (Izumi et al. 2009). The limit of detection was found to be in the sub-fmol range for most studied analytes. However, this method failed for the acidic plant hormones, especially GAs. Capillary electrophoresis with laser-induced fluorescence detection (CE–LIF) has also been applied in profiling, to simultaneously determine plant hormones containing carboxyl groups, including GA3, IAA, indole butyric acid (IBA), 1-naphthalene acetic acid (NAA), 2,4-dichloro-phenoxy acetic acid (2,4-D), ABA and JA in crude extracts of banana samples (500 mg) without further purification (Chen et al. 2011). Finally (for this summary), 20 plant growth substances (including IAA, ABA, CKs and structurally related purines) have been determined in single chromatographic runs (Farrow and Emery 2012). The methodology involved extraction from 100 mg samples of Arabidopsis thaliana leaves, purification and analysis by conventional HPLC with a fused core column. QTRAP mass analyzer has been utilised here for detection of selected analytes. Reported detection limits ranged from 2 pmol for zeatin-9-glucoside to 750 pmol for IAA. High-resolution MS (HR-MS) is not yet widely used for quantitative analysis of plant hormones, but will probably soon be employed for their routine quantification. An HR-MS approach has already been applied to identify and quantify a large number of endogenous phytohormones in tomato fruits and leaf tissues (Van Meulebroek et al. 2012). The cited authors selected eight phytohormones—GA3, IAA, ABA, JA, SA, zeatin (not mentioned if trans- or cis-), BAP and epiBL—as representatives of the major hormonal classes, and applied a simple extraction procedure followed by UHPLC-Fourier Transform Orbitrap MS separation and detection for hormonally profiling 100 mg FW samples of the tomato tissues. The samples were extracted in methanol:water:formic acid (75:20:5) over 12 h, ultra-centrifuged, then injected directly into an UHPLC system equipped with C18 column of 2.1 × 50 mm diameter (particle size 1.8 μm) and coupled to a benchtop Orbitrap mass spectrometer, equipped with a heated electrospray ionisation source (HESI), operating in both positive and negative modes. This technique provided detection limits of the analytes ranging from 0.05 to 0.42 pg/μL. Moreover, full mass scans by the Orbitrap MS provided a dataset including information on hundreds of matrix compounds. Therefore, this metabolomic profiling approach might lead to the discovery of compounds with no previously known hormonal activities or roles in plants. However, although HR-MS provides opportunities to use narrow mass windows to exclude interfering matrix compounds and selectively analyse substances (Kaufmann 2012), Van Meulebroek et al. (2012) used a relatively broad mass window of 5 ppm, to ensure that no compounds would be completely undetected.

Conclusion and perspectives

Analysis of plant hormones is very challenging because they have extremely wide ranges of physicochemical properties, and plant tissues contain trace quantities of hormones together with thousands of other substances at far higher levels. However, there have been great advances in analytical techniques used in diverse “life sciences” during the last decade. LC–MS has become the most versatile, rapid, selective and sensitive technique available for identifying and quantifying small molecules (Pan et al. 2008; Van Meulebroek et al. 2012). Thus, it is replacing all other approaches in plant hormone analysis. New technologies based on a unique ion transfer device designed to maximise ion transmission from the source to the mass analyzer, could further improve the sensitivity (typically the primary concern) of phytohormone measurements. Typical gains obtained using such device include generally 25-fold increases in peak areas and 10-fold increases in signal-to-noise ratios, which are highly valuable for phytohormone quantifications. The next challenge could be to develop robust techniques for extending the breadth of profiling, including more phytohormone precursors and metabolites, as well as those of other signalling molecules in plants. LC–MS/MS methods may be particularly useful for this as they afford capabilities for simultaneously quantifying metabolites with diverse properties of both single and multiple phytohormone groups.

Even with further anticipated advances it will be extremely challenging to quantify all phytohormones and related compounds in a single LC–MS/MS run due to their high chemical diversity and the inherent difficulty in distinguishing numerous metabolites that may have very similar chromatographic properties, share the same mass and yield very similar fragments. The extremely high levels of matrix compounds typically present in plant extracts compound the problem. However, additional orthogonal separation techniques have been recently introduced that provide high selectivity, further improve spectral quality, enhance the quality of acquired datasets and facilitate their interpretation, thus surmounting some of the difficulties. Notably, ion mobility separation (Eugster et al. 2012) and MSE have been combined in a powerful new approach called high definition mass spectrometry (Sotelo-Silveira et al. 2013), which could generate more precise datasets from explorations of endogenous phytohormone levels and their changes during developmental processes in plants. The recent progress in analytical MS technologies could also enable tissue- and cell-specific quantifications as well as analyses of levels of multiple hormones in single plant cells or subcellular compartments.

Notes

Acknowledgments

Financial support from the Ministry of Education, Youth and Sport of the Czech Republic through National Program of Sustainability, Grant no. LO 1204 is gratefully acknowledged. The authors would like to also express thanks to Sees-editing Ltd. for critical reading and editing of the manuscript.

References

  1. Abeles FB, Morgan PW, Saltveit MEJ (1992) Ethylene in plant biology. Academic Press, San DiegoGoogle Scholar
  2. Addicott FT, Lyon JL, Ohkuma K, Thiessen WE, Carns HR, Smith OE, Cornforth JW, Milborrow BV, Ryback G, Wareing PF (1968) Abscisic acid: a new name for abscisin II (dormin). Science 159:1493PubMedGoogle Scholar
  3. Addicott FT (1970) Plant hormones in the control of abscission. Biol Rev 45:485–524Google Scholar
  4. Akiyama K, Matsuzaki K, Hayashi H (2005) Plant sesquiterpenes induce hyphal branching in arbuscular mycorrhizal fungi. Nature 435:824–827PubMedGoogle Scholar
  5. Anderson JM (1985) Simultaneous determination of abscisic acid and jasmonic acid in plant extracts using high performance liquid chromatography. J Chromatogr A 330:347–355Google Scholar
  6. Åstot C, Doležal K, Moritz T, Sandberg G (1998) Precolumn derivatization and capillary liquid chromatographic/frit-fast atom bombardment mass spectrometric analysis of cytokinins in Arabidopsis thaliana. J Mass Spectrom 33:892–902PubMedGoogle Scholar
  7. Ayele BT, Magnus V, Mihaljević S, Prebeg T, Čož-Rakovac R, Ozga JA, Reinecke DM, Mander LN, Kamiya Y, Yamaguchi S, Salopek-Sondi B (2010) Endogenous gibberellin profile during Christmas rose (Helleborus niger L.) flower and fruit development. J Plant Growth Regul 29:194–209Google Scholar
  8. Bai Y, Zhang J, Bai Y, Liu H (2012) Direct analysis in real time mass spectrometry combined with single-drop liquid-liquid-liquid microextraction for the rapid analysis of multiple phytohormones in fruit juice. Anal Bioanal Chem 403:2307–2314PubMedGoogle Scholar
  9. Balcke GU, Handrick V, Bergau N, Fichtner M, Henning A, Stellmach H, Tissier A, Hause B, Frolov A (2012) An UPLC–MS/MS method for highly sensitive high-throughput analysis of phytohormones in plant tissues. Plant methods 8:47PubMedCentralPubMedGoogle Scholar
  10. Barkawi LS, Tam Y-Y, Tillman JA, Pederson B, Calio J, Al-Amier H, Emerick M, Normanly J, Cohen JD (2008) A high-throughput method for the quantitative analysis of indole-3-acetic acid and other auxins from plant tissue. Anal Biochem 372:177–188PubMedGoogle Scholar
  11. Barkawi LS, Tam Y-Y, Tillman JA, Normanly J, Cohen JD (2010) A high-throughput method for the quantitative analysis of auxins. Nat Protoc 5:1609–1618PubMedGoogle Scholar
  12. Bassi PK, Spencer MS (1985) Comparative evaluation of photo-ionisation and flame ionisation detectors for ethylene analysis. Plant Cell Environ 8:161–165Google Scholar
  13. Bassi PK, Spencer MS (1989) Methods for the quantification of ethylene produced by plants. In: Linskens HF, Jacksons JF (eds) Gases in plant and microbial cells. Modern methods of plant analysis, vol 9. Springer, Berlin, pp 309–321Google Scholar
  14. Becher JJ (1669) Physica Subterranea. In: Brown JC (ed) (2006) A history of chemistry: from the earliest times till the present day. Kessinger, pp 225. ISBN 978-1-4286-3831-0Google Scholar
  15. Belles JM, Garro R, Fayos J, Navarro P, Primo J, Conejero V (1999) Gentisic acid as a pathogen-inducible signal, additional to salicylic acid for activation of plant defenses in tomato. Mol Plant Microbe 12:227–235Google Scholar
  16. Béres T, Gemrotová M, Tarkowski P, Ganzera M, Maier V, Friedecký D, Dessoyf MA, Wessjohannf LA, Spíchal L, Strnad M, Doležal K (2012) Analysis of cytokinin nucleotides by capillary zone electrophoresis with diode array and mass spectrometric detection in a recombinant enzyme in vitro reaction. Anal Chim Acta 751:176–181PubMedGoogle Scholar
  17. Binks R, MacMillan J, Pryce RJ (1969) Plant hormones-VIII: combined gas chromatography-mass spectrometry of the methyl esters of gibberellins A1 to As and their trimethylsilyl ethers. Phytochemistry 8:271–284Google Scholar
  18. Birkett MA, Campbell CAM, Chamberlain K, Guerrieri E, Hick AJ, Martin JL, Matthes M, Napier JA, Pettersson J, Pickett JA, Poppy GM, Pow EM, Pye BJ, Smart LE, Wadhams GH, Wadhams LJ, Woodcock CM (2000) New roles for cis-jasmone as an insect semiochemical and in plant defense. Proc Natl Acad Sci USA 97:9329–9334PubMedCentralPubMedGoogle Scholar
  19. Boyer GL, Zeevaart JAD (1982) Isolation and quantitation of β-d-glucopyranosyl abscisate from leaves of Xanthium and spinach. Plant Physiol 70:227–231PubMedCentralPubMedGoogle Scholar
  20. Bruce TJA, Matthes MC, Chamberlain K, Woodcock CM, Mohib A, Webster B, Smart LE, Birkett MA, Pickett JA, Napier JA (2008) cis-Jasmone induces Arabidopsis genes that affect the chemical ecology of multitrophic interactions with aphids and their parasitoids. Proc Natl Acad Sci USA 105:4553–4558PubMedCentralPubMedGoogle Scholar
  21. Bulens I, Van de Poel B, Hertog ML, De Proft MP, Geeraerd AH, Nicolai BM (2011) Protocol: an updated integrated methodology for analysis of metabolites and enzyme activities of ethylene biosynthesis. Plant Methods 7:17PubMedCentralPubMedGoogle Scholar
  22. Burk SP, Stolwijk JAJ (1959) A highly sensitive katharometer and its application to the measurement of ethylene and other gases of biological importance. J Biochem Microb Technol Eng 1:245–259Google Scholar
  23. Campell BR, Town CD (1991) Physiology of hormone autonomous tissue lines derived from radiation-induced tumors of Arabidopsis thaliana. Plant Physiol 97:1166–1173PubMedCentralPubMedGoogle Scholar
  24. Caño-Delgado A, Yin Y, Yu C, Vafeados D, Mora-García S, Cheng J-C, Nam KH, Li J, Chory J (2004) BRL1 and BRL3 are novel brassinosteroid receptors that function in vascular differentiation in Arabidopsis. Development 131:5341–5351PubMedGoogle Scholar
  25. Cargile NL, Borchert R, McChesney JD (1979) Analysis of abscisic acid by high-performance liquid chromatography. Anal Biochem 97:331–339PubMedGoogle Scholar
  26. Chen CM (1981) Biosynthesis and enzymatic regulation of the interconversion of cytokinin. In: Guern J, Péaud-Lenoël C (eds) Metabolism and molecular activities of cytokinins. Springer, Berlin, pp 34–43Google Scholar
  27. Chen H, Guo X-F, Zhang H-S, Wang H (2011) Simultaneous determination of phytohormones containing carboxyl in crude extracts of fruit samples based on chemical derivatization by capillary electrophoresis with laser-induced fluorescence detection. J Chromatogr B 879:1802–1808Google Scholar
  28. Chiwocha SDS, Abrams S, Ambrose SJ, Cutler AJ, Loewen M, Ross ARS, Kermode AR (2003) A method for profiling classes of plant hormones and their metabolites using liquid chromatography-electrospray ionisation tandem mass spectrometry: an analysis of hormone regulation of thermodormancy of lettuce (Lactuca sativa L.) seeds. Plant J 35:405–417PubMedGoogle Scholar
  29. Chiwocha SDS, Rouault G, Abrams S, von Aderkas P (2007) Parasitism of seed of Douglas-fir (Pseudotsuga menziesii) by the seed chalcid, Megastigmus spermotrophus, and its influence on seed hormone physiology. Sex Plant Reprod 20:19–25Google Scholar
  30. Chen K-H, Miller AN, Patterson GW, Cohen JD (1988) A rapid and simple procedure for purification of indole-3-acetic acid prior to GC–SIM–MS analysis. Plant Physiol 86:822–825PubMedCentralPubMedGoogle Scholar
  31. Choe S (1999) Biosynthesis, signal transduction, action! In: Davies PJ (ed) Plant hormones. Kluwer Academic Publishers, Netherlands, pp 156–178Google Scholar
  32. Chory J, Reinecke D, Sim S, Washburn T, Brenner M (1994) A role for cytokinins in de-etiolation in Arabidopsis. Det mutants have an altered response to cytokinins. Plant Physiol 104:339–347PubMedCentralPubMedGoogle Scholar
  33. Ciha AJ, Brenner ML, Brun WA (1977) Rapid separation and quantification of abscisic acid from plant tissues using high performance liquid chromatography. Plant Physiol 59:821–826PubMedCentralPubMedGoogle Scholar
  34. Cohen JD, Bausher MG, Bialek K, Buta JG, Gocal GFW, Janzen LM, Pharis RP, Reed AN, Slovin JP (1987) Comparison of a commercial ELISA assay for indole-3-acetic acid at severa1 stages of purification and analysis by gas chromatography-mass spectrometry-selected ion monitoring-mass spectrometry using a 13C6-labeled internal standard. Plant Physiol 84:982–986PubMedCentralPubMedGoogle Scholar
  35. Cook CE, Whichard LP, Turner B, Wall ME, Egley GH (1966) Germination of witchweed (Striga lutea Lour.): isolation and properties of a potent stimulant. Science 154:1189–1190PubMedGoogle Scholar
  36. Cook CE, Whichard Leona P, Wall M, Egley Grant H, Coggon P, Luhan PA, McPhail AT (1972) Germination stimulants. II. Structure of strigol, a potent seed germination stimulant for witchweed (Striga lutea). J Am Chem Soc 17:6198–6199Google Scholar
  37. Cornforth JW, Milborow BV, Ryback G, Wareing PF (1965) Identity of sycamore ‘dormin’ with abscisin II. Nature 205:1269–1270Google Scholar
  38. Cornforth JW, Milborow BV, Ryback G (1966) Identification and estimation of (+) abscisin II (‘dormin’) in plant extracts by spectropolarimetry. Nature 210:627–628Google Scholar
  39. Creelman RA, Tierney ML, Mullet JE (1992) Jasmonic acid/methyl jasmonate accumulate in wounded soybean hypocotyls and modulate wound gene expression. Proc Natl Acad Sci USA 89:4938–4941PubMedCentralPubMedGoogle Scholar
  40. Cristescu SM, Persijn ST, Te Lintel Hekkert S, Harren FJM (2008) Laser-based systems for trace gas detection in life sciences. Appl Phys 92:343–349Google Scholar
  41. Cristescu SM, Mandon J, Arslanov D, De Pessemier J, Hermans C, Harren FJM (2013) Current methods for detecting ethylene in plants. Ann Bot 111:347–360PubMedCentralPubMedGoogle Scholar
  42. Crocker W, Zimmerman PW, Hitchcock AE (1932) Ethylene-produced epinasty of leaves and the relation of gravity to it. Contrib Boyce Thompson Inst 4:177–218Google Scholar
  43. Crocker W, Hitchcock AE, Zimmerman PW (1935) Similarities in the effects of ethylene and the plant auxins. Contrib Boyce Thompson Inst 7:231–248Google Scholar
  44. Croker SJ, Gaskin P, Hedden P, MacMillan J, MacNeil KAG (1994) Gibberellin biosynthesis in maize. Metabolic studies with GA15, GA24, GA25, GA7, and 2,3-dehydro-GA9. Phytochem Anal 5:74–78Google Scholar
  45. Crozier A, Loeferski K, Zaerr JB, Morris BO (1980) Analysis of picogram quantities of indole-3-acetic acid by high-performance liquid chromatography fluorescence procedures. Planta 150:366–370PubMedGoogle Scholar
  46. Daie J, Wyse R (1982) Adaptation of the enzyme-linked immunosorbent assay (ELISA) to the quantitative analysis of abscisic acid. Anal Biochem 119:365–371PubMedGoogle Scholar
  47. Darwin C, Darwin F (1880) The power of movement in plants. John Murray, LondonGoogle Scholar
  48. Defraia CT, Schmelz EA, Mou Z (2008) A rapid biosensor-based method for quantification of free and glucose-conjugated salicylic acid. Plant Methods 4:28PubMedCentralPubMedGoogle Scholar
  49. Dewitte W, Chiappetta A, Azmi A, Witters E, Strnad M, Rembur J, Noin M, Chriqui D, Van Onckelen H (1999) Dynamics of cytokinins in apical shoot meristems of a day-neutral tobacco during floral transition and flower formation. Plant Physiol 119:1111–1122Google Scholar
  50. Ding J, Mao L-J, Yuan B-F, Feng Y-Q (2013) A selective pretreatment method for determination of endogenous active brassinosteroids in plant tissues: double layered solid phase extraction combined with boronate affinity polymer monolith microextraction. Plant Methods 9:13PubMedCentralPubMedGoogle Scholar
  51. Dobra J, Motyka V, Dobrev P, Malbeck J, Prasil IT, Haisel D, Gaudinova A, Havlova M, Gubis J, Vankova R (2010) Comparison of hormonal responses to heat, drought and combined stress in tobacco plants with elevated proline content. J Plant Physiol 167:1360–1370PubMedGoogle Scholar
  52. Dobrev PI, Kamínek M (2002) Fast and efficient separation of cytokinins from auxin and abscisic acid and their purification using mixed-mode solid-phase extraction. J Chromatogr A 950:21–29PubMedGoogle Scholar
  53. Dobrev PI, Havlíček L, Vágner M, Malbeck J, Kamínek M (2005) Purification and determination of plant hormones auxin and abscisic acid using solid phase extraction and two-dimensional high performance liquid chromatography. J Chromatogr A 1075:159–166PubMedGoogle Scholar
  54. Du F, Ruan G, Liang S, Xie F, Liu H (2012) Monolithic molecularly imprinted solid-phase extraction for the selective determination of trace cytokinins in plant samples with liquid chromatography-electrospray tandem mass spectrometry. Anal Bioanal Chem 404:489–501PubMedGoogle Scholar
  55. Duffield PH, Netting AG (2001) Methods for the quantitation of abscisic acid and its precursors from plant tissues. Anal Biochem 289:251–259PubMedGoogle Scholar
  56. Dunlap JR, Guinn G (1989) A simple purification of indole-3-acetic acid and abscisic acid for GC–SIM–MS analysis by microfiltration of aqueous samples through nylon. Plant Physiol 90:197–201PubMedCentralPubMedGoogle Scholar
  57. Durgbanshi A, Arbona V, Pozo O, Miersch O, Sancho JV, Gómez-Cadenas A (2005) Simultaneous determination of multiple phytohormones in plant extracts by liquid chromatography-electrospray tandem mass spectrometry. J Agric Food Chem 53:8437–8442PubMedGoogle Scholar
  58. Durley RC, Kannangara T, Simpson GM (1982) Leaf analysis for abscisic, phaseic and 3-indolylacetic acids by high-performance liquid chromatography. J Chromatogr 236:181–188Google Scholar
  59. Edlund A, Eklöf S, Sundberg B, Moritz T, Sandberg G (1995) A microscale technique for gas-chromatography mass-spectrometry measurements of picogram amounts of indole-3-acetic-acid in plant tissues. Plant Physiol 108:1043–1047PubMedCentralPubMedGoogle Scholar
  60. Emery RJN, Leport L, Barton JE, Turner NC, Atkins CA (1998) Cis-isomers of cytokinins predominate in chickpea seeds throughout their development. Plant Physiol 117:1515–1523PubMedCentralPubMedGoogle Scholar
  61. Emery RJN, Ma Q, Atkins CA (2000) The forms and sources of cytokinins in developing white lupine seeds and fruits. Plant Physiol 123:1593–1604PubMedCentralPubMedGoogle Scholar
  62. Engelberth J, Schmelz EA, Alborn HT, Cardoza YJ, Huang J, Tumlinson JH (2003) Simultaneous quantification of jasmonic acid and salicylic acid in plants by vapor-phase extraction and gas chromatography-chemical ionization-mass spectrometry. Anal Biochem 312:242–250PubMedGoogle Scholar
  63. Entsch B, Parker CW, Letham DS, Summons RE (1979) Preparation and characterization, using high-performance liquid chromatography, of an enzyme forming glucosides of cytokinins. Biochim Biophys Acta 570:124–139PubMedGoogle Scholar
  64. Eugster PJ, Knochenmuss R, Wolfender JL (2012) Ion mobility spectrometry in metabolite profiling of complex plant extracts. Planta Med 78:PJ46. doi: 10.1055/s-0032-1321206 Google Scholar
  65. Farmer EE, Ryan CA (1990) Interplant communication: airborne methyl jasmonate induces synthesis of proteinase inhibitors in plant leaves. Proc Natl Acad Sci USA 87:7713–7716PubMedCentralPubMedGoogle Scholar
  66. Farrow SC, Emery RJN (2012) Concurrent profiling of indole-3-acetic acid, abscisic acid, and cytokinins and structurally related purines by high-performance-liquid chromatography tandem electrospray mass spectrometry. Plant Methods 8:42PubMedCentralPubMedGoogle Scholar
  67. Flors V, Ton J, van Doorn R, Jakab G, García-Agustín P, Mauch-Mani B (2008) Interplay between JA, SA and ABA signalling during basal and induced resistance against Pseudomonas syringae and Alternaria brassicicola. Plant J 54:81–92PubMedGoogle Scholar
  68. Fonseca S, Chico JM, Solano R (2009) The jasmonate pathway: the ligand, the receptor and the core signalling module. Curr Opin Plant Biol 12:539–547PubMedGoogle Scholar
  69. Fujioka S, Sakurai A (1997) Brasinosteroids. Nat Prod Rep 14:1–10PubMedGoogle Scholar
  70. Galil J (1948) An ancient technique for ripening sycamore fruit in East Mediterranean countries. Econ Bot 22:178–190Google Scholar
  71. Galuszka P, Frébort I, Šebela M, Sauer P, Jacobsen S, Peč P (2001) Cytokinin oxidase or dehydrogenase? Mechanism of cytokinin degradation in cereals. Eur J Biochem 268:450–461PubMedGoogle Scholar
  72. Gamoh K, Abe H, Shimada K, Takatsuo S (1996) Liquid chromatography/mass spectrometry with atmospheric pressure chemical ionization of free brassinosteroids. Rapid Commun Mass Spectrom 10:903–906Google Scholar
  73. Gane R (1934) Production of ethylene by some ripening fruits. Nature 134:1008Google Scholar
  74. Ge L, Yong JW, Tan SN, Ong ES (2006) Determination of cytokinins in coconut (Cocos nucifera L.) water using capillary zone electrophoresis-tandem mass spectrometry. Electrophoresis 27:2171–2181PubMedGoogle Scholar
  75. Gómez-Cadenas A, Pozo OJ, García-Augustín P, Sancho JV (2002) Direct analysis of abscisic acid in crude plant extracts by liquid chromatography-electrospray/tandem mass spectrometry. Phytochem Anal 13:228–234PubMedGoogle Scholar
  76. Gomez-Roldan V, Fermas S, Brewer PB, Puech-Pagès V, Dun EA, Pillot J-P, Letisse F, Matusova R, Danoun S, Portais J-C, Bouwmeester H, Bécard G, Beveridge CA, Rameau C, Rochange SF (2008) Strigolactone inhibition of shoot branching. Nature 455:189–194PubMedGoogle Scholar
  77. Gruz J, Novak O, Strnad M (2008) Rapid analysis of phenolic acids in beverages by UPLC MS/MS. Food Chem 111:789–794Google Scholar
  78. Hammond-Kosack K, Jones JDG (2000) Responses to plant pathogens. In: Buchanan BB, Gruissem W, Jones RL (eds) Biochemistry and molecular biology of plants. American Society of Plant Physiologists, and Wiley, UK, pp 1102–1155Google Scholar
  79. Harrison MA, Walton DC (1975) Abscisic acid metabolism in water-stressed bean leaves. Plant Physiol 56:250–254PubMedCentralPubMedGoogle Scholar
  80. Harrison STL (2011) Cell disruption. In: Moo-Young M (ed) Comprehensive biotechnology, 2nd edn. Elsevier, Oxford, pp 619–640Google Scholar
  81. Hedden P, Thomas SG (2012) Gibberellin biosynthesis and its regulation. Biochem J 444:11–25PubMedGoogle Scholar
  82. Hernández L, Hernández P, Rica M, Galán F (1995) Determination of zeatin in plant extracts by square wave stripping polarography and differential pulse stripping polarography. Anal Chim Acta 315:33–39Google Scholar
  83. Hillman JR (1978) Isolation of plant growth substances. Cambridge University Press, LondonGoogle Scholar
  84. Hirai N, Koshimizu K (1983) A new conjugate of dihydrophaseic acid from avocado fruit. Agric Biol Chem 47:365–371Google Scholar
  85. Hoffmann-Benning S, Kende H (1992) On the role of abscisic acid and gibberellin in the regulation of growth in rice. Plant Physiol 99:1156–1161PubMedCentralPubMedGoogle Scholar
  86. Hogge LR, Balsevich JJ, Olson DJH, Abrams GD, Jacques SL (1993) Improved methodology for liquid chromatography/continuous flow secondary-ion mass spectrometry: quantitation of abscisic acid glucose ester using reaction monitoring. Rapid Commun Mass Spectrom 7:6–11Google Scholar
  87. Holub J, Hanuš J, Hanke DE, Strnad M (1998) Biological activity of cytokinins derived from Ortho- and Meta-hydroxybenzyladenine. Plant Growth Regul 26:109–115Google Scholar
  88. Horgan R, Scott IM (1987) Cytokinins. In: Rivier L, Crozier A (eds) Principles and practice of plant hormone analysis. Academic Press, London, pp 303–365Google Scholar
  89. Horgen PA, Nakagawa CH, Irvin RT (1984) Production of monoclonal antibodies to a steroid plant growth regulator. Can J Biochem Cell Biol 62:715–721Google Scholar
  90. Havlová M, Dobrev PI, Motyka V, Štorchová H, Libus J, Dobrá J, Malbeck J, Gaudinová A, Vanková R (2008) The role of cytokinins in responses to water deficit in tobacco plants over-expressing trans-zeatin O-glucosyltransferase gene under 35S or SAG12 promoters. Plant Cell Environ 31:341–353PubMedGoogle Scholar
  91. Hradecká V, Novák O, Havlíček L, Strnad M (2007) Immunoaffinity chromatography of abscisic acid combined with electrospray liquid chromatography–mass spectrometry. J Chromatogr B 847:162–173Google Scholar
  92. Huang WE, Wang H, Zheng H, Huang L, Singer AC, Thompson I, Whiteley AS (2005) Chromosomally located gene fusions constructed in Acinetobacter sp. ADP1 for the detection of salicylate. Environ Microbiol 7:1339–1348PubMedGoogle Scholar
  93. Huelin FE, Kennett BH (1959) Nature of the olefines produced by apples. Nature 184:996Google Scholar
  94. Huo F, Wang X, Han Y, Bai Y, Zhang W, Yuan H, Liu H (2012) A new derivatization approach for the rapid and sensitive analysis of brassinosteroids by using ultra high performance liquid chromatography-electrospray ionization triple quadrupole mass spectrometry. Talanta 99:420–425PubMedGoogle Scholar
  95. Hušková R, Pěchová D, Kotouček M, Lemr K (2000) Voltammetric behaviour and determination of some cytokinines on mercury electrode. Chem Listy 94:1004–1009Google Scholar
  96. Imbault N, Moritz T, Nilsson O, Chen H-J, Bollmark M, Sandberg G (1993) Separation and identification of cytokinins using combined capillary liquid-chromatography mass-spectrometry. Biol Mass Spectrom 22:201–210Google Scholar
  97. Izumi Y, Okazawa A, Bamba T, Kobayashi A, Fukusaki E (2009) Development of a method for comprehensive and quantitative analysis of plant hormones by highly sensitive nanoflow liquid chromatography-electrospray ionization-ion trap mass spectrometry. Anal Chim Acta 648:215–225PubMedGoogle Scholar
  98. Joel DM, Steffens JC, Matthews DE (1995) Germination of weedy root parasites. In: Kigel J, Galili G (eds) Seed development and germination. Dekker, New York, pp 567–597Google Scholar
  99. Kai K, Horita J, Wakasa K, Miyagawa H (2007a) Three oxidative metabolites of indole-3-acetic acid from Arabidopsis thaliana. Phytochemistry 68:1651–1663PubMedGoogle Scholar
  100. Kai K, Wakasa K, Miyagawa H (2007b) Metabolism of indole-3-acetic acid in rice: identification and characterization of N-beta-d-glucopyranosyl indole-3-acetic acid and its conjugates. Phytochemistry 68:2512–2522PubMedGoogle Scholar
  101. Kamínek M, Vaněk T, Motyka V (1987) Cytokinin activities of N6-benzyladenosine derivatives hydroxylated on the side chain phenyl ring. J Plant Growth Regul 6:113–120Google Scholar
  102. Kang BG, Rat PM (1969) Ethylene and carbon dioxide as mediators in the response of the bean hypocotyl hook to light and auxins. Planta 87:206–216PubMedGoogle Scholar
  103. Kaufmann A (2012) The current role of high-resolution mass spectrometry in food analysis. Anal Bioanal Chem 403:1233–1249PubMedGoogle Scholar
  104. Kögl F, Kostermans D (1934) Hetero-auxin als Stoffwechselprodukt niederer pflanzlicher Organismen. XIII. Isolierung aus Hefe. Z Phys Chem 228:113–121Google Scholar
  105. Kojima M, Kamada-Nobusada T, Komatsu H, Takei K, Kuroha T, Mizutani M, Ashkari M, Ueguchi-Tanaka M, Matsuoka M, Suzuki K, Sakakibara H (2009) Highly sensitive and high-throughput analysis of plant hormones using MS-probe modification and liquid chromatography-tandem mass spectrometry: an application for hormone profiling in Oryza sativa. Plant Cell Physiol 50:1207–1214Google Scholar
  106. Kovacik J, Gruz J, Backor M, Strnad M, Repcak M (2009) Salicylic acid-induced changes of growth and phenolic metabolism in Matricaria chamomilla plants. Plant Cell Rep 28:135–143PubMedGoogle Scholar
  107. Kowalczyk M, Sandberg G (2001) Quantitative analysis of indole-3-acetic acid metabolites in Arabidopsis. Plant Physiol 127:1845–1853PubMedCentralPubMedGoogle Scholar
  108. Kramell R, Atzorn R, Schneider G, Miersch O, Brückner C, Schmidt J, Sembdner G, Parthier B (1995) Occurrence and identification of jasmonic acid and its amino acid conjugates induced by osmotic stress in barley leaf tissue. J Plant Growth Regul 14:29–36Google Scholar
  109. Kristl J, Veber M, Krajničič B, Orešnik K (2005) Determination of jasmonic acid in Lemna minor (L.) by liquid chromatography with fluorescence detection. Anal Bioanal Chem 383:886–893PubMedGoogle Scholar
  110. Kudo T, Makita N, Kojima M, Tokunaga H, Sakakibara H (2012) Cytokinin activity of cis-zeatin and phenotypic alterations induced by overexpression of putative cis-zeatin-O-glucosyltransferase in rice. Plant Physiol 160:319–331PubMedCentralPubMedGoogle Scholar
  111. Kurakawa T, Ueda N, Maekawa M, Kobayashi K, Kojima M, Nagato Y, Sakakibara H, Kyozuka J (2007) Direct control of shoot meristem activity by a cytokinin-activating enzyme. Nature 445:652–655PubMedGoogle Scholar
  112. Kuwabara A, Ikegami K, Koshiba T, Nagata T (2003) Effects of ethylene and abscisic acid upon heterophylly in Ludwigia arcuata (Onagraceae). Planta 217:880–887PubMedGoogle Scholar
  113. Laloue M (1977) Cytokinins: 7-glucosylation is not a prerequisite of the expression of their biological activity. Planta 134:273–275PubMedGoogle Scholar
  114. Liu B-F, Zhong X-H, Lu Y-T (2002) Analysis of plant hormones in tobacco flowers by micellar electrokinetic capillary chromatography coupled with on-line large volume sample stacking. J Chromatogr A 945:257–265PubMedGoogle Scholar
  115. Liu H-T, Li Y-F, Luan T-G, Lan Ch-Y, Shu W-S (2007) Simultaneous determination of phytohormones in plant extracts using SPME and HPLC. Chromatographia 66:515–520Google Scholar
  116. Liu W-C, Carns HR (1961) Isolation of abscisin, an abscission accelerating substance. Science 134:384–385PubMedGoogle Scholar
  117. Liu X, Ma L, Lin YW, Lu YT (2003) Determination of abscisic acid by capillary electrophoresis with laser-induced fluorescence detection. J Chromatogr A 1021:209–213PubMedGoogle Scholar
  118. Liu X, Li DF, Wang Y, Lu YT (2004) Determination of 1-aminocyclopropane-1-carboxylic acid in apple extracts by capillary electrophoresis with laser-induced fluorescence detection. J Chromatogr A 1061:99–104PubMedGoogle Scholar
  119. Liu X, Hegeman AD, Gardner G, Cohen JD (2012a) Protocol: high-throughput and quantitative assays of auxin and auxin precursors from minute tissue samples. Plant Methods 8:31PubMedCentralPubMedGoogle Scholar
  120. Liu Z, Wei F, Feng Y-Q (2010) Determination of cytokinins in plant samples by polymer monolith microextraction coupled with hydrophilic interaction chromatography-tandem mass spectrometry. Anal Methods 2:1676–1685Google Scholar
  121. Liu Z, Yuan B-F, Feng Y-Q (2012b) Tandem solid phase extraction followed by online trapping-hydrophilic interaction chromatography-tandem mass spectrometry for sensitive detection of endogenous cytokinins in plant tissues. Phytochem Anal 23:559–568PubMedGoogle Scholar
  122. Liu Z, Cai B-D, Feng Y-Q (2012c) Rapid determination of endogenous cytokinins in plant samples by combination of magnetic solid phase extraction with hydrophilic interaction chromatography–tandem mass spectrometry. J Chromatogr B 891–892:27–35Google Scholar
  123. Ljung K, Sandberg G, Moritz T (2004) Hormone Analysis. In: Davies JP (ed) Plant hormones, biosynthesis, signal transduction, action!. Kluwer Academic Publishers, Dordrecht, pp 717–740Google Scholar
  124. Ljung K (2012) Auxin metabolism and homeostasis during plant development. Development 140:943–950Google Scholar
  125. López-Carbonell M, Jáuregui O (2005) A rapid method for analysis of abscisic acid (ABA) in crude extracts of water stressed Arabidopsis thaliana plants by liquid chromatography-mass spectrometry in tandem mode. Plant Physiol Biochem 43:407–411PubMedGoogle Scholar
  126. López-Carbonell M, Gabasa M, Jáuregui O (2009) Enhanced determination of abscisic acid (ABA) and abscisic acid glucose ester (ABA-GE) in Cistus albidus plants by liquid chromatography-mass spectrometry in tandem mode. Plant Physiol Biochem 47:256–261PubMedGoogle Scholar
  127. López-Ráez JA, Verhage A, Fernández I, García JM, Azcón-Aguilar C, Flors V, Pozo MJ (2010) Hormonal and transcriptional profiles highlight common and differential host responses to arbuscular mycorrhizal fungi and the regulation of the oxylipin pathway. J Exp Bot 61:2589–2601PubMedCentralPubMedGoogle Scholar
  128. Ludewig M, Dörffling K, Konig WA (1982) Electron-capture capillary gas chromatography and mass spectrometry of trifluoroacetylated cytokinins. J Chromatogr 243:93–98Google Scholar
  129. MacMillan J, Pryce RJ (1968) Further investigations of gibberellins in Phaseolus multiflorus by combined gas chromatography-mass spectrometry—the occurrence of gibberellin A20 (pharbitis gibberellin) and the structure of compound b. Tetrahedron Lett 9:1537–1664Google Scholar
  130. Maeda E (1965) Rate of lamina inclination in excised rice leaves. Physiol Plant 18:813–827Google Scholar
  131. Magome H, Nomura T, Hanada A, Takeda-Kamiya N, Ohnishi T, Shinma Y, Katsumata T, Kawaide H, Kamiya Y, Yamaguchi S (2013) CYP714B1 and CYP714B2 encode gibberellin 13-oxidases that reduce gibberellin activity in rice. Proc Natl Acad Sci USA 110:1947–1952PubMedCentralPubMedGoogle Scholar
  132. Mano Y, Nemoto K (2012) The pathway of auxin biosynthesis in plants. J Exp Bot 63:2853–2872PubMedGoogle Scholar
  133. Mapelli S, Rocchi P (1983) Separation and quantification of abscisic acid and its metabolites by high-performance liquid chromatography. Ann Bot 52:407–409Google Scholar
  134. Marcussen J, Ulvskov P, Olsen CE, Rajagopal R (1989) Preparation and properties of antibodies against indoleacetic acid (IAA)-C5-BSA, a novel ring-coupled IAA antigen, as compared to two other types of IAA-specific antibodies. Plant Physiol 89:1071–1078PubMedCentralPubMedGoogle Scholar
  135. Marek G, Carver R, Ding Y, Sathyanarayan D, Zhang X, Mou Z (2010) A high-throughput method for isolation of salicylic acid metabolic mutants. Plant Methods 6:21PubMedCentralPubMedGoogle Scholar
  136. Mashiguchi K, Tanaka K, Sakai T, Sugawara S, Kawaide H, Natsume M, Hanada A, Yaeno T, Shirasu K, Yao H, McSteen P, Zhao Y, Hayashi K, Kamiya Y, Kasahara H (2011) The main auxin biosynthesis pathway in Arabidopsis. Proc Natl Acad Sci USA 108:18512–18517PubMedCentralPubMedGoogle Scholar
  137. Mattivi F, Vrhovšek U, Versinia G (1999) Determination of indole-3-acetic acid, tryptophan and other indoles in must and wine by high-performance liquid chromatography with fluorescence detection. J Chromatogr A 855:227–235PubMedGoogle Scholar
  138. Mauriat M, Moritz T (2009) Analyses of GA20ox- and GID1-over-expressing aspen suggest that gibberellins play two distinct roles in wood formation. Plant J 58:989–1003PubMedGoogle Scholar
  139. McGaw BA, Burch LR (1995) Cytokinin biosynthesis and metabolism. In: Davies PJ (ed) Plant Hormones: physiology, biochemistry and molecular biology. Kluwer Academic Publisher, Dordrecht, pp 98–117Google Scholar
  140. Mertens R, Deus-Neumann B, Weiler EW (1983) Monoclonal antibodies for the detection and quantitation of the endogenous plant growth regulator, abscisic acid. FEBS Lett 160:269–272Google Scholar
  141. Meuwly P, Metraux JP (1993) Ortho-anisic acid as internal standard for the simultaneous quantitation of salicylic acid and its putative biosynthetic precursors in cucumber leaves. Anal Biochem 214:500–505PubMedGoogle Scholar
  142. Meyer A, Miersch O, Buttner C, Dathe W, Sembdner G (1984) Occurrence of the plant growth regulator jasmonic acid in plants. J Plant Growth Regul 3:1–8Google Scholar
  143. Mitchell JW, Livingston GA (1968) Methods of studying plant hormones and growth-regulating substances. Agricultural Handbook No. 336, Agric Res Ser USDAGoogle Scholar
  144. Mok MC, Mok DWS, Armstrong DJ (1978) Differential cytokinin structure-activity relationships in Phaseolus. Plant Physiol 61:72–75PubMedCentralPubMedGoogle Scholar
  145. Mok MC (1994) Cytokinins and plant development—an overview. In: Mok DWS, Mok MC (eds) Cytokinins: chemistry, activity and function. CRC Press, Boca Raton, pp 129–137Google Scholar
  146. Mok DWS, Martin RC (1994) Cytokinin metabolic enzymes. In: Mok DWS, Mok MC (eds) Cytokinins: chemistry, activity and function. CRC Press, Boca Raton, pp 129–137Google Scholar
  147. Morris RO (1977) Mass spectroscopic identification of cytokinins. Glucosyl zeatin and glucosyl ribosylzeatin from Vinca rosea crown gall. Plant Physiol 59:1029–1033PubMedCentralPubMedGoogle Scholar
  148. Most BH, Williams JC, Parker KJ (1968) Gas chromatography of cytokinins. J Chromatogr 38:136–138PubMedGoogle Scholar
  149. Müller A, Düchting P, Weiler EW (2002) A multiplex GC–MS/MS technique for the sensitive and quantitative single-run analysis of acidic phytohormones and related compounds, and its application to Arabidopsis thaliana. Planta 216:44–56PubMedGoogle Scholar
  150. Müller MJ, Brodschelm W, Spannagl E, Zenk MH (1993) Signaling in the elicitation process is mediated through the octadecanoid pathway leading to jasmonic acid. Proc Natl Acad Sci USA 90:7490–7494Google Scholar
  151. Müller MJ, Brodschelm W (1994) Quantification of jasmonic acid by capillary gas chromatography–negative chemical-ionization mass-spectrometry. Anal Biochem 218:425–435Google Scholar
  152. Nambara E, Marion-Poll A (2005) Abscisic acid biosynthesis and catabolism. Annu Rev Plant Biol 56:165–185PubMedGoogle Scholar
  153. Neill SJ, Horgan R, Heald JK (1983) Determination of the levels of abscisic acid-glucose ester in plants. Planta 157:371–375PubMedGoogle Scholar
  154. Nejlubow DN (1901) Über die horozontale Nutation der Stengel von Pisum sativum and einiger anderen. Pflanzen Beiträge and Botanik Zentralblatt 10:128–139Google Scholar
  155. Netting AG, Milborrow BV, Duffield AM (1982) Determination of abscisic acid in Eucalyptus haemastoma leaves using gas chromatography/mass spectrometry and deuterated internal standards. Phytochemistry 21:385–389Google Scholar
  156. Nicander B, Ståhl U, Björkman PO, Tillberg E (1993) Immunoaffinity co-purification of cytokinins and analysis by high-performance liquid chromatography with ultraviolet spectrum detection. Planta 189:312–320PubMedGoogle Scholar
  157. Nordström A, Tarkowski P, Tarkowska D, Doležal K, Åstot C, Sandberg G, Moritz T (2004) Derivatization for LC-Electrospray Ionization-MS: a tool for improving reversed-phase separation and ESI responses of bases, ribosides, and intact nucleotides. Anal Chem 76:2869–2877PubMedGoogle Scholar
  158. Normanly J (2010) Approaching cellular and molecular resolution of auxin biosynthesis and metabolism. Cold Spring Harb Perspect Biol 2:a001594PubMedCentralPubMedGoogle Scholar
  159. Novák O, Tarkowski P, Tarkowská D, Doležal K, Lenobel R, Strnad M (2003) Quantitative analysis of cytokinins in plants by liquid chromatography-single-quadrupole mass spectrometry. Anal Chim Acta 480:207–218Google Scholar
  160. Novák O, Hauserová E, Amakorová P, Doležal K, Strnad M (2008) Cytokinin profiling in plant tissues using ultra-performance liquid chromatography-electrospray tandem mass spectrometry. Phytochemistry 69:2214–2224PubMedGoogle Scholar
  161. Novák O, Hényková E, Sairanen I, Kowalczyk M, Pospíšil T, Ljung K (2012) Tissue specific profiling of the Arabidopsis thaliana auxin metabolome. Plant J 72:523–536PubMedGoogle Scholar
  162. Ohkuma K, Lyon JL, Addicott FT, Smith OE (1963) Abscisin II, an abscission-accelerating substance from young cotton fruit. Science 142:1592–1593PubMedGoogle Scholar
  163. Pacáková V, Štulík K, Vlasáková V, Březinová A (1997) Capillary electrophoresis of cytokinins and cytokinin ribosides. J Chromatogr A 764:331–335PubMedGoogle Scholar
  164. Pan X, Welti R, Wang X (2008) Simultaneous quantification of major phytohormones and related compounds in crude plant extracts by liquid chromatography-electrospray tandem mass spectrometry. Phytochemistry 69:1773–1781PubMedGoogle Scholar
  165. Paré PW, Tumlinson JH (1999) Plant volatiles as a defense against insect herbivores. Plant Physiol 121:325–331PubMedCentralPubMedGoogle Scholar
  166. Parfrey LW, Lahr DJG, Knoll AH, Katz LA (2011) Estimating the timing of early eukaryotic diversification with multigene molecular clocks. Proc Natl Acad Sci USA 108:13624–13629PubMedCentralPubMedGoogle Scholar
  167. Parthier B (1991) Jasmonates, new regulators of plant growth and development: many facts and few hypotheses on their actions. Bot Acta 104:446–454Google Scholar
  168. Parthier B, Brückner C, Dathe W, Hause B, Herrmann G, Knofel H-D, Kramell H-M, Kramell R, Lehmann J, Miersch O, Reinbothe S, Sembdner G, Wasternack C, zur Nieden U (1992) Jasmonates: Metabolism, biological activities, and modes of action in senescence and stress responses. In: Karssen CM, van Loon LC, Vreugdenhil DD (eds) Progress in plant growth regulation. Kluwer Academic Publishers, Dordrecht, pp 276–285Google Scholar
  169. Pastor V, Vicent C, Cerezo M, Mauch-Mani B, Dean J, Flors V (2012) Detection, characterization and quantification of salicylic acid conjugates in plant extracts by ESI tandem mass spectrometric techniques. Plant Physiol Biochem 53:19–26PubMedGoogle Scholar
  170. Patterson SE (2001) Cutting loose. Abscission and dehiscence in Arabidopsis. Plant Physiol 126:494–500PubMedCentralPubMedGoogle Scholar
  171. Pěnčík A, Rolčík J, Novák O, Magnus V, Barták P, Buchtík R, Salopek-Sondi B, Strnad M (2009) Isolation of novel indole-3-acetic acid conjugates by immunoaffinity extraction. Talanta 80:651–655PubMedGoogle Scholar
  172. Pengelly WL, Bandurski RS, Schulze A (1981) Validation of a radioimmunoassay for indole-3-acetic acid using gas chromatography-selected ion monitoring-mass spectrometry. Plant Physiol 68:96–98PubMedCentralPubMedGoogle Scholar
  173. Perrine FM, Rolfe BG, Hynes MF, Hocart CH (2004) Gas chromatography-mass spectrometry analysis of indoleacetic acid and tryptophan following aqueous chloroformate derivatisation of Rhizobium exudates. Plant Physiol Biochem 42:723–729PubMedGoogle Scholar
  174. Pertry I, Václavíková K, Depuydt S, Galuszka P, Spíchal L, Temmerman W, Stes E, Schmülling T, Kakimoto T, Van Montagu MCE, Strnad M, Holsters M, Tarkowski P, Vereecke D (2009) Identification of Rhodococcus fascians cytokinins and their modus operandi to reshape the plant. Proc Natl Acad Sci USA 106:929–934PubMedCentralPubMedGoogle Scholar
  175. Petritis K, Koukaki G, Koussissi E, Elfakir C, Dreux M, Dourtoglou V (2003) The simultaneous determination of 1-aminocyclopropane-1-carboxylic acid and cyclopropane-1,1-dicarboxylic acid in Lycopersicum esculentum by high-performance liquid chromatography-electrospray tandem mass spectrometry. Phytochem Anal 14:347–351PubMedGoogle Scholar
  176. Pichersky E, Gershenzon J (2002) The formation and function of plant volatiles: perfumes for pollinator attraction and defense. Curr Opin Plant Biol 5:237–243PubMedGoogle Scholar
  177. Pieterse CM, Van Loon LC (2004) NPR1: the spider in the web of induced resistance signalling pathways. Curr Opin Plant Biol 7:456–464PubMedGoogle Scholar
  178. Pouvreau J-B, Gaudin Z, Auger B, Lechat M-M, Gauthier M, Delavault P, Simier P (2013) A high-throughput seed germination assay for root parasitic plants. Plant Methods 9:32PubMedCentralPubMedGoogle Scholar
  179. Prinsen E, Redig P, Van Dongen W, Esmans EL, Van Onckelen HA (1995) Quantitative analysis of cytokinins by electrospray tandem mass spectrometry. Rapid Commun Mass Spectrom 9:948–953Google Scholar
  180. Pryce RJ, MacMIllan J, McCormica A (1967) The identification of bamboo gibberellin in Phaseolus multiflorus by combined gas chromatography-mass spectrometry. Tetrahedron Lett 8:5009–5011Google Scholar
  181. Quesnelle PE, Emery RJN (2007) Cis-cytokinins that predominate in Pisum sativum during early embryogenesis will accelerate embryo growth in vitro. Can J Bot 85:91–103Google Scholar
  182. Quittenden LJ, Davies NW, Smith JA, Molesworth PP, Tivendale ND, Ross JJ (2009) Auxin biosynthesis in pea: characterization of the tryptamine pathway. Plant Physiol 151:1130–1138PubMedCentralPubMedGoogle Scholar
  183. Radhika V, Kost C, Bonaventure G, David A, Boland W (2012) Volatile emission in bracken fern is induced by jasmonates but not by Spodoptera littoralis or Strongylogaster multifasciata herbivory. PLoS ONE 7:1–9Google Scholar
  184. Ribnicky DM, Cooke TJ, Cohen JD (1998) A microtechnique for the analysis of free and conjugated indole-3-acetic acid in milligram amounts of plant tissue using a benchtop gas chromatograph-mass spectrometer. Planta 204:1–7PubMedGoogle Scholar
  185. Richmond AE, Lang A (1957) Effect of kinetin on protein content and survival of detached Xanthium leaves. Science 125:650–651Google Scholar
  186. Sandberg G, Ljung K, Alm P (1985) Precision and accuracy of radioimmunoassays in the analysis of endogenous 3-indole acetic acid from needles of Scots pine. Phytochemistry 24:1439–1442Google Scholar
  187. Sato D, Awad AA, Chae SH, Yokota T, Sugimoto Y, Takeuchi Y, Yoneyama K (2003) Analysis of strigolactones, germination stimulants for Striga and Orobanche, by high-performance liquid chromatography/tandem mass spectrometry. J Agric Food Chem 51:1162–1168PubMedGoogle Scholar
  188. Sato D, Awad AA, Takeuchi Y, Yoneyama K (2005) Confirmation and quantification of strigolactones, germination stimulants for root parasitic plants Striga and Orobanche, produced by cotton. Biosci Biotechnol Biochem 69:98–102PubMedGoogle Scholar
  189. Schmelz EA, Engelberth J, Alborn HAT, O’Donnell P, Sammons M, Toshima H, Tumlinson JH 3rd (2003) Simultaneous analysis of phytohormones, phytotoxins, and volatile organic compounds in plants. Proc Nat Acad Sci USA 100:10552–10557PubMedCentralPubMedGoogle Scholar
  190. Schmelz EA, Engelberth J, Tumlinson JH, Block A, Alborn HAT (2004) The use of vapor phase extraction in metabolic profiling of phytohormones and other metabolites. Plant J 39:790–808PubMedGoogle Scholar
  191. Schneider G, Ziethe F, Schmidt J (1997) Liquid chromatography/electrospray ionisation-tandem mass spectrometry: a tool for the identification and quantification of abscisic acid glucose ester. Chromatographia 45:78–80Google Scholar
  192. Scott IM, Yamamoto H (1994) Mass spectrometric quantification of salicylic acid in plant tissues. Phytochemistry 37:335–336Google Scholar
  193. Segal A, Gorecki T, Mussche P, Lips J, Pawliszyn J (2000) Development of membrane extraction with a sorbent interface-micro gas chromatography system for field analysis. J Chromatogr A 873:13–27PubMedGoogle Scholar
  194. Segarra G, Jáuregui O, Casanova E, Trillas I (2006) Simultaneous quantitative LC-ESI-MS/MS analyses of salicylic acid and jasmonic acid in crude extracts of Cucumis sativus under biotic stress. Phytochemistry 67:395–401PubMedGoogle Scholar
  195. Sembdner G, Schneider G, Schreiber K (1988) Methoden zur Pflanzenhormonanalyse. Jena, VEB Gustav Fischer Verlag, p 176Google Scholar
  196. Sembdner G, Parthier B (1993) The biochemistry and the physiological and molecular actions of jasmonates. Annu Rev Plant Physiol Plant Mol Biol 44:569–589Google Scholar
  197. Shapiro AD, Gutsche AT (2003) Capillary electrophoresis-based profiling and quantitation of total salicylic acid and related phenolics for analysis of early signalling in Arabidopsis disease resistance. Anal Biochem 320:223–233PubMedGoogle Scholar
  198. Sharp RE, LeNoble ME (2002) ABA, ethylene and the control of shoot and root growth under water stress. J Exp Bot 53:33–37PubMedGoogle Scholar
  199. Siame BA, Weerasuriya Y, Wood K, Ejeta G, Butler LG (1993) Isolation of strigol, a germination stimulant for Striga asiatica, from host plants. J Agric Food Chem 41:1488–1491Google Scholar
  200. Singh V, Roy S, Giri MK, Chaturvedi R, Chowdhury Z, Shah J, Nandi AK (2013) Arabidopsis thaliana FLOWERING LOCUS D is required for systemic acquired resistance. Mol Plant Microbe In 26:1079–1088Google Scholar
  201. Skoog F, Miller CO (1957) Chemical regulation of growth and organ formation in plant tissue cultured in vitro. Symp Soc Exp Biol 11:118–131PubMedGoogle Scholar
  202. Smets R, Claes V, Van Onckelen HA, Prinsen E (2003) Extraction and quantitative analysis of 1-aminocyclopropane-1-carboxylic acid in plant tissue by gas chromatography coupled to mass spectrometry. J Chromatogr A 993:79–87PubMedGoogle Scholar
  203. Schmitz RY, Skoog F (1972) Cytokinins: synthesis and biological activity of geometric and position isomers of zeatin. Plant Physiol 50:702–705PubMedCentralPubMedGoogle Scholar
  204. Sotelo-Silveira M, Cucinotta M, Chauvin A-L, Chávez Montes RA, Colombo L, Marsch-Martínez N, de Folter S (2013) Cytochrome P450 CYP78A9 is involved in Arabidopsis reproductive development. Plant Physiol 162:2779–2799Google Scholar
  205. Staswick PE, Tiryaki I (2004) The oxylipin signal jasmonic acid is activated by an enzyme that conjugates it to isoleucine in Arabidopsis. Plant Cell 16:2117–2127PubMedCentralPubMedGoogle Scholar
  206. Stepanova AN, Yun J, Robles LM, Novak O, He W, Guo H, Ljung K, Alonso JM (2011) The Arabidopsis YUCCA1 flavin monooxygenase functions in the indole-3-pyruvic acid branch of auxin biosynthesis. Plant Cell 23:3961–3973PubMedCentralPubMedGoogle Scholar
  207. Strnad M (1997) The aromatic cytokinins. Physiol Plantarum 101:674–688Google Scholar
  208. Sugawara S, Hishiyama S, Jikumaru Y, Hanada A, Nishimura T, Koshiba T, Zhao Y, Kamiya Y, Kasahara H (2009) Biochemical analyses of indole-3-acetaldoxime-dependent auxin biosynthesis in Arabidopsis. Proc Natl Acad Sci USA 106:5430–5435PubMedCentralPubMedGoogle Scholar
  209. Sundberg B, Sandberg G, Crozier A (1986) Purification of indole-3-acetic acid in plant extracts by immunoaffinity chromatography. Phytochemistry 25:295–298Google Scholar
  210. Suttle JC, Banowetz GM (2000) Changes in cis-zeatin and cis-zeatin riboside levels and biological activity during potato tuber dormancy. Physiol Plantarum 109:68–74Google Scholar
  211. Svačinová J, Novák O, Plačková L, Lenobel R, Holík J, Strnad M, Doležal K (2012) A new approach for cytokinin isolation from Arabidopsis tissues using miniaturized purification: pipette tip solid-phase extraction. Plant Methods 8:17PubMedCentralPubMedGoogle Scholar
  212. Svatoš A, Antonchick A, Schneider B (2004) Determination of brassinosteroids in the sub-femtomolar range using dansyl-3-aminophenylboronate derivatization and electrospray mass spectrometry. Rapid Commun Mass Spectrom 18:816–821PubMedGoogle Scholar
  213. Swaczynová J, Novák O, Hauserová E, Fuksová K, Šíša M, Kohout L, Strnad M (2007) New techniques for the estimation of naturally occurring brassinosteroids. J Plant Growth Regul 26:1–14Google Scholar
  214. Takagi M, Yokota T, Murofushi N, Ota Y, Takahashi N (1985) Fluctuation of endogenous cytokinin contents in rice during its life cycle—quantification of cytokinins by selected ion monitoring using deuterium-labelled internal standards. Agr Biol Chem 49:3271–3277Google Scholar
  215. Takatsuo S, Ying B, Morisaki M, Ikekawa N (1982) Microanalysis of brassinolide and its analogues by gas chromatography and gas chromatography-mass spectrometry. J Chromatogr 239:233–241Google Scholar
  216. Takatsuo S, Yokota T (1999) Biochemical analysis of natural brassinosteroids. In: Sakurai A, Yokota T, Clouse SD (eds) Brassinosteroids. Springer, Tokyo, pp 47–68Google Scholar
  217. Takei K, Yamaya T, Sakakibara H (2004) Arabidopsis CYP735A1 and CYP735A2 encode cytokinin hydroxylases that catalyze the biosynthesis of trans-zeatin. J Biol Chem 279:41866–41872PubMedGoogle Scholar
  218. Tam YY, Normanly J (1998) Determination of indole-3-pyruvic acid levels in Arabidopsis thaliana by gas chromatography-selected ion monitoring-mass spectrometry. J Chromatogr A 800:101–108PubMedGoogle Scholar
  219. Tamogami S, Kodama O (1998) Quantification of amino acid conjugates of jasmonic acid in rice leaves by high-performance liquid chromatography turboionspray tandem mass spectrometry. J Chromatogr A 822:310–315Google Scholar
  220. Tarkowská D, Kotouček M, Doležal K (2003) Electrochemical reduction of 6-benzylaminopurine at mercury electrodes and its analytical application. Collect Czech Chem Commun 68:1076–1093Google Scholar
  221. Teale WD, Paponov IA, Palme K (2006) Auxin in action: signalling, transport and the control of plant growth and development. Nat Rev Mol Cell Biol 7:847–859PubMedGoogle Scholar
  222. Thomson MJ, Mandava NB, Meudt WJ, Lusby WR, Spaulding DW (1981) Synthesis and biological activity of brassinolide and its 22 beta, 23 beta-isomer: novel plant growth-promoting steroids. Steroids 38:567–580Google Scholar
  223. Tidd BK (1964) Dissociation constants of gibberellins. J Chem Soc 295:1521–1523Google Scholar
  224. Tsuchiya Y, McCourt P (2009) Strigolactones: a new hormone with a past. Curr Opin Biol 12:556–561Google Scholar
  225. Turečková V, Novák O, Strnad M (2009) Profiling ABA metabolites in Nicotiana tabacum L. leaves by ultra-performance liquid chromatography–electrospray tandem mass spectrometry. Talanta 80:390–399PubMedGoogle Scholar
  226. Umehara M, Hanada A, Yoshida S, Akiyama K, Arite T, Takeda-Kamiya N, Magome H, Kamiya Y, Shirasu K, Yoneyama K, Kyozuka J, Yamaguchi S (2008) Inhibition of shoot branching by new terpenoid plant hormones. Nature 455:195–200PubMedGoogle Scholar
  227. Urbanová T, Tarkowská D, Novák O, Hedden P, Strnad M (2013) Analysis of gibberellins as free acids by ultra performance liquid chromatography-tandem mass spectrometry. Talanta 112:85–94PubMedGoogle Scholar
  228. van Rhijn JA, Heskampa HH, Davelaarb E, Jordib W, Lelouxa MS, Brinkmanc UAT (2001) Quantitative determination of glycosylated and aglycon isoprenoid cytokinins at sub-picomolar levels by microcolumn liquid chromatography combined with electrospray tandem mass spectrometry. J Chromatogr A: 929:31–42Google Scholar
  229. Van Meulebroek L, Vanden Bussche J, Steppe K, Vanhaecke L (2012) Ultra-high performance liquid chromatography coupled to high resolution Orbitrap mass spectrometry for metabolomic profiling of the endogenous phytohormonal status of the tomato plant. J Chromatogr A 1260:67–80PubMedGoogle Scholar
  230. VanDoorn A, Bonaventure G, Schmidt DD, Baldwin IT (2011) Regulation of jasmonate metabolism and activation of systemic signaling in Solanum nigrum: COI1 and JAR4 play overlapping yet distinct roles. New Phytol 190:640–652PubMedGoogle Scholar
  231. Varbanova M, Yamaguchi S, Yang Y, McKelvey K, Hanada A, Borochov R, Yu F, Jikumaru Y, Ross J, Cortes D, Je Ma Ch, Noel JP, Mander L, Shulaev V, Kamiya Y, Rodermel S, Weiss D, Pichersky E (2007) Methylation of gibberellins by Arabidopsis GAMT1 and GAMT2. Plant Cell 19:32–45PubMedCentralPubMedGoogle Scholar
  232. Veach YK, Martin RC, Mok DW, Malbeck J, Vankova R, Mok MC (2003) O-glucosylation of cis-zeatin in maize. Characterization of genes, enzymes, and endogenous cytokinins. Plant Physiol 131:1374–1380Google Scholar
  233. Verberne MC, Brouwer N, Delbianco F, Linthorst HJ, Bol JF, Verpoorte R (2002) Method for the extraction of the volatile compound salicylic acid from tobacco leaf material. Phytochem Anal 13:45–50PubMedGoogle Scholar
  234. Vilaró F, Canela-Xandri A, Canela R (2006) Quantification of abscisic acid in grapevine leaf (Vitis vinifera) by isotope-dilution liquid chromatography-mass spectrometry. Anal Bioanal Chem 386:306–312PubMedGoogle Scholar
  235. Vyroubalová Š, Václavíková K, Turečková V, Novák O, Šmehilová M, Hluska T, Ohnoutková L, Frébort I, Galuszka P (2009) Characterization of new maize genes putatively involved in cytokinin metabolism and their expression during osmotic stress in relation to cytokinin levels. Plant Physiol 151:433–447PubMedCentralPubMedGoogle Scholar
  236. Wada K, Shingo Marumo S, Abe H, Morishita T, Nakmura K, Uchiyama M, Mori K (1984) A rice lamina inclination test—a micro-quantitative bioassay for brassinosteroids. Agric Biol Chem 3:719–726Google Scholar
  237. Walton D, Dashek W, Galson E (1979) A radioimmunoassay for abscisic acid. Planta 146:139–145PubMedGoogle Scholar
  238. Wasternack C, Kombrink E (2010) Jasmonates: structural requirements for lipid-derived signals active in plant stress responses and development. ACS Chem Biol 5:63–77PubMedGoogle Scholar
  239. Wasternack C, Hause B (2013) Jasmonates: biosynthesis, perception, signal transduction and action in plant stress response, growth and development. an update to the 2007 review in Annals of Botany. Ann Bot 111:1021–1058PubMedCentralPubMedGoogle Scholar
  240. Watts SH, Wheeler CT, Hillman JR, Berries AMM, Crozier A, Math VB (1983) Abscisic acid in the nodulated root system of Alnus glutinosa. New Phytol 95:203–208Google Scholar
  241. Weiler EW (1979) Radioimmunoassay for the determination of free and conjugated abscisic acid. Planta 144:255–263PubMedGoogle Scholar
  242. Weiler EW (1980) Radioimmunoassays for the differential and direct analysis of free and conjugated abscisic acid in plant extracts. Planta 148:262–272PubMedGoogle Scholar
  243. Weiler EW (1982) An enzyme-immunoassay for cis(+)-abscisic acid. Physiol Plant 54:510–514Google Scholar
  244. Went FW, Thimann KV (1937) Phytohormones. Macmillan, New YorkGoogle Scholar
  245. Wilbert SM, Ericsson LH, Gordon MP (1998) Quantification of jasmonic acid, methyl jasmonate, and salicylic acid in plants by capillary liquid chromatography electrospray tandem mass spectrometry. Anal Biochem 257:186–194PubMedGoogle Scholar
  246. Witters E, Vanhoutte K, Dewitte W, Macháčková I, Benková E, Van Dongen W, Esmans EL, Van Onckelen HA (1999) Analysis of cyclic nucleotides and cytokinins in minute plant samples using phase-system switching capillary electrospray–liquid chromatography–tandem mass spectrometry. Phytochem Anal 10:143–151Google Scholar
  247. Woodward AW, Bartel B (2005) Auxin: regulation, action, and interaction. Ann Bot 95:707–735PubMedGoogle Scholar
  248. Wu Y, Hu B (2009) Simultaneous determination of several phytohormones in natural coconut juice by hollow fiber-based liquid-liquid-liquid microextraction-high performance liquid chromatography. J Chromatogr A 1216:7657–7663PubMedGoogle Scholar
  249. Xi Z, Zhang Z, Sun Y, Shi Z, Tian W (2009) Determination of indole-3-acetic acid and indole-3-butyric acid in mung bean sprouts using high performance liquid chromatography with immobilized Ru(bpy)32+–KMnO4 chemiluminescence detection. Talanta 79:216–221PubMedGoogle Scholar
  250. Xie S, Wang F, Chen Z (2012) Determination of endogenous jasmonic acid in plant samples by liquid chromatography-electrochemical detection based on derivatisation with dopamine. Analyst 138:1226–1231Google Scholar
  251. Xie X, Yoneyama K, Yoneyama K (2010) The strigolactone story. Ann Rev Phytopathol 48:93–117Google Scholar
  252. Xie X, Yoneyama K, Kisugi T, Uchida K, Ito S, Akiyama K, Hayashi H, Yokota T, Nomura T, Yoneyama K (2013) Confirming stereochemical structures of strigolactones produced by rice and tobacco. Mol Plant 6:153–163PubMedCentralPubMedGoogle Scholar
  253. Xiong X-J, Rao W-B, Guo X-F, Wang H, Zhang H-S (2012) Ultrasensitive determination of jasmonic acid in plant tissues using high-performance liquid chromatography with fluorescence detection. J Agric Food Chem 60:5107–5111PubMedGoogle Scholar
  254. Yang YY, Yamaguchi I, Kato Y, Weiler EW, Murofuchi N, Takahashi N (1993) Qualitative and semi-quantitative analyses of cytokinins using LC/APCI-MS in combination with ELISA. J Plant Growth Regul 12:21–25Google Scholar
  255. Yokota T, Watanabe S, Ogino Y (1990) Radioimmunoasay for brassinosteroids and its use for comparative analysis of brassinosteroids in stems and seeds of Phaseolus vulgaris. J Plant Growth Regul 9:151–159Google Scholar
  256. Yokota T, Sakai H, Okuno K, Yoneyama K, Takeuchi Y (1998) Alectrol and orobanchol, germination stimulants for Orobanche minor, from its host red clover. Phytochemistry 49:1967–1973Google Scholar
  257. Zhang ZL, Liu X, Li DF, Lu YT (2005) Determination of jasmonic acid in bark extracts from Hevea brasiliensis by capillary electrophoresis with laser-induced fluorescence detection. Anal Bioanal Chem 382:1616–1619PubMedGoogle Scholar
  258. Zhou R, Squires TM, Ambrose SJ, Abrams SR, Ross AR, Cutler AJ (2003) Rapid extraction of abscisic acid and its metabolites for liquid chromatography-tandem mass spectrometry. J Chromatogr A 1010:75–85PubMedGoogle Scholar
  259. Zeevaart JAD, Milborrow BV (1976) Metabolism of abscisic acid and the occurrence of epi-dihydrophaseic acid in Phaseolus vulgaris. Phytochemistry 15:493–500Google Scholar
  260. Zhang Y, Li Y, Hu Y, Li G, Chen Y (2010) Preparation of magnetic indole-3-acetic acid imprinted polymer beads with 4-vinylpyridine and β-cyclodextrin as binary monomer via microwave heating initiated polymerization and their application to trace analysis of auxins in plant tissues. J Chromatogr A 1217:7337–7344PubMedGoogle Scholar
  261. Zwanenburg B, Mwakaboko AS, Reizelman A, Anilkumar G, Sethumadhavan D (2009) Structure and function of natural and synthetic signalling molecules in parasitic weed germination. Pest Manag Sci 65:478–491PubMedGoogle Scholar

Copyright information

© Springer-Verlag Berlin Heidelberg 2014

Authors and Affiliations

  • Danuše Tarkowská
    • 1
  • Ondřej Novák
    • 1
  • Kristýna Floková
    • 1
  • Petr Tarkowski
    • 2
  • Veronika Turečková
    • 1
  • Jiří Grúz
    • 1
  • Jakub Rolčík
    • 1
  • Miroslav Strnad
    • 1
  1. 1.Laboratory of Growth Regulators, Centre of the Region Haná for Biotechnological and Agricultural ResearchInstitute of Experimental Botany ASCR and Palacký UniversityOlomoucCzech Republic
  2. 2.Department of Protein Biochemistry and Proteomics, Faculty of Science, Centre of the Region Haná for Biotechnological and Agricultural ResearchPalacký UniversityOlomoucCzech Republic

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