Conjugates of auxin and cytokinin

Phytochemistry 70 (2009) 957–969

Contents lists available at ScienceDirect

Phytochemistry journal homepage: www.elsevier.com/locate/phytochem

Review

Conjugates of auxin and cytokinin Andrzej Bajguz *, Alicja Piotrowska University of Bialystok, Institute of Biology, Swierkowa 20 B, 15-950 Bialystok, Poland

a r t i c l e

i n f o

Article history: Received 4 March 2009 Received in revised form 7 May 2009 Available online 12 June 2009 Keywords: Conjugates Auxins Cytokinins

a b s t r a c t Plant growth and developmental processes as well as environmental responses require the action and cross talk of phytohormones including auxins and cytokinins. Active phytohormones are changed into multiple forms by acylation, esterification or glycosylation, for example. It seems that conjugated compounds could serve as pool of inactive phytohormones that can be converted to active forms by de-conjugation reactions. The concept of reversible conjugation of auxins and cytokinins suggests that under changeable environmental, developmental or physiological conditions these compounds can be a source of free hormones. Phytohormones metabolism may result in a loss of activity and decrease the size of the bioactive pool. All metabolic steps are in principle irreversible, except for some processes such as the formation of ester, glucoside and amide conjugates, where the free compound can be liberated by enzymatic hydrolysis. The role, chemistry, synthesis and hydrolysis of conjugated forms of two classes of plant hormones are discussed. Ó 2009 Elsevier Ltd. All rights reserved.

Contents 1. 2.

3.

Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Auxin conjugates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.1. The role and structure of auxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.2. The chemistry and function of auxin conjugates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.3. The synthesis and hydrolysis of auxin ester conjugates in monocotyledonous plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.4. Auxin conjugate profile in dicotyledonous plants . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.5. Synthesis and hydrolysis of amide conjugates of auxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 2.6. Peptide and methylate conjugates of auxins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cytokinin conjugates . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.1. The structure, role and metabolism of cytokinins. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.2. Conjugation of adenine ring of cytokinins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3.3. Conjugation of isoprenoid chain of cytokinins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

1. Introduction Plant hormones are low-molecular weight natural products that act at micromolar (or even lower) concentrations to regulate essentially all physiological and developmental processes during a plant’s life cycle. These structurally diverse compounds include auxins, cytokinins, abscisic acid, gibberellins, ethylene, polyamines, jasmonates, salicylic acid, and brassinosteroids. Among phytohormones, auxins and cytokinins play a crucial role in various phases of plant growth and development. Auxins and cytoki* Corresponding author. Tel.: +48 8 574 572 92; fax: +48 8 574 573 02. E-mail address: [email protected] (A. Bajguz). 0031-9422/$ - see front matter Ó 2009 Elsevier Ltd. All rights reserved. doi:10.1016/j.phytochem.2009.05.006

957 958 958 958 959 959 960 963 963 963 963 964 968

nins interact in the control of many central developmental processes in plants, particularly in apical dominance and root and shoot development. It is clearly documented that auxin may regulate cytokinin level and metabolism and vice versa (Nordström et al., 2004). Despite their importance, metabolic pathways regulating the activity and the level of the two hormones have been identified recently. Several compounds in the biosynthetic and degradative pathways of auxins and cytokinins can exhibit biological activity, giving rise to a very complex network of signaling molecules at the cellular level. The concept of reversible conjugation of phytohormones suggests that under changeable physiological conditions these compounds can be a source of free hormones (Weyers and Paterson, 2001). The hormonal homeostasis has been

958

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

defined as ‘‘the maintenance of a steady state concentration of the hormones in the receptive tissue appropriate to any fixed environmental condition” (Cohen and Bandurski, 1982). The reversible conjugation could be a mechanism to regulate the pool of the physiologically active forms of auxins and cytokinins. Phytohormones metabolism may result in a loss of activity and decrease the size of the bioactive pool. All metabolic steps are in principle irreversible, except for some processes such as the formation of ester, glucoside and amide conjugates, where the free compound can be liberated by enzymatic hydrolysis. All-round studying the structure, functions, and action mechanism of auxins and cytokinins attracted the attention of many researchers, both earlier and now (Weyers and Paterson, 2001). In the area of auxin and cytokinin metabolism, the chemistry of conjugates forms is just beginning to be discovered and is providing a somewhat better picture of phytohormone homeostasis. Therefore, this review focuses on the recent progress in research to address the biology of conjugation and de-conjugation of these two classes of plant hormones. 2. Auxin conjugates 2.1. The role and structure of auxins Auxins were the first plant hormones discovered and have been extensively examined for many decades. In many bioassays, it has been shown that auxins play a critical role in plant growth and development. Auxins are thought to regulate or influence diverse responses on a whole-plant level, such as tropisms, apical dominance and root initiation, and responses on cellular level, such as cell enlargement, division, and differentiation (Hagen and Guilfoyle, 2002). Our knowledge about the physiological and molecular aspects of auxin role is rapidly expanding. Great advances have been also reported on the characterization of auxin metabolism, including conjugation and de-conjugation. Auxins belong to chemically diverse compounds, most of which have an aromatic system such as indole, phenyl or naphthalene ring with a side chain containing a carboxyl group attached (Fig. 1). Indole-3-acetic acid (IAA) is the natural auxin commonly occurring in all vascular and lower plants (Cooke et al., 2002). A

Fig. 2. The role of indole-3-acetic acid (IAA) conjugates in auxin homeostatis.

chlorinated form of IAA with high auxin activity, 4-Cl-IAA, is found in several plant species (Slovin et al., 1999). In addition to the indolic auxins, phenylacetic acid has been identified in plants and is an active auxin in the most bioassays (Ludwig-Müller and Cohen, 2002). Certain IAA precursors, such as indole-3-acetonitrile and indole-3-pyruvic acid, exhibit also stimulating properties of plant growth and development, presumably because of conversion in the tissue to the biologically active IAA (Cohen et al., 2003). Similarly, indole-3-butyric acid, identical to IAA except for two additional methylene groups in the side chain, is effective in bioassays. Indole-3-butyric acid, originally classified as synthetic auxin, is in fact an endogenous plant compound (Bartel et al., 2001). Two, main types of synthetic plant growth regulators with high auxin activity have been described: naphthalene-1-acetic acid and 2,4-related compounds, e.g., 2,4-dichlorophenoxyacetic acid. The isomer of naphthalene-1-acetic acid, naphthalene-2-acetic acid, shows lower biological activity. 2,4-Dichlorophenoxyacetic acid is commonly used herbicide (Kelley and Riechers, 2007). 2.2. The chemistry and function of auxin conjugates Plants use several mechanisms to control the levels of endogenous auxins, especially IAA. Despite, the regulation of synthesis and degradation of these phytohormones, plants may store auxins in the form of conjugates (Fig. 2). Most of plant’s endogenous IAA is found not as a free and biologically active form, but conjugated at

Fig. 1. Selected chemical structures of natural and synthetic auxins.

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

the carboxyl group. IAA is conjugated at the carboxyl group of monosaccharides, high molecular weight polysaccharides, myoinositol, choline and the carbohydrate components of glycoproteins via ester bonds. IAA can also be conjugated to single amino acids, peptides or proteins via amide bonds. These mentioned conjugates are thought to be involved in IAA storage and transport, inactivation of the hormone pathways to auxin catabolism, and as components of a homeostasis for the control of IAA levels. They can also protect IAA against peroxidative degradation, and detoxification of excess auxin. Hydrolysis of endogenous conjugates of IAA is likely to be important free IAA sources (Cohen and Bandurski, 1982; Fluck et al., 2000). IAA conjugation is a ubiquitous process in both higher and lower plants and IAA-conjugated forms are synthesized rapidly in plant tissues when auxin homeostasis is perturbed. For example, the exposure of plants to high exogenous level of natural IAA or synthetic 2,4-dichlorophenoxyacetic acid with herbicide properties leads to synthesis of conjugates with aspartic and glucose. It appears that plants use different conjugated forms to detoxify excess auxins and to store of these phytohormones, suggesting that the conjugated moiety may dictate the fate of the attached auxin for storage, transport or degradation. These modified forms of auxins may be further oxidized for example to OxIAA-conjugates, which permanently inactivate auxins (Östin et al., 1998; Kai et al., 2007). Besides IAA, indole-3-butyric acid occurs as the free acid as well as in a variety of conjugated forms, although both are usually less abundant than IAA. Indole-3-butyric acid (IBA) is conjugated to other moieties through amide- and ester-linkages. On the other hand, ester conjugates of IBA dominate over amide forms of IBA. Moreover, IBA conjugates are more easily hydrolyzed and more slowly transported in different plant systems, perhaps leaving more phytohormones at the plant base in comparison with conjugates of IAA. In addition, certain IBA conjugates are very active in bioassays (Bartel et al., 2001; Ljung et al., 2002). The formation and hydrolysis of auxin conjugates is developmentally regulated and varies significantly among plant tissues (Rampey et al., 2004). Moreover, different plant species have distinct auxin conjugate profiles. In general, monocots appear to accumulate ester conjugates, whereas dicots accumulate mostly amide conjugates (Slovin et al., 1999; Cooke et al., 2002; Jakubowska and Kowalczyk, 2004). 2.3. The synthesis and hydrolysis of auxin ester conjugates in monocotyledonous plants The maize seedlings contain primarily ester-linked conjugates including IAA–myo-inositol, IAA–myo-inositol glycosides, IAA–glucose and a large cellulosic glucan conjugates (Fig. 3) (Cohen and Bandurski, 1982). The conjugate with glucose is the most studied: the enzymes involved both in its formation and its hydrolysis has been characterized in several plants (Jackson et al., 2002). The synthesis of IAA–glucose, followed by transacylation to myo-inositol represents two potential regulatory steps for the control of IAA concentration by converting hormonally active free IAA into growth-inactive IAA ester in monocots (Jakubowska and Kowalczyk, 2004). Glucosyltransferase from maize was the first plant protein with IAA-conjugating activity (Szerszen et al., 1994). The formation of IAA–glucose from IAA and UDP–glucose by indole-3-acetylglucose synthase is the first step in the series of reactions leading to the IAA-ester conjugates found in maize. Moreover, an Arabidopsis and maize gene encoding UDP–glucosyltransferase that conjugate IAA to glucose was identified (Jackson et al., 2001). Intracellular location of this enzyme is not clear, but the pH optimum and requirement of a redox status are consistent with being in the cyto-

959

plasm. Overexpressing gene encoding this specific UDP–glucosyltransferase renders plants resistant to exogenous IAA, and disturbs gravitropism, consistent with the role in IAA inactivation. This enzyme recognizes also indole-3-butyric acid (IBA) as a substrate and therefore is responsible for formation of IBA–glucose from radiolabelled IBA fed to Arabidopsis seedlings. Moreover, the stimulation of gene expression encoding IAA–glucose synthesis by auxin in maize coleoptiles has been observed (Kowalczyk et al., 2002). In immature maize kernels, the energetically unfavourable synthesis of IAA–glucose is followed by an energetically favourable transacylation of the IAA moiety from IAA–glucose to myo-inositol. The synthesis of IAA–myo-inositol was observed for the first time in vitro in the seeds of maize, and the transferase catalyzing this reaction was partially purified and characterized (Kesy and Bandurski, 1990). Since all maize tissues hydrolyze IAA–glucose isomers, IBA–glucose and, more slowly, IAA–myo-inositol to free IAA, the mechanism is interpreted as a shuttle to adjust the free pool of IAA via the temporal storage of IAA esters (Jakubowska and Kowalczyk, 2004). Partially purified 6-O(4-O)-IAGlc hydrolase from immature kernels of maize (Zea mays) was found to be the specific enzyme catalyzing hydrolysis of stable esters of IAA and glucose. Among a range of ester conjugates tested as substrates, only 6-O(4-O)IAA–glucose and IBA–glucose isomers were effectively hydrolyzed. The enzyme is probably involved in the regulation of the IAA levels by the target release of free auxin from ester-linked conjugates, its inactive storage forms (Jakubowska and Kowalczyk, 2005).

2.4. Auxin conjugate profile in dicotyledonous plants Amide conjugates of auxins dominate over ester conjugates in dicotyledonous plants. However, conjugate profile of endogenous auxins differs during their growth and development. For example, the level of ester conjugates and free IAA in bean declines rapidly during seed maturation, so that in fully mature seeds, the esterlinked IAA represents about 13% of the total IAA pool, and only 6% is free IAA. It is noteworthy that in seeds harvested at full maturity, IAA is conjugated to several polypeptides and proteins that approximate 80% of the total IAA pool (Walz et al., 2002). IAA undergoes conjugation to yield both IAA–glucose and amide-linked IAA, and the preferential formation of either IAA–glucose or amidelinked IAA-conjugates depends on the ripening stage of the fruit of tomato (Iyer et al., 1997). Tobacco explants and leaf protoplasts produce mainly auxin–aspartate and auxin–glucose conjugates (IAA or naphthalene-1-acetic acid conjugates) (Smulders et al., 1990). Arabidopsis, the model dicotyledonous plant, is also able to form the ester-linked IAA-conjugates that constitute approximately 8–10% of the total IAA pool. However, amide-linked IAAconjugates constitute approximately 90% of the total IAA pool in Arabidopsis, and in other dicots as well (Tam et al., 2000; Ljung et al., 2002). Many experiments have shown that conjugates of IAA with aspartate are the predominant IAA constituents present in most dicotyledonous plants, e.g., Vicia faba seedlings and tomato pericarp discs (Fig. 4). The principal IAA deactivation pathway in dicots converts IAA to N-(indole-3-acetyl)-L-aspartic acid. The indole ring of IAA–aspartate is oxidized to form N-(oxindole-3-acetyl)-L-aspartic acid, which is subjected to successive glycosylations (Iyer et al., 1997; Crozier et al., 2000). Amide conjugates with amino acids (Asp, Glu, Ala, Gly, Val and Leu) are present in a variety of plants, and conjugates with other amino acids may also occur (Fig. 5). IAA–amino acid conjugates found in plant tissues may be classified into two groups: (i) based on bioassay activity and (ii) susceptibility to hydrolysis by specific enzymes (Ljung et al., 2002).

960

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

Fig. 3. Biosynthesis and hydrolysis of indole-3-acetic acid ester conjugates in Zea mays.

The first group of amide conjugates is represented by IAA–Ala and IAA–Leu, which are characterized by biological activity in bioassays. In Arabidopsis, IAA–Ala is present at highest levels in shoots, whereas IAA–Leu accumulates in roots. Both, IAA–Ala and IAA–Leu exhibit activity because they can be readily converted via amidohydrolases to active IAA and are likely storage forms of auxins (Ljung et al., 2002). The second group of auxin amide conjugates is represented by IAA–Asp and IAA–Glu. The levels of conjugates with aspartate and glutamate are increasing after addition of IAA, making them candidates for detoxification of excess IAA (Antolic´ et al., 2001). These amide conjugates play role in IAA turnover and they are not appreciably hydrolyzed via amidohydrolases. Formation of these conjugates leads irreversibly to oxidation, followed by catabolism of IAA (Ljung et al., 2002). Moreover, IAA–Glu and IAA–Asp

are not characterized by biological activity in plant species tested (Slovin et al., 1999). On the other hand, the conjugate of IAA with aspartate and its downstream metabolites must have an important function, because a mutant cell line (XIIB2) of Egyptian henbane (Hyoscyamus muticus L.) impaired in IAA–Asp biosynthesis rapidly dies at 33 °C, a temperature to which, otherwise isogenic, wildtype cell cultures are resistant (Oetiker and Aeschbacher, 1997). 2.5. Synthesis and hydrolysis of amide conjugates of auxins The auxin levels in plants are modulated by a specific group of amidohydrolases from the peptidase M20D family that release the active hormone from its conjugated storage forms. For example, IAA–amino acid hydrolase IAA–leucine (ILL2) from Arabidopsis thaliana preferentially hydrolyses the auxin–amino acid conjugate N-

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

961

Fig. 4. Nondecarboxylative catabolism and conjugation of indole-3-acetic acid in the seedlings of Vicia faba and tomato (Lycopersicon esculentum) pericarp discs.

(indol-3-acetyl)-alanine. The overall structure of ILL2 is reminiscent of dinuclear metallopeptidases from the M20 peptidase family (Bitto et al., 2009). The conjugate hydrolysis is the principal source of auxins in germinating seed and seedling development. An increased auxin pool after amide conjugate hydrolysis appears also to correlate with cell expansion and cell division. Therefore, these conjugated forms of auxins contribute to the pool of active free auxins (Rampey et al., 2004; Schuller and Ludwig-Müller, 2006). The conjugate hydrolases have different amino acid specificities both in vivo and in vitro when tested with battery of IAA–amino acid conjugates. They are also differentially expressed, implying that a variety of IAA-conjugates exist and serve different roles at varied location throughout the plant (Cohen and Bandurski,

1982). For example, IAA–Ala is hydrolyzed by IAR3–amidohydrolase (IAA–amino acid amidohydrolase) isolated from Arabidopsis and Brassica rapa (Schuller and Ludwig-Müller, 2006). On the other hand, amidohydrolase TaIAR3 isolated from monocotyledonous species wheat (Triticum aestivum) was found to hydrolyze negligible levels of IAA–Ala and no other IAA amino acid conjugates tested. TaIAR3 has also low specificity for the ester conjugates IAA–Glc and IAA–myo-inositol and high specificity for the conjugates of indole-3-butyric acid (IBA–Ala and IBA–Gly) and indole3-propionic-acid (IPA–Ala) (Campanella et al., 2004). Moreover, the regulation of auxin conjugate hydrolysis in Medicago truncatula during development and interaction with two symbionts was characterized by Campanella et al. (2008). In this plant five putative

962

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

Fig. 5. The structures of amide auxin conjugates.

auxin amidohydrolase genes (MtIAR31, MtIAR32, MtIAR33, MtIAR34, and MtIAR36) were identified, homologous to the AtIAR3 gene from A. thaliana. MtIAR31, -32, -33, and -34 have hydrolytic activity against IAA–Asp and IBA–Ala; MtIAR33, -34, and -36 hydrolyze the ester bonds of IAA–Glc. MtIAR36 solely possesses activity against IAA–Gly, –Ala, and –Ile (Campanella et al., 2008). IAA–Leu is hydrolyzed by the Arabidopsis ILR1–amidohydrolase (IAA–amino acid amidohydrolase) which requires Mn or Co for its activity and is predicted to be localized to the endoplasmic reticulum lumen. Additionally, another protein, ILR3, is an apparent transcription factor important for IAA-conjugate responsiveness. These data suggested that ILR3 might regulate ILR1 activity because ILR3 may be directly or indirectly involved in metal homeostasis, especially metal cofactors availability (Mn, Co, and Fe) which are necessary for ILR1 activity (Rampey et al., 2006). Examined IAA–amidohydrolases in these plant species are targeted to the endoplasmic reticulum lumen, although the significance of such compartmentation is unclear because there is no clear evidence on the storage location of any conjugate compounds (Rampey et al., 2004; Woodward and Bartel, 2005). The enzymes that conjugate IAA to amino acids were found in Arabidopsis. Those enzymes are in the luciferase superfamily and are related to the JAR1 (jasmonate:amino acid synthetase) enzyme that conjugates other plant hormone – jasmonic acid (JA) to amino acids, and are encoded by members of the GH3 family of auxin-induced genes. The Arabidopsis GH3 gene family is composed of 20 members that encode proteins capable of conjugating phytohormones such as auxin, jasmonates and salicylic acid to amino acid. Accumulating evidence indicates that GH3 family members func-

tion to regulate levels biologically active hormones through amino acid conjugation, thereby targeting them for degradation/storage in the case of auxins and activating them in the case of jasmonates (Staswick et al., 2005). The characterized GH3-like enzymes apparently prefer to synthesize biologically inactive conjugates of IAA (IAA–Asp, IAA–Glu) over hydrolysable and active in bioassay forms of amide conjugates (IAA–Ala, IAA–Leu) in vitro (Staswick et al., 2005; Khan and Stone, 2007). Auxin substrate for GH3 enzyme includes also indole-3-butyric acid, indole-3-pyruvic acid, phenylacetic acid, 1-naphthalacetic acid. Experiments performed on Populus  canescens indicated that GH3 became active when the plants were exposed to abiotic stresses, like bending or salinity, causing changes in wood anatomy suggesting that adjustment of the internal auxin balance in wood in response to environmental cues involves GH3 auxin conjugate synthases (Teichmann et al., 2008). Moreover, amino acid conjugates of these phytohormones can also be glucosylated (Ljung et al., 2002; Zazimalova and Napier, 2003). Recently, a rice (Oryza sativa) GH3-8 gene encoding IAA– amino acid synthetase has been shown to promote basal immunity in rice by converting active IAA to inactive IAA–Asp and thus reducing the auxin-induced cell wall loosening (Ding et al., 2008). The data obtained by Ludwig-Müller et al. (2009) suggested a developmentally controlled involvement of GH3 proteins in auxin homeostasis by conjugating excess of physiologically active free auxin to inactive IAA–amide conjugates in lower plant – moss Physcomitrella patens. Targeted knock-out of two GH3 genes (DeltaPpGH3-1/DeltaPpGH3-2) exhibited an increased sensitivity to auxin, resulting in growth inhibition. On plain mineral media mutants of moss had higher levels of free IAA and less conjugated IAA

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

than the wild type, and this effect was enhanced when auxin was supplied. The DeltaPpGH3-1/DeltaPpGH3-2 double knockout had almost no IAA amide conjugates but still synthesized ester conjugates. Therefore, the control of auxin levels is also achieved in lower plants via synthesis of auxin conjugates by members of the GH3 family. 2.6. Peptide and methylate conjugates of auxins Among amide conjugates of IAA, a 35-kDa IAA-peptide was identified in Arabidopsis seeds. The presence of peptide-bound IAA has been also demonstrated in bean seeds and strawberry fruit. Achenes and receptacle tissue of Fragaria vesca were shown to contain conjugated indole-3-acetic acid (IAA) that was not soluble in organic solvents and yielded IAA after strong alkaline hydrolysis, suggestive of IAA attached to plant proteins (76 kDa). Biochemical analysis indicated that protein may function as a chaperonin related to the hsp60 class of proteins or, alternatively, an ATP synthase (Park et al., 2006). The large size of this conjugate might contribute to the solvent insolubility of amino acid conjugates. Moreover, bean seeds apparently lack amino acid conjugates, and IAA is instead conjugated to several polypeptides ranging size of molecular weight from 3 to 60 kDa. Although, genes encoding peptides that conjugate to auxins have not been identified yet, one of these IAA-modified bean protein is similar to a soybean late seed maturation protein, suggesting that certain seed storage proteins may function in both amino acid and phytohormones storage (Walz et al., 2002). Another conjugation process of auxin leads to methylation of IAA (MeIAA). An enzyme that methylates the carboxyl side chain of IAA is a member of a family of carboxyl methyltransferases which methylate various plant hormones, such as jasmonates and salicylates. It was recently shown that IAA could be converted to its methyl ester by the Arabidopsis enzyme IAA carboxyl methyltransferase 1 (IAMT1). Furthermore, disruption of the expression levels of IAMT1 led to phenotypes indicative of disruption of IAA homeostasis (Qin et al., 2005). MeIAA has rarely been reported as an endogenous IAA metabolite in plants, probably due to its low abundance or fast turnover, and therefore its in vivo function remains unknown. MeIAA had been used as a substitute for IAA in physiological studies, and it has been observed that various auxin signaling mutants show decreased sensitivity to exogenously applied MeIAA as well as to IAA (Qin et al., 2005), suggesting that MeIAA and IAA share similar signaling components. Therefore, either MeIAA itself could initiate the auxin signaling pathway, or it must be hydrolyzed to IAA to exert hormonal function. Methylation may increase the volatility of IAA, but its function in vivo is not clear (Woodward and Bartel, 2005). Inactive methyl indole-3acetic acid ester can be hydrolyzed and activated by several esterases belonging to the AtMES esterase family of Arabidopsis (Yang et al., 2008).

3. Cytokinin conjugates 3.1. The structure, role and metabolism of cytokinins Cytokinins are a class of phytohormones that play an important role at all phases of plant development from seed germination to senescence. They act at the cellular level by inducing expression of some genes, promotion mitosis and chloroplast development but also on the organ level by releasing buds from apical dominance or by inhibiting root growth (Riefler et al., 2006). The naturally occurring cytokinins are N6-substituted adenine derivatives that contain an isoprenoid or an aromatic derivative side chain (Fig. 6). Among isoprenoid cytokinins, trans-zeatin is

963

considered central due to its general occurrence and high activity in the most bioassays. Its stereoisomer, cis-zeatin, is characterized by weak activity in bioassays. However, cis-isomers can be dominant cytokinins at particular stages of development in plants such as Cicer arietinum, and Lupinus albus. Dihydrozeatin, and N6-(D2isopentenyl)-adenine are also commonly present in lower and vascular plants (Emery et al., 1998, 2000; Sakakibara, 2006; Stirk et al., 2008). N6-Benzyladenine and its derivatives, representing aromatic cytokinins, have been detected in a number of plant species as minor components of the total cytokinins. Hydroxylated derivatives of N6-benzyladenine in meta or ortho position of benzyl group are commonly named as meta- and ortho-topolin, respectively (Strnad et al., 1997). Kinetin, the most known cytokinin, has furfuryl ring at the N6-position of adenine and was identified in both animal cellular DNA and plant tissue extracts (Barciszewski et al., 2000). Natural cytokinins behave as normal adenylate compounds in that they exist in the plant cell as mixtures of free bases, nucleosides (Fig. 7), as well as mono-, di-, and trinucleotides in apparent equilibrium. All forms of cytokinins may be reversible or irreversible conjugated with sugars, and amino acids. In most bioassays, cytokinin bases are the most active, and therefore cytokinin conjugation contributes to regulation their activity. Cytokinin conjugates seem to serve as storage, transport, and deactivated forms because they are resistant to degradation by cytokinin oxidase/ dehydrogenase (Auer, 2002; Blagoeva et al., 2004). Cytokinin conjugate formation can be divided into two major categories: conjugation with the adenine moiety or the side chain. 3.2. Conjugation of adenine ring of cytokinins The most common modifications of the adenine molecule are Nglucosylation, and N9-alanine conjugation. The adenine ring system can be glucosylated at the N3-, N7- and N9-position (Fig. 8). Glucosyl conjugates at N7- and N9-position but not at the N3-position of cytokinins such as trans-zeatin, dihydrozeatin as well as N6-(D2-isopentenyl)-adenine are usually inactive in the most of bioassays because their active free forms cannot be released by hydrolysis. The relatively high activity of 3-b-glucosyldihydrozeatin in bioassay most probably reflects the presence of specific enzymes in the plant system studied which can convert its back to free active base. Additionally, N-glucosylated cytokinins may accumulate to even higher levels than the free bases under normal circumstances as well as stress conditions. However, the precise in vivo function of these cytokinin metabolites remains still unknown (Mok and Mok, 2001; Veach et al., 2003). Moreover, among all cytokinin types of conjugates present in macroalgae, N-glucosides have not been detected (Stirk et al., 2003). The results suggest that different pathways for regulating cytokinin concentrations operate in macroalgae than in higher plants. The formation of cytokinin glucosides at the N7- and N9-position is catalyzed in Arabidopsis by two specific glucosyltransferases (UGT76C1 and UGT76C2). However, both mentioned enzymes prefer glucosylation at N7- to that N9-, which corresponds well with higher concentrations of various N7-glucosylated forms of cytokinins in Arabidopsis plastids (Sakakibara, 2006). Although the enzymes recognize a large number of adenine derivatives as substrates, the rate of N-glucosylation is higher for compounds with N6-side chains of at least three alkyl carbons and roughly correlates with cytokinin activity. Both, UDP–Glc and TDP–Glc can serve as glucosyl donors for formation of N-glucosylated cytokinins (Brzobohaty et al., 1993). Less common is alanine conjugation at N9-position of adenine moiety (Fig. 9). 9-Alanylzeatin and 9-alanyldihydrozeatin, commonly known as lupininc acid and dihydrolupininc acid,

964

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

Fig. 6. Selected chemical structures of naturally occurring cytokinins.

Fig. 7. Possible interconversion of cytokinin bases, nucleosides, and nucleotides.

respectively, were identified in lupin (Lupinus angustifolius) seeds (Duke et al., 1978). A specific transferase which catalyses the conversion of trans-zeatin, and dihydrozeatin to their N9-alanyl derivatives was discovered and purified from lupin seeds. The donor substrate is O-acetyl-L-serine. Alanine conjugates of isoprenoid cytokinins are characterized by low activity in bioassay because the lack of enzyme systems responsible for hydrolysis to active forms (Entsch et al., 1983; Mok and Mok, 2001).

3.3. Conjugation of isoprenoid chain of cytokinins Conjugation of cytokinins involves also O-glycosylation, and Oacetylation at the hydroxyl group of the side chains of cytokinins (Fig. 10) (Martin et al., 1999). The O-glycosylation of these phytohormones was extensively studied in different plant systems. Oglucosyl conjugates of isoprenoid and aromatic cytokinins are commonly present metabolites in various species of macroalgae

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

965

Fig. 8. N-Glucosylated conjugates of cytokinins.

Fig. 9. Amino acid (alanine) conjugates of cytokinins.

(Stirk et al., 2003). Among the lower plants, the mosses have emerged as a model for studying cytokinin metabolism because genetic mutants have been isolated, and they are amenable to physiological and genetic studies. For example, bryophyte Physcomitrella patents OVE mutants are characterized by overproduction of gametophores and cytokinins. The analysis of cytokinin profile of this moss revealed that cis-zeatin-riboside-O-glucoside and trans-zeatin-riboside-O-glucoside were the most abundant intracellular conjugates of cytokinins (von Schwartzenberg et al., 2007). The current development of molecular techniques and genetic transformation of mosses holds promise for further work on synthesis and hydrolysis of glucosyl conjugates of cytokinins. In contrast to the research on mosses, there is very little information on liverworts and hornworts and no endogenous cytokinin conjugates have been identified. O-Glucosylyated forms of isoprenoid and aromatic cytokinins are the most commonly found in higher plants, although, the presence of O-xylosyl conjugates has been only observed in Phaseolus species (Martin et al., 2000). Both, O-glucosylated and O-xylosylated cytokinins are considered important for storage, transport, and protection against degrading enzymes. Additionally, these metabolites can be easily converted into active cytokinin by specific b-glucosidases (Brzobohaty et al., 1993). Thus, it is believed that O-glucosides and O-xylosides of these phytohormones play an important role in cytokinin homeostasis. Moreover, O-glycosylation of cytokinins may modify the activity of cytokinins. For in-

stance, O-b-glucosylzeatin, and O-b-xylosylzeatin effectively stimulated callus growth in bean (P. vulgaris) and Lima bean (P. lunatus). These O-glycosides are relatively stable in plant tissues but they can be easily converted back to active trans-zeatin. The level of sugar derivatives shows also large fluctuations during plant development. For example, O-b-glucosylzeatin was the predominant cytokinin in fully developed leaves of Urtica dioica, whereas the dominance of free trans-zeatin was observed in young leaves (Wagner and Beck, 1992). The fact that O-glycosides may be needed for stored cytokinins would mean that trans-zeatin biosynthesis does not necessarily coincide with the cytokinin requirement of the plant tissue. These conjugates are mainly localized in plant vacuoles, however the existence of mechanism controlling their transport across membranes when the need for cytokinin increases, is still unknown (Mok et al., 1992). Moreover, it has been reported that the ratio of cytokinin conjugation process with glucose and xylose increases under the influence of stress conditions, for example in the presence of high concentrations of heavy metals such as Pb, Cu, Zn and Al as well as heat stress (Atanasova et al., 2004; Wang et al., 2004). In previous reports for tobacco callus tissue, levels of O-glucosides were also found to be markedly increased upon expression or de-repression of ipt gene encoding a key enzyme in cytokinin biosynthetic pathway (Redig et al., 1996). The transfer of glycosyl moiety from an activated glycosyl donor to hydroxyl group in the side chain of cytokinin can be mediated by specific glycosyltransferase enzymes (Fig. 11) (Auer, 2002). The cytokinin glycosyltransferases are UDP–Glc- or UDP–Xyl-requiring enzymes and belong to family 1 of the 68 families of various glycosyltransferases. These enzymes are localized mainly in plant vacuoles. Zeatin O-glucosyltransferase has been isolated from Lima bean (P. lunatus), soybean (Glycine max), rice (O. sativa), and tomato (Lycopericon esculentum), whereas zeatin O-xylosyltransferase was identified and purified from bean (P. vulgaris). The zeatin O-glucosyltransferase uses UDP–Glc and UDP–Xyl as donor substrates but has much higher affinity to UDP–Glc, whereas the zeatin O-xylosyltransferase exclusively utilizes UDP–Xyl (Mok and Mok, 2001; Meek et al., 2008). The cytokinin substrate recognition is also highly specific because only trans-zeatin, and dihydrozeatin, and their respective

966

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

Fig. 10. O-Glucosyl and O-acetyl conjugates of cytokinins.

nucleosides are O-glucosylated and O-xylosylated (Fig. 11). The stringent substrate specificity for cytokinins and sugar donor suggests that conjugation process with glucose and xylose is precisely regulated during plant development. Moreover, the ribose moiety of cytokinin nucleosides can also be O-glucosylated, forming O-bglucosyl-9-ribosylzeatin, and O-b-glucosyl-9-ribosyldihydrozeatin (Martin et al., 1999). The O-glucosylation of cytokinins is stereo-specific. The O-glucosyltransferase encoded by the P. lunatus ZOG1 gene has high affinity for trans-zeatin as the substrate, whereas the enzyme en-

coded by the maize (Z. mays) gene prefers cis-zeatin (Veach et al., 2003). The cis-zeatin-O-glucosyltransferase isolated from maize root specifically glucosylates cis-zeatin, and 9-ribosyl-cis-zeatin but do not recognize as a substrate trans-zeatin and dihydrozeatin. Therefore, it could be assumed that cis-zeatin may have a greater importance in plant growth and development than previously believed (Martin et al., 2001). Additionally, it was found that hydroxylated derivatives of N6benzyladenine (topolins) are also O-glucosylated (Fig. 10). Structurally different isomers of topolins are recognized by distinct

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

967

Fig. 11. The conjugation and de-conjugation of the cytokinin N6-isoprenoid side chain.

plant O-glucosyltransferases. Therefore, meta-topolin is the preferred substrate of zeatin O-glucosyltransferase, whereas cis-zeatin-O-glucosyltransferase recognizes ortho-topolin as the substrate. The O-glucosides of meta- and ortho-topolins are characterized by high activity in the P. lunatus callus bioassay. Probably, they can be easily converted back to active topolins. However, para-topolin cannot be metabolized by O-glucosyltransferases, and does not possess the ability to promote growth of P. lunatus (Strnad et al., 1997; Mok et al., 2005). De-conjugation process of O-glucosylated cytokinins involves the conversion of glucosides and xylosides to the correspondent active aglycones. This reaction is catalyzed by various substratespecific b-glucosidase enzymes. Experiments have been shown that b-glucosidases utilize a broad spectrum of substrates, not only cytokinin conjugate. At this time, the only reported b-glucosidase from maize seedling leaves and root cells can hydrolyze endogenous cytokinin conjugates with sugars as well as number of other artificial and natural compounds (Vévodová et al., 2001; Gu et al., 2006). For example, the b-glucosidase isolated from maize and encoded by Zm-p60.1 gene cleaved the biologically inactive hormone conjugates such as O-b-glucosylzeatin and 3-b-glucosylkinetin,

releasing active cytokinins. Tobacco protoplasts that transiently expressed Zm-p60.1 gene could use the inactive cytokinin glucosides to initiate cell division. Product of Zm-p60.1 gene was localized to the meristematic maize cells and may function in vivo to supply the developing embryo with active forms of cytokinins (Brzobohaty et al., 1993). The low specificity of b-glucosidases to substrates suggests that the hydrolysis of cytokinin conjugates is not highly regulated, in contrast to the precise control of O-glycoside synthesis (Mok and Mok, 2001). In addition to O-glucosylated cytokinins, the O-acetylation of side chain of cytokinins is less common in plants (Fig. 10). For example, the O-acetylation of trans-zeatin has been observed in a few lupine species. This process does not reduce drastically stimulating properties of cytokinins. Probably, the ability of O-acetylzeatin to promote the growth and development of P. lunatus may be derived from the conversion of this conjugate to free trans-zeatin. Although O-acetylzeatin was identified as naturally occurring metabolite, its significance and synthesis are still unknown (Letham and Zhang, 1989). In various lower and vascular plant species most auxins and cytokinins are thought to be present in both free and as conjugated

968

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969

forms. Considerable efforts have been directed at clarifying the conjugation and de-conjugation as well as factors contributing to auxin and cytokinin homeostasis, but the entire picture remains to be elucidated. References Antolic´, S., Kveder, M., Klaic´, B., Magnus, V., Kojic´-Prodic´, B., 2001. Recognition of the folder conformation of plant hormone (auxin, IAA) conjugates with glutamic and aspartic acid and their amides. J. Mol. Struct. 560, 223–237. Atanasova, L., Pissarska, M.G., Popov, G.S., Georgiev, G.I., 2004. Growth and endogenous cytokinins of juniper shoots as affected by high metal concentrations. Biol. Plant. 48, 157–159. Auer, C.A., 2002. Discoveries and dilemmas concerning cytokinin metabolism. J. Plant Growth Regul. 21, 24–31. Barciszewski, J., Siboska, G., Clark, B.F.C., Rattan, S.I.S., 2000. Cytokinin formation by oxidative metabolism. J. Plant Physiol. 158, 587–588. Bartel, B., LeClere, S., Magidin, M., Zolman, B.K., 2001. Inputs to the active indole-3acetic acid pool: de novo synthesis, conjugate hydrolysis, and indole-3-butyric acid b-oxidation. J. Plant Growth Regul. 20, 198–216. Bitto, E., Bingman, C.A., Bittova, L., Houston, N.L., Boston, R.S., Fox, B.G., Phillips, G.N., 2009. X-ray structure of ILL2, an auxin-conjugate amidohydrolase from Arabidopsis thaliana. Proteins 74, 61–71. ˇ ková, R., 2004. Blagoeva, E., Dobrev, P., Malbeck, J., Motyka, V., Gaudinova, A., Van Effect of exogenous cytokinins, auxins and adenine on cytokinin Nglucosylation and cytokinin oxidase/dehydrogenase activity in de-rooted radish seedlings. Plant Growth Regul. 44, 15–23. Brzobohaty, B., Moore, I., Kristoffersen, P., Bako, L., Campos, N., Schell, J., Palme, K., 1993. Release of active cytokinin by a b-glucosidase localized to the maize root meristem. Science 262, 1051–1054. Campanella, J.J., Olajide, A.F., Magnus, V., Ludwig-Müller, J., 2004. A novel auxin conjugate hydrolase from wheat with substrate specificity for longer side-chain auxin amide conjugates. Plant Physiol. 135, 2230–2240. Campanella, J.J., Smith, S.M., Leibu, D., Wexler, S., Ludwig-Müller, J., 2008. The auxin conjugate hydrolase family of Medicago truncatula and their expression during the interaction with two symbionts. J. Plant Growth Regul. 27, 26–38. Cohen, J.D., Bandurski, R.S., 1982. Chemistry and physiology of the bound auxins. Ann. Rev. Plant Physiol. 33, 403–430. Cohen, J.D., Slovin, J.P., Hendrickson, A.M., 2003. Two genetically discrete pathways convert tryptophan to auxin: more redundancy in auxin biosynthesis. Trends Plant Sci. 8, 197–199. Cooke, T.J., Poli, D.B., Sztein, A.E., Cohen, J.D., 2002. Evolutionary patterns in auxin action. Plant Mol. Biol. 49, 319–338. Crozier, A., Kamiya, Y., Bishop, G., Yokota, T., 2000. Biosynthesis of hormones and elicitor molecules. In: Buchanan, B., Gruissem, W., Jones, R. (Eds.), Biochemistry and Molecular Biology of Plants. American Society of Plant Physiologists, Rockville, USA, pp. 850–929. Ding, X., Cao, Y., Huang, L., Zhao, J., Xu, C., Li, X., Wang, S., 2008. Activation of the indole-3-acetic acid amido synthetase GH3-8 suppresses expansion expression and promotes salicylate- and jasmonate-independent basal immunity in rice. Plant Cell 20, 228–240. Duke, C.C., MacLeod, J.K., Summons, R.E., Letham, D.S., Parker, C.W., 1978. The structure and synthesis of cytokinin metabolites. II. Lupinic acid and O-b-Dglucopyranosylzeatin from Lupinus angustifolius. Aust. J. Chem. 31, 1291–1301. Emery, R.J.N., Leport, L., Barton, J.E., Turner, N.C., Atkins, C.A., 1998. cis-Isomers of cytokinins predominate in chickpea seeds throughout their development. Plant Physiol. 117, 1515–1523. Emery, R.J.N., Ma, Q., Atkins, C.A., 2000. The forms and sources of cytokinins in developing white lupine seeds and fruits. Plant Physiol. 123, 1593–1604. Entsch, B., Parker, C.W., Letham, D.S., 1983. An enzyme from lupin seeds forming alanine derivatives of cytokinins. Phytochemistry 22, 375–381. Fluck, R.A., Leber, P.A., Lieser, J.D., Szczerbicki, S.K., Varnes, J.G., Vitale, M.A., Wolfe, E.E., 2000. Choline conjugates of auxins. I. Direct evidence for the hydrolysis of choline–auxin conjugates by pea cholinesterase. Plant Physiol. Biochem. 38, 301–308. Gu, R., Zhao, L., Zhang, Y., Chen, X., Bao, J., Zhao, J., Wang, Z., Fu, J., Liu, T., Wang, J., Wang, G., 2006. Isolation of a maize beta-glucosidase gene promoter and characterization of its activity in transgenic tabacco. Plant Cell Rep. 25, 1157– 1165. Hagen, G., Guilfoyle, T., 2002. Auxin-responsive gene expression: genes, promoters and regulatory factors. Plant Mol. Biol. 49, 373–385. Iyer, M., Cohen, J.D., Slovin, J.P., 1997. Molecular manipulation of IAA metabolism in tomato. Plant Physiol. 114S, 158. Jackson, R.G., Lim, E.K., Li, Y., Sandberg, G., Hoggett, J., Ashford, D.A., Bowles, D.J., 2001. Identification and biochemical characterization of an Arabidopsis indole3-acetic acid glucosyltransferase. J. Biol. Chem. 276, 4350–4356. Jackson, R.G., Kowalczyk, M., Li, Y., Higgings, G., Ross, J., Sandberg, G., Bowles, D.J., 2002. Over-expression of an Arabidopsis gene encoding a glucosyltransferase of indole-3-acetic acid; phenotypic characterization of transgenic lines. Plant J. 32, 573–583. Jakubowska, A., Kowalczyk, S., 2004. The auxin conjugate 1-O-indole-3-acetyl-b-Dglucose is synthesized in immature legume seeds by IAGlc synthase and may be used for modification of some high molecular weight compounds. J. Exp. Bot. 55, 791–801.

Jakubowska, A., Kowalczyk, S., 2005. A specific enzyme hydrolyzing 6-O(4-O)indole-3-ylacetyl-beta-D-glucose in immature kernels of Zea mays. J. Plant Physiol. 162, 207–213. Kai, K., Horita, J., Wakasa, K., Miyagawa, H., 2007. Three oxidative metabolites of indole-3-acetic acid form Arabidopsis thaliana. Phytochemistry 68, 1651– 1663. Kelley, K.B., Riechers, D.E., 2007. Recent developments in auxin biology and new opportunities for auxinic herbicide research. Pestic. Biochem. Physiol. 89, 1–11. Kesy, J.M., Bandurski, R.S., 1990. Partial purification and characterization of indol-3ylacetylglucose: myo-inositol indol-3-ylacetyltransferase (indoleacetic acidinositol synthase). Plant Physiol. 94, 1598–1604. Khan, S., Stone, J.M., 2007. Arabidopsis thaliana GH3.9 influences primary root growth. Planta 226, 21–34. Kowalczyk, S., Jakubowska, A., Bandurski, R.S., 2002. 1-Naphthalene acetic acid induces indole-3-ylacetyl-glucose synthase in Zea mays seedlings tissue. Plant Growth Regul. 38, 127–134. Letham, D.S., Zhang, R., 1989. Cytokinin translocation and metabolism in lupin species. II. New nucleotide metabolites of cytokinins. Plant Sci. 64, 161–165. Ljung, K., Hull, A.K., Kowalczyk, M., Marchant, A., Celenza, J., Cohen, J.D., Sandberg, G., 2002. Biosynthesis, conjugation, catabolism and homeostasis of indole-3acetic acid in Arabidopsis thaliana. Plant Mol. Biol. 49, 249–272. Ludwig-Müller, J., Cohen, J.D., 2002. Identification and quantification of three active auxins in different tissues of Tropaeolum majus. Physiol. Plant. 115, 320–329. Ludwig-Müller, J., Jülke, S., Bierfreund, N.M., Decker, E.L., Reski, R., 2009. Moss (Physcomitrella patens) GH3 proteins act in auxin homeostasis. New Phytol. 181, 323–338. Martin, R.C., Mok, M.C., Mok, D.W.S., 1999. Isolation of a cytokinin gene, encoding zeatin O-glucosyltransferase from Phaseolus lunatus. Proc. Natl. Acad. Sci. USA 96, 284–289. Martin, R.C., Cloud, K.A., Mok, M.C., Mok, D.W.S., 2000. Substrate specificity and domain analysis of zeatin O-glucosyltransferases. J. Plant Growth Regul. 32, 289–293. Martin, R.C., Mok, M.C., Habben, J.E., Mok, D.W.S., 2001. A maize cytokinin gene encoding an O-glucosyltransferase specific to cis-zeatin. Proc. Natl. Acad. Sci. USA 98, 5922–5926. Meek, L., Martin, R.C., Shan, X., Karplus, P.A., Mok, D.W.S., Mok, M.C., 2008. Isolation of legume glycosyltransferases and active site mapping of the Phaseolus lunatus zeatin O-glucosyltransferase ZOG1. J. Plant Growth Regul. 27, 192–201. Mok, D.W.S., Mok, M.C., 2001. Cytokinins metabolism and actions. Ann. Rev. Plant Physiol. Plant Mol. Biol. 52, 89–119. Mok, D.W.S., Mok, M.C., Martin, R.C., Bassil, N., Shaw, G., 1992. Immuno-analyses of zeatin metabolic enzymes of Phaseolus. In: Kaminek, M., Mok, D.W.S., Zazˇimalová, E. (Eds.), Physiology and Biochemistry of Cytokinins in Plants. SPB Academic Publishing, The Hague, pp. 17–25. Mok, M.C., Martin, R.C., Dobrev, P.I., Vanková, R., Ho, P.S., Sakakibara, K.Y., Sakakibara, H., Mok, D.W.S., 2005. Topolins and hydroxylated thidiazuron derivatives are substrates of cytokinin O-glucosyltransferase with position specificity related to receptor recognition. Plant Physiol. 137, 1057–1066. Nordström, A., Tarkowski, P., Tarkowska, D., Norbaek, R., Ästot, C., Dolezal, K., Sandberd, G., 2004. Auxin regulation of cytokinin biosynthesis in Arabidopsis thaliana: a factor of potential importance for auxin–cytokinin-regulated development. Proc. Natl. Acad. Sci. USA 21, 8039–8044. Oetiker, J.H., Aeschbacher, G., 1997. Temperature-sensitive plant cells with shunted indole-3-acetic acid conjugation. Plant Physiol. 114, 1385–1395. Östin, A., Kowalczyk, M., Bhalerao, R.P., Sandberg, G., 1998. Metabolism of indole-3acetic acid in Arabidopsis. Plant Physiol. 118, 285–296. Park, S., Cohen, J.D., Slovin, J.P., 2006. Strawberry fruit protein with a novel indoleacyl modification. Planta 224, 1015–1022. Qin, G., Gu, H., Zhao, Y., Ma, Z., Shi, G., Yang, Y., Pichersky, E., Chen, H., Liu, M., Chen, Z., 2005. Regulation of Arabidopsis leaf development by an indole-3-acetic acid carboxyl methyltransferase in Arabidopsis. Plant Cell 17, 2693–2704. Rampey, R.A., LeClere, S., Kowalczyk, M., Ljung, K., Sandberg, G., Bartel, B., 2004. A family of auxin-conjugate hydrolases that contributes to free indole-3-acetic acid levels during Arabidopsis germination. Plant Physiol. 135, 978–988. Rampey, R.A., Woodward, A.W., Hobbs, B.N., Tierney, M.P., Lahner, B., Salt, D.E., Bartel, B., 2006. An Arabidopsis basic helix-loop-helix leucine zipper protein modulates metal homeostasis and auxin conjugate responsiveness. Genetics 174, 1841–1857. Redig, P., Shaul, O., Inze, D., van Montagu, M., van Onckelen, H., 1996. Levels of endogenous cytokinins, indole-3-acetic acid and abscisic acid during the cell cycle of synchronized tobacco BY-2 cells. FEBS Lett. 391, 175–180. Riefler, M., Novak, O., Strnad, M., Schmülling, T., 2006. Arabidopsis cytokinin receptor mutants reveal functions in shoot growth, leaf senescence, seed size, germination, root development, and cytokinin metabolism. Plant Cell 18, 40– 54. Sakakibara, H., 2006. Cytokinins: activity, biosynthesis and translocation. Ann. Rev. Plant Biol. 57, 431–449. Schuller, A., Ludwig-Müller, J., 2006. A family of auxin conjugate hydrolases from Brassica rapa: characterization and expression during clubroot disease. New Phytol. 171, 145–157. Slovin, J.P., Bandurski, R.S., Cohen, J.D., 1999. Auxin. In: Hooykaas, P.J.J., Hall, M.A., Libbenga, K.R. (Eds.), Biochemistry and Molecular Biology of Plant Hormones. Elsevier Science, London, pp. 115–140. Smulders, M.J.M., Van de Ven, E.T.W.M., Croes, A.F., Wullems, G.J., 1990. Metabolism of 1-naphthaleneacetic acid in explants of tobacco: evidence for release of free hormone from conjugates. J. Plant Growth Regul. 9, 27–34.

A. Bajguz, A. Piotrowska / Phytochemistry 70 (2009) 957–969 Staswick, P.E., Serban, B., Rowe, M., Tiryaki, I., Maldonado, M.T., Maldonado, M.C., Suza, W., 2005. Characterization of an Arabidopsis enzyme family that conjugates amino acids to indole-3-acetic acid. Plant Cell 17, 616–627. Stirk, W.A., Novák, O., Strnad, M., van Staden, J., 2003. Cytokinins in macroalgae. Plant Growth Regul. 41, 13–24. Stirk, W.A., Novák, O., Václavíková, K., Tarkowski, P., Strnad, M., van Staden, J., 2008. Spatial and temporal changes in endogenous cytokinins in developing pea roots. Planta 227, 1279–1289. Strnad, M., Hanuš, J., Vaneˇk, T., Kaminek, M., Ballantine, J., Fussell, B., Hanke, D.E., 1997. meta-Topolin, a highly active aromatic cytokinin from poplar leaves (Populus  canadensis Moench., cv. Robusta). Phytochemistry 45, 213–218. Szerszen, J.B., Szczyglowski, K., Bandurski, R.S., 1994. iaglu, a gene from Zea mays involved in conjugation of growth hormone indole-3-acetic acid. Science 265, 1699–1701. Tam, Y.Y., Epstein, E., Normanly, J., 2000. Characterization of auxin conjugates in Arabidopsis. Low steady-state levels of indole-3-acetyl-aspartate, indole-3acetyl-glutamate, and indole-3-acetyl-glucose. Plant Physiol. 123, 589–595. Teichmann, T., Bolu-Arianto, W.H., Olbrich, A., Langenfeld-Heyser, R., Göbel, C., Grzeganek, P., Feussner, I., Hänsch, R., Polle, A., 2008. GH3::GUS reflects cellspecific developmental patterns and stress-induced changes in wood anatomy in the poplar stem. Tree Physiol. 28, 1305–1315. Veach, Y.K., Martin, R.C., Mok, D.W.S., Malbeck, J., Vanˇková, R., Mok, M.C., 2003. OGlucosylation of cis-zeatin in maize. Characterisation of genes, enzymes and endogenous cytokinins. Plant Physiol. 131, 1374–1380. Vévodová, J., Marek, J., Zouhar, J., Brzobohaty´, B., Su, X.D., 2001. Purification, crystallization and preliminary X-ray analysis of a maize cytokinin glucoside specific beta-glucosidase. Acta Crystallogr. D Biol. Crystallogr. 57, 140–142. von Schwartzenberg, K., Núñez, M.F., Blaschke, H., Dobrev, P.I., Novák, O., Motyka, V., Strnad, M., 2007. Cytokinins in the bryophyte Physcomitrella patens: analyses of activity, distribution, and cytokinin oxidase/dehydrogenase overexpression reveal the role of extracellular cytokinins. Plant Physiol. 145, 786–800. Wagner, B.M., Beck, E., 1992. Cytokinins in Urtica dioica plants: production, metabolic and fluxes. In: Kaminek, M., Mok, D.W.S., Zazˇimalová, E. (Eds.), Physiology and Biochemistry of Cytokinins in Plants. SPB Academic Publishing, The Hague, pp. 53–59. Walz, A., Park, S., Slovin, J.P., Ludwig-Müller, J., Momonoki, Y.S., Cohen, J.D., 2002. A gene encoding a protein modified by the phytohormones indoleacetic acid. Proc. Natl. Acad. Sci. USA 99, 1718–1723. Wang, Z., Xu, Q., Huang, B., 2004. Endogenous cytokinin levels and growth responses to extended photoperiods for creeping bentgrass under heat stress. Crop Sci. 44, 209–213. Weyers, J.D.B., Paterson, N.W., 2001. Plant hormones and the control of physiological processes. New Phytol. 152, 375–407. Woodward, A.W., Bartel, B., 2005. Auxin: regulation, action and interaction. Ann. Bot. 95, 707–735.

969

Yang, Y., Xu, R., Ma, C.J., Vlot, A.C., Klessig, D.F., Pichersky, E., 2008. Inactive methyl indole-3-acetic acid ester can be hydrolyzed and activated by several esterases belonging to the AtMES esterase family of Arabidopsis. Plant Physiol. 147, 1034– 1045. Zazimalova, E., Napier, R.M., 2003. Points of regulation for auxin action. Plant Cell Rep. 21, 625–634.

Andrzej Bajguz, born in 1970 (Bialystok, Poland). In 1994, he obtained MSc degree in Biology at Warsaw University, Branch in Bialystok, PhD in 1998 at the Nicholas Copernicus University in Torun´. He works at the University of Bialystok. Author or co-author of several original and review papers, chapters, and books. Member of Polish Biochemical Society, Polish Botanical Society, Polish Society of Experimental Plant Biology and Federation of European Societies of Plant Physiology. Field of scientific interest: phytohormones (auxins, cytokinins and brassinosteroids) and plant growth regulators (ecdysteroids), their physiology and biochemistry in lower and higher plants; environmental stresses.

Alicja Piotrowska, born in 1976 (Siemiatycze, Poland). In 2000, she obtained MSc degree in Biology, PhD in 2006 at University of Bialystok. She works at the University of Bialystok. Co-author of several original and review papers, chapter, and book. Member of Polish Botanical Society, Polish Society of Experimental Plant Biology and Federation of European Societies of Plant Physiology. Field of scientific interest: phytohormones (auxins and cytokinins), their physiology and biochemistry in plants; environmental stresses.