, Volume 244, Issue 3, pp 545–555 | Cite as

Plant ecdysteroids: plant sterols with intriguing distributions, biological effects and relations to plant hormones



Main conclusion

The present review summarises current knowledge of phytoecdysteroids’ biosynthesis, distribution within plants, biological importance and relations to plant hormones.

Plant ecdysteroids (phytoecdysteroids) are natural polyhydroxylated compounds that have a four-ringed skeleton, usually composed of either 27 carbon atoms or 28–29 carbon atoms (biosynthetically derived from cholesterol or other plant sterols, respectively). Their physiological roles in plants have not yet been confirmed and their occurrence is not universal. Nevertheless, they are present at high concentrations in various plant species, including commonly consumed vegetables, and have a broad spectrum of pharmacological and medicinal properties in mammals, including hepatoprotective and hypoglycaemic effects, and anabolic effects on skeletal muscle, without androgenic side-effects. Furthermore, phytoecdysteroids can enhance stress resistance by promoting vitality and enhancing physical performance; thus, they are considered adaptogens. This review summarises current knowledge of phytoecdysteroids’ biosynthesis, distribution within plants, biological importance and relations to plant hormones.


Phytoecdysteroids Ecdysteroids 20-Hydroxyecdysone Plant hormones Signalling molecules 



2,4-Dichlorophenoxy acetic acid




Abscisic acid










Indole-3-acetic acid


Jasmonic acid




Methyl jasmonate






Ecdysteroids (ECs) are steroidal hormones that have been originally found in the animal kingdom (Butenandt and Karlson 1954) to control moulting (ecdysis) and other metamorphotic processes in insects (Dinan 2001). Structurally, they are polyhydroxylated compounds that have a four-ringed skeleton, usually composed of either 27 carbon atoms or 28–29 carbon atoms (biosynthetically derived from cholesterol or other plant sterols, respectively). Since their discovery about 300 compounds of ecdysteroid character have also been detected in over 100 terrestrial plant families (annual and perennial), 20-hydroxyecdysone (20E, Fig. 1) being the most widely distributed (Lafont et al. 2002). This group of naturally occurring plant ecdysteroids was designated phytoecdysteroids (PEs) to differentiate them from those isolated from animal sources. However, this is not a strict division since some ECs are present in both animals and plants (e.g., ecdysone, 20E and ajugasterone C—Fig. 1). In addition to higher plants (both angiosperms and gymnosperms), PEs have also been discovered in ferns, fungi and algae (Dinan et al. 2009). There is evidence that most plant species have the genetic capacity to produce PEs, although they have only been detected in a small proportion (5–6 %) of 250,000 species tested (Imai et al. 1969). The inability to detect these compounds in some species could be due to suppression of gene transcription (Dinan 2009; Lafont et al. 2002), insufficient purification during sample preparation and/or instrumental limitations. Latter mentioned argument is supported by the results obtained by Dinan et al. (2001) who found in Arabidopsis, considered as PEs-negative plant, certain amount of PEs when using radioimmunoassay (RIA) as sensitive analytical detection method. Thus, the picture about PEs presence in plant species may change as technical challenges are solved and applied in the course of time. Recent studies also indicate that the occurrence of PEs may be related to phylogenetic position (Dinan 1998; Savchenko et al. 1998; Zibareva 2000). For instance, in the genus Chenopodium, PEs are present in most members of the subgenus Chenopodium, but no PEs have been detected in the subgenus Ambrosia (Dinan 1998). This review addresses PEs’ distribution in plants, their biosynthesis, biological importance and relation to known plant signalling molecules.
Fig. 1

Chemical structure of 20-hydroxyecdysone—the most widely distributed representative of ecdysteroids in nature

Distribution of phytoecdysteroids within plants

Phytoecdysteroids are frequently detectable in newly emerged tissues (young leaves) and reproductive organs (flowers, anthers, seeds) of annual plants, but less frequently in stems and roots (Dinan et al. 2001). This may imply that the highest concentrations of PEs are found in tissues that are important for plants’ survival or organs needed for the next generation of plants (Dinan et al. 2009). This is in accordance with the hypothesis that PEs may serve as protectors against predators (discussed in more detail below in the section “Biological importance of phytoecdysteroids”). Results consistent with this hypothesis have been obtained for Chenopodium album, where levels of PEs are reportedly highest in the anthers (protecting developing pollen), seeds and young leaves (Dinan 1992). In addition, cotyledons and the first two true leaves of spinach (another annual plant with rich PEs content) can reportedly synthesise PEs more rapidly and abundantly than later leaves (Grebenok and Adler 1993).

Comparing distribution of PEs in annual and perennial plants, it is believed that annual plants transfer PEs present in seeds to the developing shoots at the beginning of their growth (Grebenok et al. 1991), while many perennials (e.g. the Siberian herb Maral root Leuzea carthamoides) cycle PEs between their underground (perennial) and aerial part (deciduous) during the season, i.e. in spring the highest concentration of PEs can be found in the newly developing shoot while in autumn, the PEs are transferred from the shoot to the root where their level rises (Kholodova et al. 1979).

Biosynthesis of phytoecdysteroids

Insects and arthropods cannot synthesise sterols as they lack the ability to cyclise squalene (Fig. 2) to form them and thus must acquire those they need (for building cell membranes and as EC precursors) from dietary sources (Nes and McKean 1977). For ECs formation, C27 sterols are required, so several biosynthetic pathways must have evolved in insects and arthropods that allow them to metabolise C28 and C29 plant sterols. In contrast, plants possess a complete sterol biosynthetic pathway, thus many are capable of biosynthesising PEs (although our knowledge of PEs’ biosynthesis and localisation of the biosynthetic enzymes in plant cells is far from complete). In addition, lathosterol (Fig. 2) has been identified as a C27 phytosterol precursor for PEs in the model plant species spinach (Adler and Grebenok 1995), while in many other plants and in the model insect Drosophila, ECs are synthesised from the C27 sterol cholesterol (Fig. 2; Niwa and Niwa 2014). Originally, the plant sterol (triterpenoid) biosynthetic pathway has been described as a cytosolic pathway starting from mevalonic acid (MVA; Fig. 2) as a precursor (Goldstein and Brown 1990). However, it was recently proved that the isoprenoid building blocks (isopentenyl diphosphate and dimethylallyl diphosphate; IPP and DMAPP, respectively; Fig. 2) required for terpene synthesis can be formed not only by the MVA pathway in cytosol but also by the 2C-methyl-erythritol-4-phosphate (MEP; Fig. 2) pathway operating in plastids (Lichtenthaler 1999; Rohmer 1999). For example, for many years it was thought that isoprene units needed for formation of geranylgeranyl diphosphate, a precursor of the diterpenoid plant hormones gibberellins (GAs), are formed via the MVA pathway (MacMillan 1998). However, recent research with isotope-labelled precursors has shown that the MEP pathway in the plastid provides most of the isoprene units used to form GAs in Arabidopsis seedlings, and the contribution from the cytosolic MVA pathway is minor (Kasahara et al. 2002). Moreover, IPP serves as the basic building unit for the biosynthesis of all terpenoids, including C30 triterpenoid sterols (Piironen et al. 2000). The IPP used in their synthesis has been shown to derive exclusively from the MVA pathway. However, in germinating corn seedlings MVA may not be an intermediate in the synthesis of phytosterols and other pathways can reportedly direct carbon fluxes into IPP (Guo et al. 1995). Thus, this observation raises doubts about the pivotal role of MVA in the plant sterol pathway. However, the only studies to date on biosynthesis of PEs have been based on labelling experiments with 14C-mevalonic acid (Grebenok and Adler 1993; Bakrim et al. 2008). Thus, knowledge of PE biosynthesis is limited to parts mediated via the cytosolic sterol synthesis pathway.
Fig. 2

Simplified biosynthetic pathway of phytoecdysteroids (PEs) with relation to biosynthesis of other plant sterols including brassinosteroids (BRs)

In spite of many biosynthetic studies aiming to reveal biochemical background of ECs formation and published during last three decades, our understanding of the biosynthetic pathway(s) for these compounds still remains limited. This is especially true for PEs biosynthesis since these studies have been focused more to the ECs biosynthesis in invertebrates while only small attention has been paid to PEs synthesis in plants. To our knowledge, the sites of PEs production have not been found yet and it is not even known if their biosynthesis takes place only in specialised cells or in all cells of particular tissue/organism. Only biosynthetic enzyme purified so far is ecdysone 20-monooxygenase catalysing the oxidation of E to 20E (Fig. 2; Grebenok et al. 1996; Canals et al. 2005). This compound is Δ7 sterol possessing a reduced side chain at C-24 (Fig. 1). From this point of view, its precursor for its biosynthesis should be also C-24 reduced Δ7 sterol. This hypothesis has been confirmed using spinach as a model plant where lathosterol has been found as Δ7 sterol reduced at C-24 as already mentioned above (Grebenok and Adler 1993). Δ7 sterols are also found in related chenopods (Salt and Adler 1985) and in some other plant families (Nes 1977). Nevertheless, many other plants have been reported to possess C-24 alkylated PEs (Horn and Bergamasco 1985) being presumably formed from C-24 alkylated sterols. These plants produce 24-alkyl Δ5 sterols as well as cholesterol that is not alkylated at C-24 (Fig. 2). It is not, however, known whether cholesterol or lathosterol is the preferred substrate for PEs biosynthesis. Both these sterols have been reported to be present for instance in Chenopodium album and Chenopodium quinoa (Xu et al. 1990).

Biological importance of phytoecdysteroids

Unlike the very well-known hormonal activity of ECs in animals, there are numerous indications that plant ecdysteroids do not possess hormonal activity in planta. Notably, they have at most slight activity in various plant hormone activity assays (for instance, auxin, cytokinin, brassinosteroid and gibberellin bioassays) (Hendrix and Jones 1972; Dreier and Towers 1988; Macháčková et al. 1995). These results are consistent with apparently not universal PEs’ presence in terrestrial plant species tested to date. However, this only accounts for less than 2 % of the global flora (Dinan 2001) and, as mentioned above, methods routinely used for their detection may be insufficiently sensitive. Furthermore, PE levels in PE-positive plants are often much higher than the trace levels of phytohormones, e.g., 1–2 % dry weight in Leuzea carthamoides (Koudela et al. 1995), and no PE receptor has been reported to date. The only known ecdysteroid-binding proteins are members of the nuclear receptor superfamily with a characteristic domain structure in arthropods (Laudet 1997). For all these reasons, PEs are usually regarded as plant secondary metabolites, and not plant hormones. Despite their non-hormonal properties in plants, PEs seem to participate in the regulation of some physiological processes in photosynthesizing organisms. For example, they appear to influence the size and growth of cells of the alga Chlorella vulgaris (Bajguz and Dinan 2004) and growth of the cyanobacterium Nostoc (Maršálek et al. 1992). The main hypothesis regarding the function of PEs in plants is that they provide protection against non-adapted insects (invertebrates) and/or soil nematodes (Bergamasco and Horn 1983; Kubo and Hanke 1986). Thus, increasing plants’ endogenous PEs levels, or modulating their PEs profiles, by breeding or genetic modification, could provide potent strategies for protecting some crop plants. PEs acting either alone or in concert with other signalling molecules may deter consumption of plants (antifeedants), or lead to endocrine disruption and/or death of phytophagous invertebrates. This hypothesis is supported by results of several experiments involving either PEs exogenous applications or transgenic/mutant plants with elevated endogenous PEs levels. For instance, Udalova et al. (2004) found that spraying tomato plants with a solution of αα-ecdysone reduced infestation by the root-knot nematode Meloidogyne incognita. Similarly, Soriano et al. (2004) found that exposing cereal cyst nematodes to exogenously applied 20E at concentrations higher than 10−6 M greatly reduced their capacity to invade wheat roots. Furthermore, Schmelz et al. (1999) observed that spinach plants inoculated with two types of cereal cyst nematodes and one type of root-knot nematode were less damaged when levels of endogenous PEs in the plants were elevated by treatment with methyl jasmonate (MeJA). Generally speaking, the production of endocrine disruptors is a common defence mechanism of many plants against plant-eating insects (Bergamasco and Horn 1983). The principle of PEs action as disruptors consists in affecting the growth, development or reproduction of insects. Apparently, PEs do not show any toxicity to mammals (Dinan 2001). The susceptibility of insects to dietary PEs varies depending upon the insect species and the structure of PE (Kubo and Klocke 1983; Horn and Bergamasco 1985; Harborne 1988). For instance, 20E can cause 50 % feeding inhibition for spring wheat aphid (Schizaphis graminum; it can cause yellowing and premature death of wheat leaves) at 650 ppm in the diet, but has no effect on feeding of corn earworm (Heliothis zea; it is polyphagous major agricultural pest resistant to many pesticides and causing corn and other crop plants damage) even at higher concentration (3000 ppm) in the diet (Kubo and Klocke 1983). Further, the experiments with pink bollworm Pectinophora gossypiella revealed that 20E is capable to inhibit 95 % ecdysis of this organism at 50 ppm while 20E analogue called ponasterone A (ponA; missing hydroxyl group at position C-25—Fig. 1) is already effective at concentration of 2 ppm (Kubo and Klocke 1983).

Some PEs produced by plants also appear to act as allelochemicals, i.e., chemicals that are released by plants into the soil and stimulate or impair the growth and/or development of neighbouring organisms (plants, insects, fungi or other microbes). For example, PEs including 20E and its acetonides isolated from leaves of Chenopodium album reportedly influence seed germination and seedling growth of both monocotyledonous (onion) and dicotyledonous (lettuce and tomato) plants (DellaGreca et al. 2005; Bakrim et al. 2007). Furthermore, experiments with PE extracts containing 20E from the monocotyledon Asparagus dumosus (used in traditional Asian medicine, mainly as a diuretic and antiseptic agent) indicate that 20E appears to have antibacterial and antifungal properties (Ahmad et al. 1996). In addition, seeds of quinoa (Chenopodium quinoa Willd., Amaranthaceae), a functional food and nutraceutical, have very high contents of PEs (mainly 20E), 4–12 times more than spinach (Kumpun et al. 2011), and compounds secreted by intact quinoa seeds into water during initial stages of germination reportedly have anti-diabetic properties, as they can significantly lower fasting blood glucose in obese, hyperglycemic mice (Graf et al. 2014). Other therapeutic properties in mammals—including anabolic, performance-enhancing, anti-osteoporotic and wound healing effects—have also been described by various authors (e.g., Slama and Lafont 1995; Kapur et al. 2010; Seidlova-Wuttke et al. 2010; Syrov and Khushbaktova 1996; Lafont and Dinan 2003).

Last, but not least, there are indications that PEs participate in regulation of photosynthesis in plants. Holá et al. (2013) found that exogenous application of 20E to leaves of Tetragonia tetragonioides (New Zealand spinach) enhances their net photosynthetic rate (P N), but not their photosynthetic electron transport rate or content of photosynthetic pigments. The increase in P N was statistically significant during the 4–6 h after treatment, but not later. Moreover, the first phase of the Calvin cycle (light-independent reactions of photosynthesis)—fixation of CO2 into organic matter—is also positively affected by exogenous application of some PEs (Macek et al. 2008; Holá et al. 2012). During the first Calvin cycle reaction, the enzyme RuBisCO (ribulose-1,5-bisphosphate carboxylase/oxygenase) catalyses carboxylation of ribulose-1,5-bisphosphate, RuBP, by CO2, and exogenously applied 20E can increase the RuBP yield (most strongly with equimolar concentrations of 20E and RuBisCO under in vitro conditions).

Relations of phytoecdysteroids and plant hormones

Nine groups of plant hormones have been identified and studied to date: auxins, cytokinins, gibberellins, abscisic acid, jasmonates, ethylene, brassinosteroids and (the most recently found) strigolactones—signalling molecules expanding the family of isoprenoid plant hormones (Tarkowská et al. 2014). Plants’ intricate signalling networks are further complicated by links and interactions with synthesis and metabolic pathways of secondary metabolites. To date, no review concerning the relations of phytoecdysteroids and plant hormones has been published, but there are some indications that these substances might interact to some extent and thus influence some physiological processes and developmental events in plants. Thus, some information regarding these interactions is summarised here.


Jasmonates—jasmonic acid (JA) and its derivatives—are well-known chemical messengers that play major roles in plant responses to both biotic and abiotic stresses (Wasternack and Hause 2013). JA usually accumulates rapidly in wounded plants following mechanical damage (Glauser et al. 2008; Glauser et al. 2009) or after attacks of plant by herbivorous or phytophagous insects (Howe and Jander 2008). In addition, the volatile methyl ester of JA (MeJA) is usually generated in target plant tissue and plays a key role in rapid transmission of JA signalling (Farmer and Ryan 1990). JA-mediated processes, such as synthesis of defence proteins and biosynthesis of secondary compounds, can be induced by application of exogenous MeJA, which is converted within plants to the biologically active JA-Ile (JA conjugated with the amino acid isoleucine). These responses have been exploited to assess roles and effects of PE induction in plants’ insect protection systems (see above), initially when Schmelz et al. (1998) investigated the phytochemicals that were rapidly induced by damage treatment and applications of MeJA, using hydroponically grown spinach as a model plant. Addition of MeJA to the plants’ root systems stimulated accumulation of root PEs (20E) in a dose-dependent manner, while mechanically damaged roots exhibited two to three fold higher 20E concentrations within 2 days. Forty-eight hours later, small increases of 20E levels in shoots were also detectable, but its concentrations remained unchanged in shoots following shoot herbivory by an insect (Spodoptera exigua). Thus, the cited authors concluded that signals mediating wound-induced accumulation of PEs are transmitted via endogenous JA generated in the roots. Based on these findings and using the same inducible system (induction of 20E production by spinach plants following root damage or MeJA application), the same research group showed that attacks by the dark-winged fungus gnat (Bradysia impatiens) raise 20E levels in spinach roots four- to nearly sevenfold (Schmelz et al. 2002). Moreover, induction of 20E production in roots with MeJA resulted in a twofold increase in 20E levels and 50 % reduction in B. impatiens larval establishment. The cited authors therefore concluded that PEs can act as inducible defences against insect herbivory. This finding was subsequently validated by findings that plant-parasitic nematodes exposed to either exogenous 20E or elevated endogenous levels in plants show abnormal moulting, immobility, reduced invasiveness and impaired development leading to their death (Soriano et al. 2004).


Phytoecdysteroids share some structural resemblance with brassinosteroids (BRs), which show some biological activity in insects as weak ecdysteroid antagonists (Dinan and Hormann 2005). Chemically, both of these triterpenoid families comprise C27 to C29 polyhydroxylated steroids with an oxygenated B-ring. However, the B-ring in BRs often bears a carbonyl group at C-6 and may be expanded to form a lactone (in brassinolide and its analogues), while the ECs have a characteristic 14α-hydroxy-7-en-6-one grouping. Further, hydroxyl groups at C-2, C-3 and C-22 are found in both families, but their orientations and locations of further hydroxyls differ between them. In addition, the junction of the A- and B-rings is in cis-orientation in the skeleton of ECs while BRs have an A/B-trans-configuration. Due to these structural differences BR receptors are unlikely to recognise ECs and vice versa, so plant BR and insect EC receptors show high specificity, explaining why PEs do not interfere with BR signalling pathways in plants where these two families of compounds co-exist. It has been proved that ECs show very weak or no activity in BR-responsive plant bioassays (Dreier and Towers 1988; Macháčková et al. 1995). Similarly, BRs do not interfere with EC signalling in insects (which could otherwise have major effects, as BRs are present, albeit at very low concentrations, in plant tissues consumed by insect herbivores). However, some synthetic structural analogues of both ECs and BRs (castasterone with various modifications) have been prepared to test their potential agonist or antagonist activity (Voigt et al. 2001) in the Drosophila melanogaster BII cell bioassay (for EC activity) and rice lamina inclination bioassay (for BR activity). Only one tested compound showed distinct PE agonist activity, at relatively high concentration (1 × 10−5 M), almost 2000-fold weaker activity than 20E. In the BR bioassay, most of the tested substances showed activity, with indications that biological activity declines with increasing structural deviation of the test compound from castasterone (used as a natural BR standard).

As mentioned above in the “Biological importance of phytoecdysteroids” section, some PEs (notably 20E) can affect various processes associated with photosynthesis, and BRs (at least 24-epibrassinolide, epiBL) share this property. For example, in pea plants, exogenous epiBL induces alterations in the thermodynamic parameters of photosynthetic membranes, reorganisation of the main pigment–protein complexes and partial unstacking of thylakoid membranes (Dobrikova et al. 2014). The cited authors claim that BR-induced changes in photosynthetic membranes are probably involved in the stress tolerance of plants. epiBL can also increase photosynthesis yields at even lower concentrations than the PE concentrations required to affect RuBisCO activity (Rothová et al. 2014). Moreover, exogenously applied epiBL induces changes in PEs contents in plant tissues (Kamlar et al. 2015), to degrees that depend on the developmental stage of leaves (of the same plant) and the epiBL concentration applied (10−8 or 10−6 M in the cited experiments). In control plants, young leaves had ca. sevenfold higher contents of the major PEs than the older leaves. Treatment with epiBL led to a reduction in endogenous 20E content in the young leaves at 10−6 M concentration, but an increase at 10−8 M. These effects were weaker, only temporary and apparent within 4 h of treatment in older leaves. Exogenous epiBL also changed the leaves’ PE profiles. Young leaves of control plants had ca. tenfold higher polypodine B (polB, Fig. 1) contents but twofold lower ajugasterone C (ajuC; Fig. 1) and stachysterone C (stachC; Fig. 1) levels than older leaves (and similar levels of the trace compound ponA). Application of epiBL at 10−6 M led to reductions in stachC and ponA levels, but increases in ajuC abundance (which were stronger when epiBL was applied at 10−8 M). These observations confirm that exogenous BRs can induce detectable changes in levels of individual endogenous PEs in the same tissue within hours.


Cytokinins (CKs) comprise one of the most important classes of endogenous growth- and development-regulating substances in plants. Chemically, they are adenine species substituted at the N6-position with either an isoprenoid or aromatic side-chain. They occur in plant tissues as free species, ribosides, ribotides or glycosides, and have no structural similarity with PEs. Although they are mainly known to have pronounced effects on plant development, at least one CK (isopentenyladenosine, iPR), is also present in animal cells (Faust and Dice 1991). Furthermore, iPR has anti-proliferative activity against mammalian cancer cell lines (Rajabi et al. 2010; Casati et al. 2011). This is not entirely surprising since callus consists of clusters of dedifferentiated plant cells that proliferate indefinitely in a disorganised manner, like human cancer cells, and a signature effect of iPR (and other CKs) is induction of re-differentiation of callus to form adventitious buds (Tanimoto and Harada 1982). Moreover, in insects (traditionally rich sources of ECs) an iPR monophosphate conjugate with ecdysone has been detected. Tsoupras et al. (1983) identified this compound as 22-N 6-(isopentenyl)adenosine monophosphoric ester of ecdysone by mass spectrometry and NMR, and showed that it was present at very high levels (50 nmol/g) in newly laid eggs of the migratory locust (Locusta migratoria). They also found that the eggs contained even higher (more than double) levels of a 2-deoxyecdysone conjugate with adenosine monophosphate (Tsoupras et al. 1982a). Later investigation revealed that this insect can produce adenosine 22-phosphate conjugates of 2-deoxy-20-hydroxyecdysone, 20-hydroxyecdysone and 20-hydroxyecdysone acetate (Tsoupras et al. 1982b). All these conjugates produced by female locusts are further progressively hydrolysed during embryonic development, yielding free highly biologically active hormones (Lagueux et al. 1979). Therefore, it is believed that females of migratory locust and other related insect species (Isaac et al. 1982) serve as suppliers of hormonal molecules for their offspring, which the embryos cannot synthesise de novo before they reach advanced developmental stages. Another indication that CKs may influence the development of insects is that the aromatic CK 6-benzylaminopurine (BAP) can stimulate growth of Drosophila cells (Becker and Roussaux 1981). In addition, dietary provision of another aromatic CK, kinetin (6-furfurylaminopurine, known to delay senescence in plants), can retard ageing of Zaprionus paravittiger fruit flies and prolong their lifespan (Sharma et al. 1995). This anti-ageing effect was attributed to a reduction in age-specific death rates throughout the adult lifespan rather than slowing of development. This study also describes retardation of the larval and pupal stages of the developing insects by kinetin. Overall, the findings summarised in this section indicate that interactions between CKs and ECs warrant further attention.


Three major PEs (20E-3-acetate, 20E and its hydroxylated analogue polypodine B) have been found in tissues of the perennial plant Serratula tinctoria and in vitro cultures (cell suspension cultures and calluses) of this plant (Corio-Costet et al. 1993). Supplementation of cultures of this species transformed by Agrobacterium rhizogenes (with a characteristic hairy root phenotype) with the synthetic auxin 2,4-dichlorophenoxy acetic acid (2,4-D) had dose-dependent negative effects on their PE contents. Growth of the hairy roots slowed on media containing 0.05–0.5 mg/l of 2,4-D, and addition of 1 mg/l stopped their growth entirely due to tissue necrosis (Corio-Costet et al. 1996). Reductions in EC concentrations in hairy roots of another plant species (the European herbaceous plant Ajuga reptans) have also been observed following addition of 0.1 mg/l of natural auxin (indole-3-acetic acid, IAA) to the cultivation medium, despite increases in the roots’ growth rate due to increases in the number of root apical meristems (Uozumi et al. 1995). To our knowledge, no published study has addressed effects of exogenously applied ECs on levels of endogenous auxins in planta. Tests of PEs’ auxin-like activity in a wheat coleoptile assay revealed that ecdysone has no biological activity, except slight inhibitory effects at non-physiologically high concentrations (Macháčková et al. 1995). Thus, no notable synergistic or antagonistic relationship between PEs and auxin has been found to date.

Phytoecdysteroids and other plant hormones

We are aware of just one study concerning relations between gibberellins (GAs) and PEs. In this study, Macháčková et al. (1995) found that two insect moulting hormones (20E and E) had no effects in a dwarf maize GA bioassay, but slight gibberellin-like activity in dwarf rice bioassays (0.4- to 4-fold weaker effects than gibberellic acid). Nevertheless, its action was unequivocally and dose-dependently stimulatory in the latter assay, and the results also indicate a slight synergistic action of ecdysone and gibberellic acid.

Infinitesimal stimulation of ethylene production in Chenopodium rubrum and dwarf maize by ecdysone has also been observed in bioassays within 3–5 h of its application. However, this effect subsequently disappeared leading to the conclusion that ecdysone has negligible effects on ethylene production in the test system used. To our knowledge, nothing is known about possible relations between PEs and other plant hormones like abscisic acid and strigolactones.


PEs clearly have intriguing distributions, but their roles in plants are still obscure despite many recent findings. Their likeliest functions still seem to be in defences against insect herbivores, which have potentially profound implications for (and applications in) agriculture. Furthermore, they clearly have potent pharmacological and medicinal properties that warrant much more attention (particularly as plants comprise a major part of human diets). For example, the numerous indications that they are very effective, non-androgenic, adaptogens and elicitors of anabolic effects on skeletal muscle suggest possible uses as natural drugs to boost people’s stress resistance, post-damage muscle regeneration and sports performance. In addition, PE contents of many more plants should be screened, using sensitive modern analytical instruments to allow detection of PEs present at low concentrations to broaden knowledge of their distributions, profiles and roles in both perennial and annual plants. Last, but not least, further analysis of the origin(s) of isoprenoid units used in PEs’ biosynthesis is required.

Author contribution statement

DT designed the outline of the article, composed the manuscript and figures. MS provided scientific feedback and critical comments and revised content. Both authors read and approved the manuscript.



Financial support from the Ministry of Education, Youth and Sport of the Czech Republic through the National Program of Sustainability (Grant No. LO 1204) is gratefully acknowledged. The authors would like to also express thanks to Sees-editing Ltd., Prof. Claus Wasternack and Dr. Juraj Harmatha for their critical reading and editing of the manuscript.


  1. Adler JH, Grebenok RJ (1995) Biosynthesis and distribution of insect-molting hormones in plants—a review. Lipids 30:257–262. doi: 10.1007/BF02537830 CrossRefPubMedGoogle Scholar
  2. Ahmad VU, Khaliq-Uz-Zaman SM, Ali MS, Perveen S, Ahmed W (1996) An antimicrobial ecdysone from Asparagus dumosus. Fitoterapia 67:88–91Google Scholar
  3. Bajguz A, Dinan L (2004) Effects of ecdysteroids on Chlorella vulgaris. Physiol Plant 121:349–357. doi: 10.1111/j.1399-3054.2004.00329.x CrossRefGoogle Scholar
  4. Bakrim A, Lamhamdi M, Sayah F, Chibi F (2007) Effects of plant hormones and 20-hydroxyecdysone on tomato (Lycopersicum esculentum) seed germination and seedlings growth. Afr J Biotechnol 6:2792–2802CrossRefGoogle Scholar
  5. Bakrim A, Maria A, Sayah F, Lafont R, Takvorian N (2008) Ecdysteroids in spinach (Spinacia oleracea L.): biosynthesis, transport and regulation of levels. Plant Physiol Biochem 46:844–854. doi: 10.1016/j.plaphy.2008.06.002 CrossRefPubMedGoogle Scholar
  6. Becker JL, Roussaux J (1981) 6-Benzylaminopurine as a growth factor for Drosophila melanogaster cells grown in vitro. In: Guern J, Peaud-Lenoël C (eds) Metabolism and molecular activities of cytokinins. Springer, Berlin, pp 319–328CrossRefGoogle Scholar
  7. Bergamasco R, Horn DHS (1983) Distribution and role of insect hormones in plants. Endocrinology of insects. A. R. Liss Inc., New York, pp 627–654Google Scholar
  8. Butenandt A, Karlson P (1954) Über die Isolierung eines Metamorphose-hormons der Insekten in kristallisierter Form. Z Naturforsch 9B:389–391Google Scholar
  9. Canals D, Irurre-Santilari J, Casas J (2005) The first cytochrome P450 in ferns. FEBS J 272:4817–4825. doi: 10.1111/j.1742-4658.2005.04897.x CrossRefPubMedGoogle Scholar
  10. Casati S, Ottria R, Baldoli E, Lopez E, Maier JAM, Ciuffreda P (2011) Effects of cytokinins, cytokinin ribosides and their analogs on the viability of normal and neoplastic human cells. Anticancer Res 31:3401–3406PubMedGoogle Scholar
  11. Corio-Costet MF, Chapuis C, Moulilett JF, Delbeckque JP (1993) Sterol and ecdysteroid profiles of Serratula tinctoria (L.): plant and cell cultures producing steroids. Insect Biochem Mol Biol 23:175–180. doi: 10.1016/0965-1748(93)90098-D CrossRefGoogle Scholar
  12. Corio-Costet MF, Chapuis L, Delbecque JP (1996) Serratula tinctoria (Dyer’s savory): in vitro culture and the production of ecdysteroids and other secondary metabolites. In: Bajaj YPS (ed) Biotechnology in agricultural and forestry. Trees IV, Medicinal and Aromatic Plants, vol 37. Springer, Berlin, pp 384–401. doi: 10.1007/978-3-662-08618-6_23
  13. DellaGreca M, D’Abrosca B, Fiorentino A, Previtera L, Zarrelli A (2005) Structure elucidation and phytotoxicity of ecdysteroids from Chenopodium album. Chem Biodivers 2:457–462. doi: 10.1002/cbdv.200590025 CrossRefPubMedGoogle Scholar
  14. Dinan L (1992) The analysis of phytoecdysteroids in single (preflowering stage) specimens of fat hen, Chenopodium album. Phytochem Anal 3:132–138. doi: 10.1002/pca.2800030309 CrossRefGoogle Scholar
  15. Dinan L (1998) A strategy towards the elucidation of the contribution made by phytoecdysteroids to the deterrence of invertebrate predators on plants. Russ J Plant Physiol 45:296–305Google Scholar
  16. Dinan L (2001) Phytoecdysteroids: biological aspects. Phytochemistry 57:325–339. doi: 10.1016/S0031-9422(01)00078-4 CrossRefPubMedGoogle Scholar
  17. Dinan L (2009) The Karlson lecture. Phytoecdysteroids: what use are they? Arch Insect Biochem Physiol 72:126–141. doi: 10.1002/arch.20334 CrossRefPubMedGoogle Scholar
  18. Dinan L, Hormann R (2005) Ecdysteroid agonists and antagonists. In: Gilbert LI, Iatrou K, Gill S (eds) Comprehensive molecular insect science, vol 3. Elsevier, Amsterdam, pp 197–242CrossRefGoogle Scholar
  19. Dinan L, Savcenko T, Whiting P (2001) On the distribution of phytoecdysteroids in plants. Cell Mol Life Sci 58:121–1132. doi: 10.1007/PL00000926 CrossRefGoogle Scholar
  20. Dinan L, Harmatha J, Volodin V, Lafont R (2009) Phytoecdysteroids: diversity, biosynthesis and distribution. In: Smagghe G (ed) Ecdysone: structures and functions. Springer, Berlin, pp 3–45. doi: 10.1007/978-1-4020-9112-4_1
  21. Dobrikova AG, Vladkova RS, Rashkov GD, Todinova SJ, Krumova SB, Apostolova EL (2014) Effects of exogenous 24-epibrassinolide on the photosynthetic membranes under non-stress conditions. Plant Physiol Biochem 80:75–82. doi: 10.1016/j.plaphy.2014.03.022 CrossRefPubMedGoogle Scholar
  22. Dreier SI, Towers GHN (1988) Activity of ecdysterone in selected plant growth bioassays. J Plant Physiol 132:509–512. doi: 10.1016/S0176-1617(88)80073-7 CrossRefGoogle Scholar
  23. Farmer EE, Ryan CA (1990) Interplant communication: airborne methyl jasmonate induces synthesis of proteinase inhibitors in plant leaves. Proc Natl Acad Sci USA 87:7713–7716. doi: 10.1073/pnas.87.19.7713 CrossRefPubMedPubMedCentralGoogle Scholar
  24. Faust JR, Dice JF (1991) Evidence for isopentenyladenine modification on a cell cycle-regulated protein. J Biol Chem 266:9961–9970PubMedGoogle Scholar
  25. Glauser G, Grata E, Dubugnon L, Rudaz S, Farmer EE, Wolfender J-L (2008) Spatial and temporal dynamics of jasmonate synthesis and accumulation in Arabidopsis in response to wounding. J Biol Chem 283:16400–16407. doi: 10.1074/jbc.M801760200 CrossRefPubMedGoogle Scholar
  26. Glauser G, Dubugnon L, Mousavi SAR, Rudaz S, Wolfender J-L, Farmer EE (2009) Velocity estimates for signal propagation leading to systemic jasmonic acid accumulation in wounded Arabidopsis. J Biol Chem 284:34506–34513. doi: 10.1074/jbc.M109.061432 CrossRefPubMedPubMedCentralGoogle Scholar
  27. Goldstein JL, Brown MS (1990) Regulation of the mevalonate pathway. Nature 343:425–430. doi: 10.1038/343425a0 CrossRefPubMedGoogle Scholar
  28. Graf BL, Poulev A, Kuhn P, Grace MH, Lila MA, Raskin I (2014) Quinoa seeds leach phytoecdysteroids and other compounds with anti-diabetic properties. Food Chem 163:178–185. doi: 10.1016/j.foodchem.2014.04.088 CrossRefPubMedPubMedCentralGoogle Scholar
  29. Grebenok RJ, Adler JH (1993) Ecdysteroid biosynthesis during the ontogeny of spinach leaves. Phytochemistry 33:341–347. doi: 10.1016/0031-9422(93)85514-R CrossRefGoogle Scholar
  30. Grebenok RJ, Ripa PV, Adler JH (1991) Occurrence and levels of ecdysteroids in spinach. Lipids 26:666–668. doi: 10.1007/BF02536433 CrossRefGoogle Scholar
  31. Grebenok RJ, Galbraith DW, Benveniste I, Feyereisen R (1996) Ecdysone 20-monooxygenase, a cytochrome P450 enzyme from spinach, Spinacia oleracea. Phytochemistry 420:927–933. doi: 10.1016/0031-9422(96)00094-5 CrossRefGoogle Scholar
  32. Guo DA, Vekatramesh M, Nes WD (1995) Developmental regulation of sterol biosynthesis in Zea mays. Lipids 30:203–219. doi: 10.1007/BF02537823 CrossRefPubMedGoogle Scholar
  33. Harborne JB (1988) In: Introduction to ecological biochemistry, 3rd edn. Academic Press, New York, pp 120–146Google Scholar
  34. Hendrix SD, Jones RL (1972) The activity of β-ecdysone in four gibberellin bioassays. Plant Physiol 50:199–200. doi: 10.1104/pp.50.1.199 CrossRefPubMedPubMedCentralGoogle Scholar
  35. Holá D, Rothova O, Kocova M, Fridrichova L, Macek T (2012) Phytoecdysteroids together with brassinosteroids stimulate oxygen-evolving activity of photosystem II. Plant Biology Congress. Book of Abstracts, Freiburg, p 771Google Scholar
  36. Holá D, Kočová M, Rothová O, Tůmová L, Kamlar M, Macek T (2013) Exogenously applied 20-hydroxyecdysone increases the net photosynthetic rate but does not affect the photosynthetic electron transport or the content of photosynthetic pigments in Tetragonia tetragonioides L. Acta Physiol Plant 35:3489–3495. doi: 10.1007/s11738-013-1379-6 CrossRefGoogle Scholar
  37. Horn DHS, Bergamasco R (1985) Chemistry of ecdysteroids. In: Kerkut GA, Gilbert LI (eds) Comprehensive insect physiology, biochemistry and pharmacology, vol 7. Pergamon Press, New York, pp 185–248Google Scholar
  38. Howe G, Jander G (2008) Plant immunity to insect herbivores. Annu Rev Plant Biol 59:41–66. doi: 10.1146/annurev.arplant.59.032607.092825 CrossRefPubMedGoogle Scholar
  39. Imai S, Toyosato T, Sakai M, Sato Y, Fujioka S, Murata E, Goto M (1969) Screening results of plants for phytoecdysones. Chem Pharm Bull 17:335–339CrossRefPubMedGoogle Scholar
  40. Isaac RE, Rose ME, Rees HH, Goodwin TW (1982) Identification of ecdysone-22-phosphate and 2-deoxyecysone-22-phosphate in eggs of the desert locust, Schistocerca gregaria, by fast atom bombardment mass spectrometry and NMR spectroscopy. J Chem Soc Chem Commun 4:249–251. doi: 10.1039/c39820000249 CrossRefGoogle Scholar
  41. Kamlar M, Rothova O, Salajkova S, Tarkowska D, Drasar P, Kocova M, Harmatha J, Hola D, Kohout L, Macek T (2015) The effect of exogenous 24-epibrassinolide on the ecdysteroid content in the leaves of Spinacia oleracea L. Steroids 97:107–112. doi: 10.1016/j.steroids.2014.12.024 CrossRefPubMedGoogle Scholar
  42. Kapur P, Wuttke W, Jarry H, Seidlova-Wuttke D (2010) Beneficial effects of beta-ecdysone on the joint, epiphyseal cartilage tissue and trabecular bone in ovariectomized rats. Phytomedicine 17:350–355. doi: 10.1016/j.phymed.2010.01.005 CrossRefPubMedGoogle Scholar
  43. Kasahara H, Hanada A, Kuzuyama T, Takagi M, Kamiya Y, Yamaguchi S (2002) Contribution of the mevalonate and methylerythritol phosphate pathways to the biosynthesis of gibberellins in Arabidopsis. J Biol Chem 277:45188–45194. doi: 10.1074/jbc.M208659200 CrossRefPubMedGoogle Scholar
  44. Kholodova YD, Baltaev U, Volovenko VO, Gorovits MB, Abubakirov NK (1979) Phytoecdisones of Serratula xeranthemoides. Khim Priv Soedin 2:171–174Google Scholar
  45. Koudela K, Tenora I, Bajer J, Maťhová A, Sláma K (1995) Stimulation of growth and development in Japanese quails after oral administration of ecdysteroid-containing diet. Eur J Entomol 92:349–354Google Scholar
  46. Kubo I, Hanke FJ (1986) Chemical methods for isolating and identifying phytochemicals biologically active in insects. In: Miller JR, Miller TA (eds) Insect plant interactions. Springer, New York, pp 225–249CrossRefGoogle Scholar
  47. Kubo I, Klocke JA (1983) Isolation of phytoecdysones as insect ecdysis inhibitors and feeding deterrents. In: Hedin EA (ed) Plant resistance to insects. American Chemical Society, Washington, DC, pp 329–346CrossRefGoogle Scholar
  48. Kumpun S, Maria A, Crouzet S, Evrard-Todeschi N, Girault J-P, Lafont R (2011) Ecdysteroids from Chenopodium quinoa Willd., an ancient Andean crop of high nutritional value. Food Chem 125:1226–1234. doi: 10.1016/j.foodchem.2010.10.039 CrossRefGoogle Scholar
  49. Lafont R, Dinan L (2003) Practical uses for ecdysteroids in mammals including humans: an update. J Insect Sci 3:7CrossRefPubMedPubMedCentralGoogle Scholar
  50. Lafont R, Harmatha J, Marion-Poll F, Dinan L (2002) Ecdybase—the ecdysone handbook, 3rd edn. Cybersales, Praha.
  51. Lagueux M, Hetru C, Goltzene F, Kappler C, Hoffmann JA (1979) Ecdysone titre and metabolism in relation to cuticulogenesis in embryos of Locusta migratoria. J Insect Physiol 25:709–723. doi: 10.1016/0022-1910(79)90123-9 CrossRefGoogle Scholar
  52. Laudet V (1997) Evolution of the nuclear receptor superfamily: early diversification from an ancestral orphan receptor. J Mol Endocrinol 19:207–226. doi: 10.1677/jme.0.0190207 CrossRefPubMedGoogle Scholar
  53. Lichtenthaler HK (1999) The 1-deoxy-d-xylulose-5-phosphate pathway of isoprenoid biosynthesis in plants. Annu Rev Plant Physiol Plant Mol Biol 50:47–65. doi: 10.1146/annurev.arplant.50.1.47 CrossRefPubMedGoogle Scholar
  54. Macek T, Uhlik O, Kamlar M, Harmatha J, Kohout L (2008) Method of increasing of the photosynthetic carbon dioxide assimilation yield (IOCB AS CR). PCT Int Appl WO 2008(125069):A2Google Scholar
  55. Macháčková I, Vágner M, Sláma K (1995) Comparison between the effects of 20-hydroxyecdysone and phytohormones on growth and development in plants. Eur J Entomol 92:309–316Google Scholar
  56. MacMillan J (1998) Gibberellin metabolism. Pure Appl Chem 50:995–1004. doi: 10.1351/pac197850090995 Google Scholar
  57. Maršálek B, Šimek M, Smith RJ (1992) The effect of ecdysone on the cyanobacterium Nostoc 6720. Z Naturforsch 47c:726–730Google Scholar
  58. Nes WR (1977) Biochemistry of plant sterols. Adv Lipid Res 15:233–324CrossRefGoogle Scholar
  59. Nes WR, McKean ML (1977) Biochemistry of steroids and other isopentenoids. University Park Press, Baltimore, pp 411–533Google Scholar
  60. Niwa R, Niwa RS (2014) Enzymes for ecdysteroid biosynthesis: their biological functions in insects and beyond. Biosci Biotechnol Biochem 78:1283–1292. doi: 10.1080/09168451.2014.942250 CrossRefPubMedGoogle Scholar
  61. Piironen V, Lindsay DG, Miettinen TA, Toivo J, Lampi A-M (2000) Plant sterols: biosynthesis, biological function and their importance to human nutrition. J Sci Food Agric 80:939–966. doi: 10.1002/(SICI)1097-0010(20000515)80:7<939:AID-JSFA644>3.3.CO;2-3 CrossRefGoogle Scholar
  62. Rajabi M, Signorelli P, Gorincioi E, Ghidoni R, Santaniello E (2010) Antiproliferative activity of N6-isopentenyladenosine on MCF-7 breast cancer cells: cell cycle analysis and DNA-binding study. DNA Cell Biol 29:687–691. doi: 10.1089/dna.2010.1073 CrossRefPubMedGoogle Scholar
  63. Rohmer M (1999) The discovery of a mevalonate-independent pathway for isoprenoid biosynthesis in bacteria, algae and higher plants. Nat Prod Rep 16:565–574. doi: 10.1039/a709175c CrossRefPubMedGoogle Scholar
  64. Rothová O, Holá D, Kočová M, Tůmová L, Hnilička F, Hniličková H, Kamlar M, Macek T (2014) 24-Epibrassinolide and 20-hydroxyecdysone affect photosynthesis differently in maize and spinach. Steroids 85:44–57. doi: 10.1016/j.steroids.2014.04.006 CrossRefPubMedGoogle Scholar
  65. Salt TA, Adler JH (1985) Diversity of sterol composition in the family Chenopodiaceae. Lipids 20:594–601. doi: 10.1007/BF02534285 CrossRefGoogle Scholar
  66. Savchenko T, Whiting P, Šik V, Underwood E, Sarker SD, Dinan L (1998) Distribution and identities of phytoecdysteroids in the genus Briza (Gramineae). Biochem Syst Ecol 26:781–791. doi: 10.1016/S0305-1978(98)00044-1 CrossRefGoogle Scholar
  67. Schmelz EA, Grebenok RJ, Galbraith DW, Bowers WS (1998) Damage-induced accumulation of phytoecdysteroids in spinach: a rapid root response involving the octadecanoic acid pathway. J Chem Ecol 24:339–360. doi: 10.1023/A:1022588610232 CrossRefGoogle Scholar
  68. Schmelz EA, Grebenok RJ, Galbraith DW, Bowers WS (1999) Insect-induced synthesis of phytoecdysteroids in spinach, Spinacia oleracea. J Chem Ecol 25:1739–1757. doi: 10.1023/A:1020969413567 CrossRefGoogle Scholar
  69. Schmelz EA, Grebenok RJ, Ohnmeiss TE, Bowers WS (2002) Interactions between Spinacia oleracea and Bradysia impatiens: a role for phytoecdysteroids. Arch Insect Biochem Physiol 51:204–221. doi: 10.1002/arch.10062 CrossRefPubMedGoogle Scholar
  70. Seidlova-Wuttke D, Christel D, Kapur P, Nguyen BT, Jarry H, Wuttke W (2010) Beta-ecdysone has bone protective but no estrogenic effects in ovariectomized rats. Phytomedicine 17:884–889. doi: 10.1016/j.phymed.2010.03.021 CrossRefPubMedGoogle Scholar
  71. Sharma SP, Kaur P, Rattan SIS (1995) Plant-growth hormone kinetin delays ageing, prolongs the life span and slows development of the fruit fly Zaprionus paravittiger. Biochem Biophys Res Commun 216:1067–1071. doi: 10.1006/bbrc.1995.2729 CrossRefPubMedGoogle Scholar
  72. Slama K, Lafont R (1995) Insect hormones—ecdysteroids: their presence and actions in vertebrates. Eur J Entomol 92:355–377Google Scholar
  73. Soriano IR, Riley IT, Potter MJ, Bowers WS (2004) Phytoecdysteroids: a novel defense against plant-parasitic nematodes. J Chem Ecol 30:651–654. doi: 10.1023/B:JOEC.0000045584.56515.11 CrossRefGoogle Scholar
  74. Syrov VN, Khushbaktova ZA (1996) Wound-healing effects of ecdysteroids. Doklady Akademii Nauk Respubliki Uzbekistana 12:47–50Google Scholar
  75. Tanimoto S, Harada H (1982) Effect of cytokinin and anticytokinin on the initial stage of adventitious bud differentiation in the epidermis of Torenia stem segments. Plant Cell Physiol 23:1371–1376Google Scholar
  76. Tarkowská D, Novák O, Floková K, Tarkowski P, Turečková V, Grúz J, Rolčík J, Strnad M (2014) Quo vadis plant hormone analysis? Planta 240:55–76. doi: 10.1007/s00425-014-2063-9 CrossRefPubMedGoogle Scholar
  77. Tsoupras G, Hetru C, Luu B, Lagueux M, Constantin E, Hoffmann JA (1982a) The major conjugates of ecdysteroids in young eggs and in embryos of Locusta-migratoria. Tetrahedron Lett 23:2045–2048. doi: 10.1016/S0040-4039(00)87256-1 CrossRefGoogle Scholar
  78. Tsoupras G, Luu B, Hoffmann JA (1982b) Isolation and identification of three ecdysteroid conjugates with a C-20 hydroxy group in eggs of Locusta migratoria. Steroids 40:551–560. doi: 10.1016/0039-128X(82)90075-7 CrossRefPubMedGoogle Scholar
  79. Tsoupras G, Luu B, Hoffmann JA (1983) A cytokinin (isopentenyl-adenosyl-mononucleotide) linked to ecdysone in newly laid eggs of Locusta migratoria. Science 220:507–509. doi: 10.1126/science.220.4596.507 CrossRefPubMedGoogle Scholar
  80. Udalova ZV, Zinov’eva SV, Vasil’eva IS, Paseshnichenko VA (2004) Correlation between the structure of plant steroids and their effects on phytoparasitic nematodes. Appl Biochem Microbiol 40:93–97. doi: 10.1023/B:ABIM.0000010362.79928.77 CrossRefGoogle Scholar
  81. Uozumi N, Makino S, Kobayashi T (1995) 20-Hydroxyecdysone production in Ajuga hairy root controlling intracellular phosphate content based on kinetic model. J Ferment Bioeng 80:362–368. doi: 10.1016/0922-338X(95)94205-6 CrossRefGoogle Scholar
  82. Voigt B, Whiting P, Dinan L (2001) The ecdysteroid agonist/antagonist and brassinosteroid-like activities of synthetic brassinosteroid/ecdysteroid hybrid molecules. Cell Mol Life Sci 58:1133–1140. doi: 10.1007/PL00000927 CrossRefPubMedGoogle Scholar
  83. 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 111:1021–1058. doi: 10.1093/aob/mct067 CrossRefPubMedPubMedCentralGoogle Scholar
  84. Xu S, Patterson GW, Lusby WR, Schmid KM, Salt TA (1990) The distribution and phylogenetic significance of desmethylsterols in Chenopodium and Atriplex: coexistence of Δ7- and Δ5-sterols. Lipids 25:61–64. doi: 10.1007/BF02562429 CrossRefGoogle Scholar
  85. Zibareva L (2000) Distribution and levels of phytoecdysteroids in plants of the genus Silene during development. Arch Insect Biochem Physiol 43:1–8. doi: 10.1002/(SICI)1520-6327(200001)43:1<1:AID-ARCH1>3.0.CO;2-D CrossRefPubMedGoogle Scholar

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© Springer-Verlag Berlin Heidelberg 2016

Authors and Affiliations

  1. 1.Laboratory of Growth Regulators, Centre of the Region Haná for Biotechnological and Agricultural Research, Institute of Experimental Botany AS CR, Faculty of SciencePalacký UniversityOlomoucCzech Republic

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