2002 Brassinosteriods

BRASSINOSTEROIDS
M I N I R E V I E143
W
Brassinosteroid phytohormones - structure, bioactivity

and applications
Marco António Teixeira Zullo1* and Günter Adam2
1 Instituto Agronômico, Centro de Recursos Genéticos Vegetais, CP 28, 13001-970 Campinas, SP, Brasil; 2 Institut für
Pflanzenbiochemie, Weinberg 3, D-06120 Halle, Alemanha. * Corresponding author: mzullo@cec.iac.br
Received: 20/09/2002, Accepted: 07/11/2002
Brassinosteroids are a new class of plant hormones with a polyoxygenated steroid structure showing pronounced plant
growth regulatory activity. This review covers their natural occurrence, biological and chromatographic methods for
their detection, biosynthesis and metabolism, biological activity, structure-activity relationships and prospective agri-
cultural uses.
Key words: brassinolide, brassinosteroids.
Hormônios vegetais brassinosteroídicos – estrutura, bioatividade e aplicações: Os brassinosteróides são uma nova
classe de hormônios vegetais com estrutura esteroídica polioxigenada, dotados de pronunciada atividade reguladora do
crescimento vegetal. Esta revisão cobre sua ocorrência natural, os métodos biológicos e cromatográficos para sua detecção,
sua biossíntese e metabolismo, a atividade biológica, as relações estrutura-atividade e suas perspectivas de uso agrícola.
Palavras-chave: brassinolídeo, brassinosteróides.
INTRODUCTION unexpected response, combining elongation (typical of

gibberellins) with swelling and curvature of the treated
Since the 1930’s, USDA researchers have found that
internode. These researchers proposed that the rape pol-
pollen extracts promote plant growth (Mandava, 1988) and
len contained a new group of lipidic plant hormones, which
a first paper was published in 1941 reporting that applica-
they called brassins (Mitchell et al., 1970). Mandava,
tion of hexanic extracts of maize pollen to the first intern-
Mitchell and co-workers reported the occurrence of an
ode of young bean seedlings produced marked elongation
active fraction of the brassins containing, mainly, glucosyl
of the treated internode (Mitchell and Whitehead, 1941), a
esters of fatty acids (Mandava and Mitchell, 1972;
response also obtained using extracts of immature bean
Mandava et al., 1973). Further work revealed that although
seeds (Mitchell et al., 1951).
these esters promoted elongation, they were not able to
In the 1960’s the USDA started a research program reproduce all of the observed response (Grove et al., 1978).
aimed to find new plant hormones, leaded by J. W. Mitchell Brassins, however, were able to increase plant growth, crop
(Maugh II, 1981). The hypothesis to be tested was that yield and seed viability (Mitchell and Gregory, 1972; Gre-
pollens would have an elevated concentration of plant hor- gory, 1981; Meudt et al., 1984).
mones and there was a great probability of finding new
In 1975 a research project to identify and synthesize
physiologically active substances in them.
the active compounds in brassins, evaluate their effect on
Employing the bean second internode bioassay, pol- the yield of selected crops (such as wheat, maize, soybean
len extracts of around 60 different plant species were tested and potato), and evaluate their growth regulating proper-
(Mandava and Mitchell, 1971): rape (Brassica napus L.) ties in green-houses and in the field was begun (Mandava,
and alder tree (Alnus glutinosa L.) pollens produced an 1988; Steffens, 1991).
Braz. J. Plant Physiol., 14(3):143-181, 2002

144 M.A.T. ZULLO AND G. ADAM
For isolating the brassins active compounds, about ure 1] have been isolated from plant sources and fully char-
250 kg of bee collected rape pollen were extracted with acterized by the usual spectroscopic methods. The vast
isopropanol in batches of 25 kg (Mandava et al., 1978). majority of the more than 50 hitherto known natural
The extracts were partitioned between carbon tetrachlo- brassinosteroids were detected in different organs of plants
ride, methanol and water. The methanolic fractions were in several families by gas or liquid chromatography com-
chromatographed in a series of silica columns, a process bined with mass spectrometry and comparison with au-
that reduced the biologically active material to 100 g. Fi- thentic samples (Adam and Marquardt, 1986; Singh and
nal purification, followed by the bean second internode Bhardwaj, 1986; Mandava, 1988; Abreu, 1991; Takatsuto,
assay, was accomplished by column chromatography and 1994; Fujioka and Sakurai, 1997a; Adam et al., 1999;
high performance liquid chromatography, affording 10 mg Fujioka, 1999).
of a crystalline substance, called brassinolide (1, figure Brassinosteroids can be derived from the 5α-
1). Its structure was elucidated by spectroscopic methods cholestane carbon skeleton bearing the following struc-
including X-ray analysis and can be systematically desig- tural characteristics:
nated as (22R,23R,24S)-2α,3α,22,23-tetrahydroxy-24-
i) ring A mono- to trioxygenated, always oxygenated
methyl-B-homo-7-oxa-5α-cholestan-6-one (Grove et al.,
at carbon 3;
1979; see the Appendix for notation of steroids).
ii) ring B presenting a 6-oxo-7-oxalactone or a 6-oxo
At the time brassinolide (1) was pointed out to be the
function or full saturation;
first phytosteroid with plant growth-promoting activity,
iii) all-trans junctions of rings A - D;
even in very minute amounts and concentrations, present-
ing a 6-oxo-7-oxalactone function and two groups of cis- iv) 22α,23α-dihydroxylated, mostly alkylated at car-
vicinal hydroxyls, one at carbons 2 and 3 and the other at bon 24, sometimes methylated at carbon 25 and sometimes
carbons 22 and 23, resembling in some way the unsaturated between carbons 24 and 28.
ecdysteroids. These characteristics lead to considering as natural
A few years later Japanese scientists isolated brassinosteroids the 3-oxygenated (20β)-5α-cholestane-
castasterone (9; Yokota et al., 1982a; figure 1) from chest- 22α,23α-diols of plant origin, bearing additional alkyl or
nut gall tissue, the ketone that was then thought to be the oxy substituents (see general structures 52 and 52a, figure
putative precursor of brassinolide. Soon afterwards they 2). They can also occur conjugated especially with sugars
were able to identify, as a mixture of brassinolide-like sub- or fatty acids. Brassinosteroid analogues are compounds
stances (Ikekawa et al., 1984), the Dystilium factors A1 that show any structural similarity with natural
and B, isolated from the leaves of D. racemosum (Marumo brassinosteroids and/or brassinolide activity (Zullo et al.,
et al., 1968), that were active in the rice lamina inclina- 2002).
tion bioassay. The occurrence of the natural brassinosteroids is de-
After the first chemical syntheses of brassinolide (Fung scribed in table 1, according to the plant source. They were
and Siddall, 1980; Ishiguro et al., 1980), a great effort is isolated or detected in algae, pteridophytae, gimnosperms
being made to synthesize it and similar compounds (the and angiosperms (mono- and dicotyledons), indicating a
brassinosteroids), to isolate new brassinosteroids, eluci- probable ubiquitous distribution in the plant kingdom.
date their biosynthetic routes, verify their biological ac- Some of the biosynthetic precursors of the
tivities and their agricultural applications. brassinosteroids, such as cathasterone (53; Fujioka et al.,
1995; figure 3), 6-deoxocathasterone (54), 3-epi-6-
deoxocathasterone (55; Fujioka et al., 2000b) and 6-deoxo-
NATURAL BRASSINOSTEROIDS
28-norcathasterone (54a, Yokota et al., 2001), as well as
Since the isolation of brassinolide (1), a series of catabolites, such as cryptolide (56; Watanabe et al., 2000),
brassinosteroids [such as dolicholide (3), 28- are in some instances considered as brassinosteroids them-
homodolicholide (4), castasterone (9), dolichosterone (11), selves, but they do not fulfill all the structural require-
28-homodolichosterone (12) and typhasterol (25); see fig- ments.

BRASSINOSTEROIDS 145
Figure 1. Natural brassinosteroids.
Continue

Figure 1. Natural brassinosteroids (continued).
Figure 2. General formulae of natural brassinosteroids.

Table 1. Occurrence of natural brassinosteroids
Species Family Plant part Brassinosteroids Reference
Alnus glutinosa (L.) Gaertn. Betulaceae pollen 1, 9 Plattner et al., 1986

Apium graveolens L. Umbelliferae seeds 7 Schmidt et al., 1995c
Arabidopsis thaliana (L.) Heynh. Brassicaceae bri1-5 seedling 1, 9, 13, 30 Fujioka et al., 2000a
root-callus suspension cultures 1, 8 Konstantinova et al., 2001
seeds 6, 9 Schmidt et al., 1997
seeds, siliques 1, 9, 25, 36, 42, 43 Fujioka et al., 1998
shoots 9, 25, 36, 42 Fujioka et al., 1996
Banksia grandis Willd. Proteaceae pollen 1, 9 cf. Takatsuto, 1994
Beta vulgaris L. Chenopodiaceae seeds 9, 14 Schmidt et al., 1994
Brassica campestris var. Brassicaceae seeds, sheat 1, 2, 5, 9, 10, 13 Abe et al., 1982, 1983; Arima et al., 1984;
pekinensis Lour. Ikekawa and Takatsuto, 1984
Brassica napus L. Brassicaceae pollen 1, 9 Grove et al., 1979; Ikekawa et al., 1984
Cannabis sativa L. Cannabaceae seeds 9, 26 Takatsuto et al., 1996b; Gamoh et al., 1996
Cassia tora L. Fabaceae seeds 1, 9, 13, 25, 26 Park et al., 1993a, 1994b
Castanea crenata Sieb. Et Zucc Fagaceae galls 1, 9, 13, 36 Abe et al., 1983; Ikeda et al., 1983;
Ikekawa et al., 1984; Yokota et al., 1982a
shoots 9, 36 Arima et al., 1984
Catharanthus roseus G. Don. Apocynaceae crown gall cells 1, 9 Park et al., 1989
culture cells 1, 9, 25, 26, 35, 36, 39, 43 Choi et al., 1993, 1997; Fujioka et al., 2000b
Cistus hirsutum Theill. Cistaceae pollen 1, 9 cf. Takatsuto, 1994
Citrus sinensis Osbeck Rutaceae pollen 1, 9 Motegi et al., 1994
Citrus unshiu Marcov. Rutaceae pollen 1, 9, 25, 26 Abe, 1991; cf. Takatsuto, 1994
Cryptomeria japonica D. Don. Taxodiaceae pollen, anthers 2, 3, 4, 25, 32 cf. Takatsuto, 1994; Yokota et al., 1998;
Watanabe et al., 2000
BRASSINOSTEROIDS
Cucurbita moschata Duchesne Cucurbitaceae seeds 1, 9 Jang et al., 2000

Cupressus arizonica E. Greene Cupressaceae pollen 1, 9, 10, 11, 25, 26, 32, 36, 42, 45 Griffiths et al., 1995
Daucus carota ssp. sativus L. Apiaceae seeds 1, 9, 14 Schmidt et al., 1998
Diospyros kaki Thunb. Ebenaceae seeds 9 cf. Takatsuto, 1994
Distylium racemosum Sieb et Zucc. Hammamelidaceae galls 9, 13 Ikekawa et al., 1984
leaves 1, 5, 9, 13, 32 Abe et al., 1994
Dolichos lablab Adans. Leguminosae seeds 1, 3, 4, 5, 9, 11, 12, 36, 37 Baba et al., 1983; Yokota et al., 1982b, 1983b, 1984
Echium plantagineum L. Boraginaceae pollen 1 cf. Takatsuto, 1994
Equisetum arvense L. Equisetaceae strobilus 5, 9, 11, 13 Takatsuto et al., 1990a
Eriobottrya japonica Lindl. Rosaceae flower buds 9 cf. Takatsuto, 1994
Erythronium japonicum Decne Liliaceae pollen, anthers 25 Yasuta et al., 1995
Eucalyptus calophylla R. Br. Myrtaceae pollen 1 cf. Takatsuto, 1994
Eucalyptus marginata Sn. Myrtaceae pollen 11 cf. Takatsuto, 1994
Fagopyrum esculentum Moench Polygonaceae pollen 1, 9 Takatsuto et al., 1990b
Gingko biloba L. Gingkoaceae seeds 26 Takatsuto et al., 1996a
Gypsophyla perfoliata L. Caryophyllaceae seeds 6 Schmidt et al., 1996
Helianthus annuus L. Asteraceae pollen 1, 9, 13 Takatsuto et al., 1989
Hydrodiction reticulatum (L.) Lagerheim Hydrodictyaceae green alga 10, 14 Yokota et al., 1987a
Lilium elegans Thunb. Araceae pollen 1, 9, 25, 26 Susuki et al., 1994b
Lilium longiflorum Thunb. Araceae anthers 1, 9, 25, 32, 47, 48, 51 Abe, 1991; Abe et al., 1994; Asakawa et al., 1994;

147
Continue
Table 1. Occurrence of natural brassinosteroids (continued).
Species Family Plant part Brassinosteroids Reference 148
Soeno et al., 2000b

culture cells 26, 47, 48 Soeno et al., 2000a
pollen 47, 48 Asakawa et al., 1996
Lolium perene L. Poaceae pollen 15 Taylor et al., 1993
Lychnis viscaria L. Caryophyllaceae seeds 14, 34 Friebe et al., 1999
Lycopersicon esculentum Mill. Solanaceae 2 month old plants 9, 36, 42, 43, 45 Bishop et al., 1999
roots and shoots 38, 46 Yokota et al., 2001
shoots 9, 13, 36 Yokota et al., 1997
Marchantia polymorpha L. Marchantiaceae whole bodies 9 Kim et al., 2002

Ornithopus sativus Brot. Fabaceae seeds 9, 14 Schmidt et al., 1993a
shoots 9, 14, 36, 38, 39 Spengler et al., 1995
Oryza sativa L. Poaceae bran 27, 28, 36 Abe et al., 1995a
seeds 9, 26, 36 Ikekawa and Takatsuto, 1984; Park et al., 1993c, 1994a
shoots 1, 9, 11 Abe et al., 1984b; Shim et al., 1998
Perilla frutescens Britton. Labiatae seeds 2, 9 Park et al., 1993b, 1994a
Phalaris canariensis L. Poaceae seeds 9, 26 Shimada et al., 1996
Pharbitis purpurea Voigt Convolvulaceae seeds 9, 13 Suzuki et al., 1985
Phaseolus vulgaris L. Fabaceae seeds 1, 3, 9, 16, 17, 18, 19, 20, 21, 22, Kim, 1988; Kim et al., 1988; Yokota et al., 1983c, 1987b;
23,24, 25, 26, 29, 31, 36, 37, 40, Kim et al., 2000
41, 44, 49, 50
Phoenix dactilifera L. Arecaceae pollen 14 Zaki et al., 1993
Picea sitchensis (Bong.) Carr Pinaceae shoots 9, 25 Yokota et al., 1985
Pinus silvestris Lour. Pinaceae cambial region 1, 9 Kim et al., 1990
Pinus thubergii Parl. Pinaceae pollen 9, 25 Yokota et al., 1983a
Pisum sativum L. Fabaceae seeds 1, 7, 9, 25, 36 Yokota et al., 1996
M.A.T. ZULLO AND G. ADAM
shoots 1, 9, 36 Nomura et al., 1997

Psophocarpus tetragonolobus DC Fabaceae seeds 1, 9, 10, 36 cf. Takatsuto, 1994
Raphanus sativus L. Brassicaceae seeds 1, 9, 26, 28 Schmidt et al., 1991, 1993b
Rheum barbarum L. Polygonaceae panicles 1, 9, 14 Schmidt et al., 1995a
Robinia pseudo-acacia Oswald Fabaceae pollen 9, 25, 36 Abe et al., 1995b
Secale cereale L. Poaceae seeds 9, 10, 13, 25, 26, 33, 36 Schmidt et al., 1995b
Solidago altissima L. Asteraceae stem 1 cf. Takatsuto, 1994
Sporobolus stapfianum Gand. Graminae 1, 3, 9 Sasse et al., 1998
Thea sinensis L. Theaceae leaves 1, 9, 10, 13, 25, 26 Abe et al., 1983, 1984a; Ikekawa and Takatsuto, 1984;
Morishita et al., 1983
Triticum aestivum L. Poaceae grain 9, 25, 26, 32, 36 Yokota et al., 1994
Tulipa gesneriana L. Liliaceae pollen 25 Abe, 1991; cf. Takatsuto, 1994
Typha latifolia G. F. W. Mey. Typhaceae pollen 25 Schneider et al., 1983
Vicia faba L. Fabaceae pollen 1, 6, 9, 11, 13 Ikekawa et al., 1988; Park et al., 1988
seeds 1, 9 Park et al., 1988; Park and Hyun, 1987
Zea mays L. Poaceae pollen 9, 10, 13, 25, 26 Suzuki et al., 1986; Gamoh et al., 1990a
seeds 9, 13, 26, 36 Park et al., 1995
Some other natural products (57-75; figure 4) related The biological detection of brassinosteroids was ini-
to brassinosteroids, many of them occurring in the imma- tially performed by the bean second internode bioassay, a
ture Phaseolus vulgaris L. seeds (Kim, 1991; Kim et al., test gradually substituted by the rice lamina inclination
1994) have been reported, but their structures are incom- bioassay (Wada et al., 1981, 1984; Takeno and Pharis,
pletely elucidated (Fujioka, 1999). They include configu- 1982; Kim et al., 1990) and the wheat leaf unrolling bio-
rational isomers of known brassinosteroids (57-59, 61-69), assay (Wada et al., 1985; Takatsuto, 1994), while the im-
of a brassinosteroid catabolite (60) and brassinosteroids munological methods are less used (Horgen et al., 1984;
bearing extra oxygen or carbonyl bearing carbon atom in Yokota et al., 1990; Schlagenhaufer et al., 1991; Taylor et
ring A (70-75). al., 1993).
Due to the need to detect brassinosteroids in plant
sources, a micromethod was very quickly developed for
screening and quantification of these kinds of compounds
(Takatsuto et al., 1982), that consists in reacting the cis-
dihydroxy function with methaneboronic acid, affording
its methane- or bismethaneboronate, e.g. brassinolide
bismethaneboronate (76). Eventually isolated hydroxyl is
trimethylsilylated after boronation to afford, e.g. in case
of 2-deoxybrassinolide (7), the mixed derivative (77; fig-
ure 5). The derivatives are analysed by gas chromatogra-
phy, with retention times sensitive to small variations in
the structure of the brassinosteroids. The boronates are
analysed by mass spectrometry, either by electron impact
or chemical ionization. Selective scan ion monitoring can
also be used, due to the regularity of the fragmentation
pattern of the different types of brassinosteroids (Adam et
al., 1996, 1999). The method is being routinely used in
Figure 3. Biosynthetic precursors and a catabolite of the detection of brassinosteroids because its detection limit
brassinosteroids. is less than 10 pg (Ikekawa and Takatsuto, 1984; Ikekawa
et al., 1984; Takatsuto, 1994).
DETECTION AND CHROMATOGRAPHIC High performance liquid chromatography is the
ANALYSIS OF BRASSINOSTEROIDS method of choice for final purification in the isolation of
Due to the very small amount in which brassinosteroids natural brassinosteroids, but it is unusual for their detec-
are found in plants (ca. 10-100 µg.kg-1 in pollen, 1-100 tion in plant sources (Konstantinova et al., 2001). Reversed
µg.kg-1 in immature seeds and 10-100 ng.kg-1 in shoots phase high performance liquid chromatography is less sen-
and leaves; Adam and Marquardt, 1986; Mandava, 1988; sitive than gas chromatography for the detection and quan-
Takatsuto, 1994), special methods were developed for their tification of brassinosteroids, with a detection limit in the
detection and identification, as their isolation in pure state range of 25-100 pg, but with response linearity in the range
would demand a great amount of plant material and be of 25 pg-40 ng, in the best instances, and precision around
very tedious and expensive. The extraction of 3 %. Using the vicinal hydroxyls as derivatization sites
brassinosteroids from the plant material can be achieved the α-naphthylboronic (78; Gamoh et al., 1988), 9-
with partition and chromatographic processes in which phenanthrylboronic (79; Takatsuto et al., 1989, 1990b), 1-
extraction with methanol or methanol/ethyl acetate fol- cyanoisoindolyl-2-m-phenylboronic (80; Gamoh and
lowed by partition between water/chloroform and 80% Takatsuto, 1989), dansylaminophenylboronic (81, Gamoh
methanol/n-hexane is used as a standard procedure. A sen- et al., 1990a), m-aminophenylboronic (82; Gamoh et al.,
sitive bioassay is necessary to monitor the brassinosteroid 1992) and ferrocenylboronic (83; Gamoh et al., 1990b)
containing fractions during the chromatographic steps. acids reacted with brassinosteroids to obtain, quantita-

tively, the derivatives 84 (figure 6). The naphthylboronates 81 are indicated for fluorimetric detection with detection
84 are ultraviolet detected, with absorption maxima at limits of, respectively, 50 pg, 20 pg and 25 pg. These meth-
280 nm and detection limit of 100 pg, the ferrocenyl- ods have also been used to identify and quantify
boronates 84 are electrochemically detected with detec- brassinosteroids in plant sources (Takatsuto et al., 1989,
tion limit of 50 pg, while the boronates derived from 79- 1990a, b; Gamoh and Takatsuto, 1994; Motegi et al., 1994).
Figure 4. Natural products related to brassinosteroids with incompletely elucidated structure.
Figure 5. Brassinosteroid derivatives for gas chromatography/mass spectrometry.

A quite different strategy for the detection of The first system used for studying brassinosteroid bio-
brassinosteroids employs dansylhydrazine (85; figure 6) synthesis was feeding culture cells of Catharantus roseus
to prepare the fluorescent dansylhydrazones of 6- G. Don. (Madagascar periwinkle) with deuterium labeled
oxobrassinosteroids followed by the dansylaminophe- precursors of brassinolide (1), where campesterol (87) was
nylboronation of vicinal hydroxyls. The sensitivity of the the main component of the sterol fraction, and trace the
method is only 1.5 ng for the hydrazone of 24- deuterium labeling in the brassinosteroid fraction. The dis-
epicastasterone (86) and it has the advantage that even coveries of brassinosteroid biosynthesis deficient mutants
precursors lacking the 22α,23α-diol side chain can be de- in Arabidopsis thaliana, Pisum sativum and Lycopersicon
tected. The subsequent dansylaminophenylboronation of esculentum allowed the clarification of some steps in
the hydrazone brings the sensitivity of the method to brassinolide biosynthetic pathways. In this case, partial
100 pg (Winter et al., 1999).
recovery of their growth or development is rescued by
exogenous application of brassinolide and its precursors
or putative precursors. The blocked step is recognized as
the one in which administration of a compound does not
change the mutant phenotype. It was soon recognized that
campesterol biosynthesis deficient mutants presented
brassinosteroid deficiency, and in this case analysis of ste-
rol composition aids to locate the specific biosynthetic step
blocked. It should be mentioned that brassinosteroid-in-
sensitive mutants, i.e., mutants that can respond to all other
plant hormones but not to brassinosteroids, were also rec-
ognized in Arabidopsis, pea and tomato (Clouse and Sasse,
1998; Clouse and Feldmann, 1999).
It was verified that brassinolide biosynthesis begins
by the reduction of campesterol (87) to campestanol (88),
which is oxidized to 6α-hydroxycampestanol (89) and this
to 6-oxocampestanol (90; Suzuki et al., 1995a). Parallel
experiments showed that cathasterone (53) is the biosyn-
thetic precursor of typhasterol (25) and teasterone (26;
Fujioka et al., 1995), but the conversion of 6-
oxocampestanol (90) to cathasterone (53) or to teasterone
(26) could not be demonstrated (Fujioka et al., 1995;
Suzuki et al., 1995b; Fujioka and Sakurai, 1997b). It was
Figure 6. Brassinosteroid derivatives for reversed phase observed that teasterone (26) and typhasterol (25) are
high performance liquid chromatography.
interconvertible in periwinkle, tobacco and tomato and that
typhasterol (25) is oxidized to castasterone (9) and then to
BIOSYNTHESIS AND METABOLISM brassinolide (1; Suzuki et al., 1994a, 1995a). In periwinkle,
OF BRASSINOSTEROIDS tobacco and rice castasterone (9) is also isomerized to 3-
The elucidation of brassinosteroid biosynthesis epicastasterone (18; Suzuki et al., 1995a). As a result, a
(Sakurai, 1999) and metabolism (Adam and Schneider, probable biosynthetic route to brassinolide (1) is shown in
1999; Schneider, 2002) is important for determining what figure 7. Due to the initial conversion of campestanol (88)
are their biologically active forms and for understanding to 6α-hydroxycampestanol (89), this route is called the
how their endogenous levels are regulated to promote ad- “early C-6 oxidation pathway” (Fujioka and Sakurai,
equate plant growth and development. 1997a, b).

Figure 7. Biosynthesis of brassinolide via the early C-6 oxidation pathway.
The 6-deoxobrassinosteroids, presenting weak there is no conversion of 6-deoxoteasterone (43), 3-

brassinolide activity, were initially considered as inacti- dehydro-6-deoxoteasterone (45) or 6-deoxotyphasterol
vation products of the 6-oxobrassinosteroids. As they were (42) to their 6-oxo counterparts. These studies could not
detected in an increasing number of plant species and, in say how campestanol (88) is converted to 6-
many cases, were observed the presence of the pair 6- deoxoteasterone (43). Recent studies, however, have re-
deoxocastasterone (36)/castasterone (9), the hypothesis vealed that 6-deoxotyphasterol (42) is converted to
arose that the 6-deoxobrassinosteroids could also be bio- typhasterol (25) in Arabidopsis, to a marginal extent
synthetic precursors of 6-oxobrassinosteroids. In cultured (Noguchi et al., 2000).
periwinkle, tobacco and rice cells the conversion of 6- Although there are important differences in the bio-
deoxocastasterone (36) to castasterone was observed (9; synthesis of plant sterols, a commonly accepted route for
Choi et al., 1996). In cultured cells of periwinkle the pres- the biosynthesis of campesterol (87) and campestanol (88)
ence of 6-deoxotyphasterol (42) and 6-deoxoteasterone is depicted in figure 9 (Asami and Yoshida, 1999). Plant
(43) was also observed and, using labeled compounds, the sterols are biosynthesized from mevalonic acid, which
conversions of 6-deoxoteasterone (43) to 3-dehydro-6- originates squalene-2,3-oxide (91) that cyclizes to
deoxoteasterone (45) and then to 6-deoxotyphasterol (42), cycloartenol (92). This compound is homologated to 24-
to 6-deoxocastasterone (36) and to castasterone (9), as methylenecycloartenol (93) and demethylated to
shown in figure 8, could be demonstrated (Choi et al., cycloeucalenol (94), which is isomerized to obtusifoliol
1997). This route is called the “late C-6 oxidation path- (95). Subsequent demethylation gives rise to 4α-methyl-
way”. It was verified that both early and late C-6 oxida- 5α-ergosta-8,14,24(28)-trien-3β-ol (96), that is reduced to
tion pathways operate simultaneously in periwinkle, but 4α-methylfecosterol (97). Isomerization of the ∆8(9) double

Figure 8. Biosynthesis of brassinolide via the late C-6 oxidation pathway.
bond to ∆7(8) originates 24-methylenelophenol (98), that methyldesmosterol (102), that is reduced to campesterol (87).
is demethylated to episterol (99). This compound suffers Oxidation of this sterol to (24R)-24-methyl-4-cholesten-3-one
dehydrogenation to 5-dehydroepisterol (100) and hydro- (103) is followed by saturation of the olefinic double bond to
genation to 24-methylenecholesterol (101). Isomerization of (24R)-methyl-5α-cholestan-3-one (104) and reduction of the
the ∆ 24(28) double bond to ∆ 24(25) produces 24- carbonylic function for campestanol (88) production.
Figure 9. Biosynthesis of campestanol (88) from (3S)-squalene-2,3-oxide (91).

The 14α-demethylation of obtusifoliol (95) to 4α- Feldmann, 1999). In tomato the late C-6 oxidation path-
methyl-5α-ergosta-8,14,24(28)-trien-3β-ol (96) is encoded way seems to be the major route in brassinolide biosyn-
by a CYP51 enzyme (steroid 14α-demethylase) and thesis. Analysis of the brassinosteroid fraction in the ex-
Arabidopsis antisense AtCYP51 transgenic plants showed treme dwarf (dx) tomato mutant showed that brassinolide
dwarfism during early development, slow growth during biosynthesis is blocked in the conversion of 6-
maturation, a high obtusifoliol (95) content but did not deoxocastasterone (36) to castasterone (9; Bishop et al.,
show phytosterol deficiency (Kushiro et al., 2001). In the 1999), as evidenced by the low castasterone (9) and high
Arabidopsis fackel mutant seedlings a high content of 96 6-deoxocastasterone (36) contents in the mutant compared
and ∆8,14-unsaturated sterols was observed, and the wild to the wild type.
phenotype was not rescued by brassinosteroid application, The conversion of teasterone (26) to typhasterol (25)
indicating the blockage of the conversion of 96 to 4α- and then to castasterone (9) was also observed in cultured
methylfecosterol (97; Jang et al., 2000; Schrick et al., cells of Marchantia polymorpha (Park et al., 1999; Kim et
2000). Two Arabidopsis mutants, dwf7 and ste1, were rec- al., 2001), while in Phaseolus vulgaris the in vitro enzy-
ognized to be unable to perform the conversion of episterol matic conversion of teasterone (26) to typhasterol (25) was
(99) to 5-dehydroepisterol (100), and a third one, dwf5, is confirmed to be a two-step reaction with the intermediacy
blocked in the conversion of 5-dehydroepisterol (100) to of 3-dehydroteasterone (32; Kim et al., 2000). Although
24-methylenecholesterol (101). The Arabidopsis dwf1 this is evidence that the pathways depicted in figure 10
mutant is defective in the conversion of the last compound are common for brassinolide (1) biosynthesis in plant spe-
to campesterol (87). The mutants dim and cbb1 are defec- cies other than Arabidopsis, pea and tomato (Nomura et
tive in the conversion of 24-methyldesmosterol (102) to al., 2001), they may not be simply extended to the synthe-
campesterol (87). The Arabidopsis mutants det2 and dwf6 sis of other lactones. In the case of 28-norbrassinosteroids
are defective in the reduction of (24R)-24-methyl-4- one would expect that they could be derived from choles-
cholesten-3-one (103) to (24R)-methyl-5α-cholestan-3-one terol (105), in a series of reactions similar to those occur-
(104). The blocked biosynthetic step in the garden pea lkb ring from campesterol (87). Metabolic experiments with
mutant is the conversion of 24-methylenecholesterol (101) deuterium labeled castasterone (9) in Arabidopsis, rice,
to campesterol (87; Nomura et al., 1999), more probably tomato and periwinkle detected 28-norcastasterone (13)
the isomerization of 101 to 24-methyldesmosterol (102). as a catabolite of castasterone (9; Fujioka et al., 2000a).
A small number of brassinosteroid biosynthesis mu- The detection of 28-nortyphasterol (30) in Arabidopsis
tants were recognized in the steps between campestanol (Fujioka et al., 2000a) and of 6-deoxo-28-norcathasterone
(88) and brassinolide (1; figure 10). In the Arabidopsis (54a), 6-deoxo-28-nortyphasterol (46) and 6-deoxo-28-
dwf4 mutant the conversions of campestanol (88) to 6- norcastasterone (38) in tomato (Yokota et al., 2001) are
deoxocathasterone (54) and of 6-oxocampestanol (90) to indications that both early and late oxidation pathways are
cathasterone (53) are blocked, indicating that both sub- operative for the synthesis of 28-norbrassinosteroids from
strates (88 and 90) are recognized by the same 22α-hy- a suitable precursor such as cholestanol (88a). Feeding
droxylase (Choe et al., 1998). In the Arabidopsis mutants experiments with labeled campestanol (88), cholestanol
cpd (Szekeres et al., 1996), dwf3 and cbb3, the blocked (88a) and cholesterol (105) in Arabidopsis, tobacco and
brassinolide biosynthesis steps are the 23-hydroxylations periwinkle revealed that cholesterol (105) is converted to
of 6-deoxocathasterone (54) to 6-deoxoteasterone (43) and 4-cholesten-3-one (103a), cholestanol (88a) and 6-oxo-
of cathasterone (53) to teasterone (26). The tomato dpy cholestanol (90a), but the conversion ratios of cholesterol
mutant, an intermediate dwarf with severely altered mor- (105) to cholestanol (88a) are much smaller than those of
phology, is rescued by spraying with 6-deoxoteasterone campestanol (88) to cholestanol (88a, Nakajima et al.,
(43) and subsequent precursors of brassinolide (1) in the 2002), so that it is unlikely that 28-norbrassinosteroids are
late C-6 oxidation pathway, but not by 6-deoxocathasterone preferably biosynthesized from cholesterol (105) but more
(54), cathasterone (53) or their precursors (Clouse and probably from campesterol (87; see figure 11).

Figure 10. Biosynthesis of brassinolide (1) from campestanol (88).

Figure 11. Biosynthesis of 28-norcastasterone (13).

The brassinosteroid metabolism was mainly studied epiteasterone (110) and 3-O-β-D-glucopyranosyl-(1→4)-
in cultured cells of tomato and serradella using the corre- β-D-galactopyranosyl-24-epiteasterone (111; Kolbe et al.,
sponding 5,7,7-tris-tritiated brassinosteroids (Kolbe et al., 1997; figure 12). The enzymatic conversion of 24-
1992) as monitors in the feeding experiments. Cell sus- epiteasterone (107) to 3-dehydro-24-epiteasterone (106)
pension cultures of tomato convert 3-dehydro-24- was monitored in cytosolic tomato and Arabidopsis
epiteasterone (106 ), a putative precursor of 24- thaliana fractions using fluorescent tagging and HPLC
epibrassinolide (6), to 24-epiteasterone (107) and 24- analysis. Inhibition experiments with cathasterone (53),
epityphasterol (108; Kolbe et al., 1998) and also the con- 6-deoxocathasterone (54) and 6-deoxoteasterone (43) in-
jugated brassinosteroids 3-O-β-D-glucopyranosyl-24- dicated that the corresponding 3β-dehydrogenase is rather
epiteasterone (109; Kolbe et al., 1998), 3-O-β-D- substrate specific for β-dehydrogenation of 24-
glucopyranosyl-(1→6)-β-D-glucopyranosyl-24- epiteasterone (107; Stündl and Schneider, 2001).
Figure 12. Metabolism of 3-dehydro-24-epiteasterone

(106) in cultured cells of L. esculentum.
In tomato, 24-epicastasterone (14) is hydroxylated and

glucosylated at C-25 or C-26, yielding 112 and 113, or is de-
hydrogenated to 3-dehydro-24-epicastasterone (114), that is
reduced to 3,24-diepicastasterone (19). This compound can
be glucopyranosylated at C-2 or C-3 yielding 115 and 116 or
hydroxylated at C-25 resulting in 25-hydroxy-3,24-
diepicastasterone (117; Hai et al., 1996; figure 13).
24-Epibrassinolide (6; figure 14) was transformed to
the glucopyranosides 118 and 119, while 25-hydroxy-24-
epibrassinolide (120), obtained by enzymatic hydrolysis
of 118, was transformed exclusively to the 25-glucoside
118 in cultured cells of tomato (Schneider et al., 1994;
Hai et al., 1995). These hydroxylations are performed by
two distinct enzymes, and 25-hydroxylase proved to be a
cytochrome P450 protein, while the 26-hydroxylase seems
to be a flavin-containing monooxygenase (Winter et al.,
1997).
It was verified that serradella (Ornithopus sativus Brot.)
cell cultures degrade 24-epicastasterone (14) up to 2α,3β,6β-
trihydroxy-5α-pregnane-20-one (121; Kolbe et al., 1994; fig-
ure 15), and fatty esters 122-124 of 3,24-diepicastasterone (19)
were also produced (Kolbe et al., 1995). Trihydroxyketone
121 is formed via transformation of 24-epicastasterone (14)
to 3,24-diepicastasterone (19), which is oxidized to 20R-hy-
Figure 13. Metabolism of 24-epicastasterone (14) in cul- droxy-3,24-diepicastasterone (125) and further to the
tured cells of L. esculentum. pregnanedione 126 followed by reduction (figure 15).

Figure 14. Metabolism of 24-epibrassinolide (6) by cultured cells of L. esculentum.
Figure 15. Metabolism of 24-epicastasterone (14) by cell culture of O. sativus.
In the same system, 24-epibrassinolide (6; figure 16) In first experiments on microbial transformations of
was transformed into 2α,3β-dihydroxy-B-homo-7-oxa-5α- brassinosteroid, incubation of 24-epibrassinolide (6) with the
pregnane-6,20-dione (127; Kolbe et al., 1994), the esters fungus Cunninghamella echinulata yielded 12β-hydroxy-24-
128-130 (Kolbe et al., 1996) and into 25-hydroxy-3,24- epibrassinolide (137) and the same 12β-hydroxylation was
diepibrassinolide (131 ) and 20R-hydroxy-3,24- also observed with 24-epicastasterone (14; Voigt et al., 1993a).
diepibrassinolide (132), through the initial conversion of On the other hand, the fungus Cochliobolus lunatus trans-
24-epibrassinolide (6) to 3,24-diepibrassinolide (133;
formed 24-epicastasterone (14) to the corresponding 15β-hy-
Kolbe et al., 1996).
droxylated compound 138 (Voigt et al., 1993b; figure 18).
A purified recombinant Brassica napus steroid
The analogue 2α,3α-dihydroxy-6-oxocholestane
sulfotransferase expressed by Escherichia coli catalyses the
(139), when incubated with the fungus Mycobacterium
enzymatic sulfonation of brassinosteroids and precursors spe-
cifically at position 22, as exemplified in figure 17 (Rouleau vaccae, yielded 2α,3α,6α-trihydroxy-5α-androstane-17-
et al., 1999). It exhibited highest affinity for 24-epicathasterone one (140) and 2α-hydroxy-4-androstene-3,17-dione (141;
(134), followed by 24-epiteasterone (107). Vorbrodt et al., 1991; figure 18).

Figure 16. Metabolism of 24-epibrassinolide (6) by culture cells of O. sativus.
BIOLOGICAL ACTIVITY ON INSECTS

Brassinosteroids show striking structural similarities
with arthropod hormones of the ecdysteroid type such as
20-hydroxyecdysone (142; Adler and Grebenok, 1995;
Lafont 1997), which led to several studies on the bioactiv-
ity of brassinosteroids and analogues on insects. Inhibit-
ing and antiecdysone effects have been observed in the
Figure 17. Sulfonation of brassinosteroids and precursors course of such investigations (Richter and Koolman, 1991).
by a Brassica napus steroid sulfotransferase. Thus, from a series of tested compounds, castasterone
(9) and 22,23-diepi-28-homobrassinolide (143), a synthetic
22β,23β-stereoisomer of 28-homobrassinolide (2), inhib-
ited the evagination of imaginal disks of the Phormia terra-
novae fly (Hetru et al., 1986). The 22,23-diepi-28-
homocastasterone analogue 144 and lactone 143, to a lesser
extent, bound competitively to ecdysteroid receptors from
larvae of the Calliphora vicina blowfly, representing first
antiecdysones (Lehmann et al., 1988). Compounds 143 and
144 were shown to act as weak inhibitors of binding of the
ecdysteroid ponasterone A to the intracellular ecdysteroid
receptor from the epithelial cell line from Chironimus
tentans and gave morphological effects and inhibition of
chitin synthesis similar to the moulting hormones (Spindler
et al., 1992). Compound 144 also exhibited a binding af-
finity to an ecdysteroid receptor in last instar larvae of the
Galleria mellonella wax moth similar to ecdysone (Sobek
et al., 1993) but did not act as ecdysone antagonist in the
salivary gland degeneration in Amblyomma hebraeum
Figure 18. Brassinosteroid transformations by fungi. (Charrois et al., 1996). In the Drosophila melanogaster

II cell bioassay natural brassinosteroids showed no ago-

nistic or antagonist activity (Dinan et al., 2001). When a
series of synthetic brassinosteroid/ecdysone hybrid mol-
ecules was checked in the same test system, only com-
pound 145, exhibiting the 14α-hydroxy-7-en-6-one func-
tion characteristic of ecdysteroids like 142, showed a weak
agonistic activity (Voigt et al., 2001). Using cultured imagi- Figure 20. Brassinosteroid biotransformation by P.
nal wing discs from last-instar larvae of the Spodoptera americana.
littoralis cotton leafworm, both native brassinosteroids 24-
epibrassinolide (6) and 24-epicastasterone (14) caused 50% The above-mentioned results indicate a series of bio-
competition for binding with the tritiated ecdysteroid logical effects of brassinosteroids on insects including in
ponasterone A but no induction of evagination (Smagghe vitro cell culture and in vivo whole larvae. More detailed
et al., 2002). biological and biochemical studies using the structural
multitude of brassinosteroids are necessary and could lead
to new strategies to influence ecdysteroid-dependent steps
of insect development and new pathways for insect pest
control.
BIOLOGICAL ACTIVITY AND STRUCTURE-

ACTIVITY RELATIONSHIPS
The biological activity of brassinosteroids was initially
evaluated by the bean second internode assay (Grove et
al., 1979; Thompson et al., 1981, 1982; Mandava, 1988).
In this test auxins and cytokinins are not detected and gib-
berellins elongate the treated and upper internodes.
Figure 19. Structures of compounds 142-145. Brassinosteroids promote cell division and elongation,
swelling, curvature and splitting of the treated internode:
In other studies the cockroach Periplaneta americana these morphological alterations are concentration depen-
has been used as preferred model. In feeding experiments dent.
22,23-diepi-28-homobrassinolide (143) provoked moult- The bean first internode assay, used for evaluating the
ing retardation by about 11 days with the highest applied auxin-induced growth, was also employed for testing the
doses (Richter et al., 1987). Similarly to the effects of the structure-activity relationships of brassinosteroids (Th-
hormone 20-hydroxyecdysone (142) dose-dependent ompson et al., 1982; Meudt and Thompson, 1983;
neurodepressing effects were observed with compounds Mandava, 1988; Fuendjiep et al., 1989).
144 and, to a lesser extent, compound 143 on Periplaneta The rice lamina inclination assay, based on a test origi-
americana indicating an ecdysteroid agonistic activity nally developed for auxins (Maeda, 1965), was modified
(Richter and Adam, 1991). Also the first evidence for a for brassinosteroid detection (Wada et al., 1981, 1984).
metabolic transformation of a brassinosteroid in insects While this assay has a limit of detection of 50 ppm for
has been shown recently with this species (Schmidt et al., indolacetic acid, the limit is 0.5 ppb for brassinolide (1)
2000). Thus, an organspecific epimerization of the and 5 ppb for 28-homobrassinolide (2). A modification,
brassinosteroid to 2,24-diepicastasterone (146; figure 20) employing rice lamina of the whole seedlings pre-treated
could be detected in female insects when 24- with IAA, diminished the limit of brassinolide (1) detec-
epicastasterone (14) was fed to the cockroach. The me- tion to 0.1 ppb (Takeno and Pharis, 1982). This test is con-
tabolite was observed only in the ovaries but not in the sidered as specific for brassinosteroids and is employed to
testes of the insect and was identified by GC-MS com- detect and follow the purification of these natural prod-
parison with a synthesized sample (Voigt et al., 2002). ucts (Takatsuto, 1994; Adam et al., 1996).

The wheat leaf unrolling bioassay, introduced in 1985, bioactivity of 7-dehydro-24-epicastasterone to one tenth
responds to brassinolide (1) and castasterone (9) at a limit (153; Takatsuto et al., 1987) while in the pair 22,23,24-
of detection of 0.5 ng.mL -1 (0.5ppb), with complete un- triepicastasterone (155 )/7-dehydro-22,23,24-
rolling at brassinolide concentrations equal or higher than triepicastasterone (154) the biological activity decreases
10 ng.mL -1. In this assay gibberellic acid and cytokinins about one hundred times. The introduction of a hydroxyl
produce a small effect in the concentration range of 0.1- at 5α decreases the brassinolide activity ca. 1,000 times
10 µg.mL -1 and zeatin causes slight to complete unrolling when moving from 7-dehydro-24-epicastasterone (153) to
at 1ng-1 µg.mL-1, while abscisic acid, indolacetic acid and 7-dehydro-5α-hydroxy-24-epicastasterone (156) and about
indolacetonitrile inhibit unrolling (Wada et al., 1985). 100 times in the pair 7-dehydro-22,23,24-triepicastasterone
Other assays are less frequently employed to evaluate (155)/7-dehydro-5α-hydroxy-22,23,24-triepicastasterone
brassinosteroids structure-activity relationships, such as the (157; Takatsuto et al., 1987). A less dramatic decrease in
mung bean epicotyl elongation assay (Gregory and bioactivity on the rice lamina inclination assay has also
Mandava, 1982), the radish (Takatsuto et al., 1983b, 1984) been reported when a 5α-hydroxyl function is introduced
and tomato (Takatsuto et al., 1983b) hypocotyl elongation on 28-homocastasterone (10; Brosa et al., 1998; Brosa,
assays, and auxin-induced ethylene production by etiolated 1999; Ramírez et al., 2000a). While the introduction of a
mung bean segments (Arteca et al., 1985). 5α-fluoro group in 28-homocastasterone (10) decreases its
Although the above biological assays are not equiva- bioactivity by one order of magnitude, the same introduc-
lent, they allowed the establishment of relatively safe struc- tion in 28-homoteasterone (28) or in 28-homotyphasterol
tural activity relationships (Adam and Marquardt, 1986; (27) slightly increases their bioactivity (Ramírez et al.,
Singh and Bhardwaj, 1986; Mandava, 1988; Abreu, 1991), 2000b). The absence of an oxygen function at ring B de-
with the aid of a series of brassinosteroid analogues. creases the brassinolide activity significantly, as in the case
of 6-deoxocastasterone (36) that shows only 1 % of the
As a general rule, the most bioactive brassinosteroids
castasterone bioactivity of castasterone (9; Yokota et al.,
are of the 6-oxo-7-oxalactone type, followed by the 6-oxo
1983c; see structures of compounds 147-157 in figure 21).
brassinosteroids and the 6-deoxo brassinosteroids, that are
almost inactive (Mandava, 1988). The effect of ring A substituents on brassinolide ac-
tivity was studied in some detail in the 28-
Transforming 6-oxo-7-oxalactone to ether, thialactone,
homobrassinolide (2) series (Takatsuto et al., 1987): chang-
lactam, 6-oxa-7-oxolactone, 6-aza-7-oxalactone and 6-aza-
ing the hydroxyls from 2α,3α to 3α,4α either in 28-
7-thiolactone (Okada and Mori, 1983a; Kishi et al., 1986;
homobrassinolide (2) or in 6-oxa-7-oxo-28-
Takatsuto et al., 1987) dramatically reduce the brassinolide
homobrassinolide (150) reduces the bioactivity of 158 and
activity [e.g. brassinolide (1, 10,000) ≈ 28-
159 in one order of magnitude. 2-Deoxy-28-
homobrassinolide (2, ≈ 10,000) > 6-deoxo-28-
homobrassinolide (160) is about 100 times less active than
homobrassinolide (147, 100) ≈ 7-aza-28-homobrassinolide
28-homobrassinolide (2), while 28-homotyphasterol (27)
(148, 100) > 7-thia-28-homobrassinolide (149, 10), while
is about ten times less active than 28-homocastasterone
6-oxa-7-oxo-28-homobrassinolide (150) presents about 1%
(10). Their 3β-isomers 3-epi-2-deoxy-28-homobrassinolide
of the bioactivity of 28-homobrassinolide (2; Takatsuto et
(161) and 28-homoteasterone (28) are also ten times less
al., 1987) and 6-aza-7-oxo-28-homobrassinolide (151;
Anastasia et al., 1984) and 6-aza-7-thia-28- active than 28-homobrassinolide (2) and 28-
homobrassinolide (152) are inactive (Okada and Mori, homocastasterone (10), respectively. While 3-dehydro-2-
1983b)]. Moving from the lactone to the 6-ketone it is deoxy-28-homobrassinolide (162) and 2,3-dideoxy-28-
observed that the brassinolide activity decreases from homobrassinolide (163) are about ten times less active than
100% to 50% in the pair brassinolide (1)/castasterone (9; 28-homobrassinolide (2), 3-dehydro-28-homoteasterone
Takatsuto et al., 1983a) in the rice lamina inclination as- (164) and 2,3-dideoxy-28-homoteasterone (165) are, re-
say, while 24-epicastasterone (14) is about 3 times more spectively, ten and one hundredfold less active than 28-
active than castasterone (9) in the bean second internode homocastasterone (10). 3-Dehydroteasterone (32),
assay (Thompson et al., 1982). The introduction of a C-7/ secasterone (33) and 2,3-diepisecasterone (166) show, re-
C-8 double bond in 24-epicastasterone (14) reduces the spectively, 74%, 59% and 89% of the bioactivity of 24-

epicastasterone (14) in the rice lamina inclination assay homoteasterone (28) yields compounds active at the rice
(Voigt et al., 1995; see structures of compounds 158-166 lamina inclination assay at dosages equal or higher than
in figure 22). The replacement of 3-hydroxy function by a 50 ng per plant, but not as active as their parent compounds
3-fluoro group in either 28-homotyphasterol (27) or 28- (Galagovsky, 2001).
Figure 21. Structures of brassinosteroid analogues 147-157.

Although brassinosteroids with cis A/B ring fusion (5β 22,23-diepi-28-homobrassinolide (143) becomes two times
configuration) have not yet been isolated from natural higher than for 2,3,5,22,23-pentaepi-28-homobrassinolide
sources, they were initially synthesized to explore their (168; 14 % and 6 % respectively). In the 6-oxo series, 28-
potential antiecdysteroid activity (Brosa et al., 1994). homocastasterone (10) presents 97 % of the brassinolide
Evaluation of the brassinolide activity of these compounds activity, while 2,3,5-triepi-28-homocastasterone (169) only
by the rice lamina inclination assay, employing the Bahia 51 % (Brosa et al., 1996). However, evaluation of synthe-
cultivar and a single brassinosteroid concentration, 1µg sized 5-epibrassinolide (170a, Seto et al. 1998) and 2,3,5-
per segment, showed that either 28-homobrassinolide (2) triepibrassinolide (170b) showed a nearly complete loss of
or 2,3,5-triepi-28-homobrassinolide (167) showed 87% of bioactivity in the rice lamina inclination assay indicating that
the brassinolide (1) bioactivity. When the configuration trans-fusion of rings A/B play an essential role (Seto et al.,
of the side chain changes to 22β,23β the bioactivity of 1999; see structures of compounds 167-170 in figure 23).
Many papers have dealt with the relationship between In contrast to these findings, 26,28-dinorbrassinolide (175)
the side chain structure and brassinolide activity. When and 26,28-dinorcastasterone (176) only show bioactivity
the 24-substituents of the 22α,23α-brassinosteroids were only at 10 mg per plantlet against activity at 0.01 mg per
examined, the order of brassinolide activity is brassinolide plantlet for brassinolide (1) and 0,1 mg per plantlet for
(1) > 24-epibrassinolide (6) > 28-homobrassinolide (2) > castasterone (9; Thompson et al., 1982). As the side chain
24-epi-28-homobrassinolide (171) > dolicholide (3) > 28- is reduced so is also the biological activity: in this way the
homodolicolide (4) > 28-norbrassinolide (5), a decreasing bisnorcholanelactone 177 and the androstanelactone 178
order that is also observed in the 6-oxo series in the bean show only 2 % and 0.001 % of the brassinolide (1)
second internode assay (Mandava, 1988) and in the rice activity, respectively (Kondo and Mori, 1983; see
lamina inclination assay (Takatsuto et al., 1983a). structures of compounds 171-178 in figure 24). 21-
Introduction of a methyl group at C-25 increases the Carboxipregnanelactones (Cerny et al., 1987) and 22-
brassinolide activity ten times, at least in the pairs alkoxybisnorcholanelactones (Kerb et al., 1983) also show
brassinolide (1)/25-metilbrassinolide (172) and brassinolide activity.
dolichosterone (11)/25-methyldolichosterone (16; Mori When the configuration of the side chain hydroxyls
and Takeuchi, 1988), while the removal of the methyl was analyzed, those presenting 22α,23α stereochemistry
groups at C-25, resulting in 26,27-dinorbrassinolide (173) are more active than those presenting 22β,23β configura-
or in 26,27-dinorcastasterone (174) does not affect the tion, whether for 6-oxobrassinosteroids or for lactones, no
brassinolide activity at the rice lamina inclination assay, matter what bioassay is employed (Thompson et al., 1979,
compared to their parent compounds (Takatsuto et al., 1984). 1981, 1982; Takatsuto et al., 1983a, 1987). When the ef-

fect of the alkyl substituent at C-24 and the stereochemis- 1991; Suzuki et al., 1993). This kind of conjugation is con-
try of the hydroxyls at C-22 and C-23 were analyzed jointly, sidered to be a mechanism for brassinosteroids deactiva-
the relative order of bioactivity changes according to the tion. Luo et al. (1998) prepared a series of methyl ethers
alkyl substituent, the ring B structure and the bioassay em- of brassinolide (1) to prevent such conjugation and, em-
ployed: even so the 22α,23α,24α isomers are always the ploying the rice lamina inclination assay, verified that,
most actives (Thompson et al., 1981, 1982; Takatsuto et while brassinolide 23-methyl ether (182) showed weak or
al., 1983a). The 22α,23β- or the 22β,23α-brassinosteroids low activity even at high dosage (1,000 ng per plant), the
present little bioactivity (Takatsuto et al., 1983a, b; 22-methyl ether 183 showed an activity comparable to 24-
Fuendjiep et al., 1989). Elimination of one hydroxyl, as in epibrassinolide (6) at dosages up to 100 ng per plant and
23-deoxy-28-norbrassinolide (179) decreases the bioactiv- the 22,23-dimethyl ether 184 even at dosages up to 1,000
ity (Kondo and Mori, 1983; Takatsuto et al., 1983b) [as ng per plant (see structures of compounds 179-184 in fig-
occurs with cathasterone (53; Fujioka et al., 1995) and the ure 25). Another way of deactivation was the recently
cholestanelactone 180 (Takematsu, 1982)], that is sup- shown enzymatic sulfonation of several brassinosteroids,
pressed with the elimination of both side chain hydroxyls including 24-epibrassinolide (6), with a steroid sulfotransferase
(Thompson et al., 1982; Kondo and Mori, 1983; Takatsuto from Brassica napus. This sulfonation abolished the biologi-
et al., 1983b). Side chain brassinosteroid glycosides are cal activity in the bean second internode bioassay and was
less bioactive than their aglycones, as happens to the 23- demonstrated to be specific for the hydroxyl at position 22 of
O-β-D-glucopyranosylbrassinolide (181; Yokota et al., brassinosteroids (Rouleau et al., 1999).

A series of brassinosteroid analogues, such as 17-esters indolacetic acid, promoted rice lamina inclination at
of androstanelactones (Kohout, 1989), 2α,3α,17β- dosages as low as 0.01 ng and 0.001 ng per plant,
trihydroxy-5α-androstane-6-one (Gaudinova et al., 1995), respectively. Very recently the heterodimer hybrid 197 of
hemiesters, orthoesters and ketals of 2α,3α-cholestanediol 24-epicastasterone and dexamethasone was synthesized to
(Kerb et al., 1982a), 22-ethers of lactone 177 (Kerb et al., study the regulation of protein-protein interactions, to
1983), esters of 28-homobrassinolide (Kerb et al., 1982b)
trigger signal transduction pathways and to detect ligand-
and 2-deoxybrassinosteroids (Abe and Yuya, 1993), also
protein receptor interactions (Kolbe et al., 2002). A series
show brassinolide activity. The spirostanes 185-188
of inclusion complexes of brassinosteroid (Durán Caballero
(Marquardt et al., 1989; Arteaga et al., 1997), spirosolanes
et al., 1999) or of the spirostanic brassinosteroid analogues
189 and 190 (Quyen et al., 1994a), epiminocholestanes 191
(De Azevedo et al., 2001a) in cyclodextrins were prepared
and 192 (Quyen et al., 1994a) and solanidanes 193 and
194 (Quyen et al., 1994b) analogues also show bioactivity. aiming to improve the brassinolide activity what was
A series of the first ten nonsteroidal brassinosteroid achieved with the 24-epibrassinolide/β-cyclodextrin
analogues was synthesized recently (Andersen et al., 2001), inclusion complex 198 (De Azevedo et al., 2001b, 2002;
and the compounds 195 and 196, when co-applied with see structures of compounds 185-198 in figure 26).

It is usually assumed that a brassinosteroid is linked

to its receptor site through three points: the 2α,3α-hy-
droxyls (Wada and Marumo, 1981), the B ring lactone and
the 22α,23α-hydroxyls (Kishi et al., 1986). It was formerly
considered that the receptor affinity to the 2α,3α-hydroxyls
would be greater than to the 22α,23α-hydroxyls, as varia-
tions in side chain structure are less influential in
brassinolide activity than variations in the ring A structure
(Takatsuto et al., 1983b). A study on quantitative struc-
ture-activity relationships indicates, however, that the con-
tributions of the ring A and the side chain hydroxyls con-
figurations are 25 % and 35 % of the total of brassinolide
activity, but also that the activity of a brassinosteroid or
analogue would be greater as greatest would be the simi- Figure 27. Summary of metabolic reactions observed in
larity between the compound and brassinolide (1) itself 24-epibrassinosteroids.
(Brosa et al., 1996). Further improvement of the method-
In regard to the more flexible side-chain conforma-
ology for predicting the activity of a brassinosteroid or
tion, a series of 10 brassinosteroids were investigated by
analogue takes into account the putative H-bonding inter-
means of detailed NMR investigations, molecular model-
actions in the brassinosteroid-receptor complex (Brosa,
ing studies, and compared with data from X-ray analysis.
1999).
For the most bioactive compound brassinolide (1) the
A look at the reactions involved in the metabolism of majority of conformations in solution showed a side-chain
a brassinosteroid (see the general formula in figure 27 sum- bent towards the β-face of the steroid skeleton, whereas
marizing the reactions observed in 24-epibrassinosteroids) for the less active members like 24-epibrassinolide con-
suggests that, as different enzymes catalyze different transformations with straight side-chains or side-chains bent
formations, and as these enzymes can be located at differ- towards α-face are preferred (see partial structures 203 and
ent organelles inside the cell, there is not a single receptor 204 in figure 28; Stoldt et al., 1997; Drohsin et al., 2001).
site for a brassinosteroid but there are different receptor While these models are valuable approaches for the de-
sites in different enzymes in which different brassinosteroid sign of new brassinosteroid analogues, it must be remem-
molecules are able to exhibit one of the many bered that they may not furnish the actual active confor-
brassinosteroid physiological activities. Each receptor site mation of a brassinosteroid inside the receptor site of an
must need different structural requirements for exhibiting enzyme.
the maximal activity, and this may be the reason why there
are different structure-activity relationships according to
the bioassay employed.
The almost rigid structure of the steroidal nucleus of
the brassinosteroids is confirmed by molecular orbital cal-
culations, nuclear magnetic resonance experiments and X-
ray diffraction studies, revealing that, in the 5α-series, the
A and C rings assume a chair conformation, observed also
in the ring B of 6-oxobrassinosteroids, while in the 6-oxo-
7-oxalactones the 7-membered B ring tends to lie in the
same plane as rings C and D. In the 5β-series brassinos-
teroids, the ring A also adopts a chair conformation, but it
sets almost perpendicularly to the plane formed by the rings Figure 28. Hypothetical active conformations of the ste-
B, C and D (see partial structures 199-202 in figure 28). roidal nuclei and side chains of brassinosteroids.

In summary, the following structural features are im- and 1988 with samples given by Professor Nobuo Ikekawa)
portant for exhibiting a high brassinolide activity: i) a 6- with 24-epibrassinolide (6), 24-epicastasterone (14),
oxo-7-oxalactone function; ii) 2α,3α hydroxyls; iii) 22,23,24-triepibrassinolide (205 ) and 22,23,24-
22α,23α hydroxyls; iv) 24-alkyl substitution; v) A/B-trans triepicastasterone (155) allowed the observation of in-
ring conjunction. creases in crop yields in wheat (up to 18 % in seed weight
per ear), soybean (up to 22 % in seed weight per plant)
and bean (up to 83 % in seed weight per plant in the Ca-
PROSPECTIVE AGRICULTURAL USES
rioca-80 cultivar).
Since the beginning of the research on the isolation of
brassinosteroids from plant sources, brassins proved to be
able to promote plant growth (Mitchell and Gregory, 1972),
as well as its acceleration (Gregory, 1981; Braun and Wild,
1984; Meudt et al., 1984).
With the isolation of brassinolide (1) and the synthe-
sis of similar compounds, brassinosteroids were shown to
be useful to increase crop yield: by using brassinolide (1)
bean crop yield increased, shown by the increase of 41 %
to 51 % in the weight of seeds per plant and the leaf weight
of two different lettuce varieties increased by about 25 %
(Meudt et al., 1983).
Treatment of rice plantlets with a 5 ppm solution of
brassinolide (1) caused an increase of 22 % in fresh weight
and 31.5 % in dry weight of seeds per plant in the Taebaik
cultivar (Lim, 1987). It was also reported to increase plant
growth speed, root size, and root and stem dry weight (Kim
and Sa, 1989), to reduce the toxicity of 2,4-D and butachlor
to the plantlets (Choi et al., 1990) and to increase the per-
centage of ripe grains when cultivated at low temperature
(Irai et al., 1991). Figure 29. Structures of brassinosteroid analogues 205-213.
In barley (cv. Nosovsky 9), brassinolide (1), 28-
Application of 24-epibrassinolide (6) at 1 ppb in-
homobrassinolide (2) and 24-epibrassinolide (6) increased
creased the root growth of chick-pea, cv. Pusa 256, by
the activity of endospermic α-amylase, the weight of seeds
25 % (Singh et al., 1993). The application of the same com-
per ear, the weight of 1,000 seeds and the crop yield, be-
pound to three different chick-pea cultivars, at the flower-
sides increasing the stem diameter, causing an increased
ing stage, caused increases in seed yield, crop index, 100
resistance to lodging (Prusakova et al., 1995).
seeds dry weight and in protein and soluble sugars of the
In corn (cv. Kwangok) the ear fresh weight increased by seeds (Ramos, 1995; Ramos et al., 1995, 1997). In this
about 7 % and seed dry weight increased by 11 % to 14 % by case the crop yield (in kg.ha-1) increased by 86 %, 76 %
using brassinosteroids, while in the cv. Danok 1 the effect of and 61 % for the cultivars IAC India-4, IAC Mexico and
these treatments was depressive (Lim and Han, 1988). IAC Marrocos, respectively.
The application of 24-epibrassinolide (6) or 22,23,24- The application of 24-epibrassinolide (6) or 24-
triepibrassinolide (205; figure 25) on wheat increased epicastasterone (14) on coffee caused no significant effect
panicle weight by 25-33 % and seed weight by 4-37 % on seed setting, seed size or yield (Mazzafera and Zullo,
and decreased the sterile portion of the ear by 25-62 % 1990). Coffea stenophylla calli grew up to 237 % between
(Takematsu et al., 1988). 60 and 130 days of culturing in the presence of 24-
Experiments performed at the Instituto Agronomico epibrassinolide (6), compared with growth of up to 49 %
(M. A. T. Zullo, unpublished results obtained between 1986 in the absence of this compound (Ramos et al., 1987).

It was noted that brassinolide (1) promoted potato tu- Micropropagation processes of tropical plants, such
ber development, inhibited its germination during storage as cassava (Manihot esculenta Crantz), yam (Dioscorea
and increased resistance to infections by Phytophthora alata L.) and pineapple (Ananas comosus L. Merril), can
infestans and Fusarium sulfureum (Kazakova et al., 1991). be improved by the use of 28-homocastasterone (10) or
28-Homobrassinolide (2) and its 22,23-diepimer 143 3β-acetyl-28-homoteasterone, as suggested in a prelimi-
caused an increase of 5-24 % in the size and of 23-59 % nary study (Bieberach et al., 2000). Treatment of shoot
in the fresh weight of azuki bean plants, as well as in- apices of the marubakaido apple rootstock [Mallus
creases of 3-30 % in the size and of 8-28 % in the fresh prunifolia (Willd.) Borkh] with 5α-fluoro-28-
weight of rape plants. The application of 22,23,24-triepi- homocastasterone increased the apple rootstock multipli-
28-homobrassinolide (206) increased tomato fruit setting cation rate up to 112 % (Schaefer et al., 2002).
by 43-111 %, while with 28-homobrassinolide (2) this in- Growth stimulating effects were also found in studies
crease was 118-129 % (Mori et al., 1986). on higher fungi when the cultivation of Psilocybe cubensis
The application of brassinolide (1) on orange trees as well as of Gymnopilus purpuratus in the presence of
during flowering increased fruit setting, while when ap- 10 -2 ppm brassinosteroid 143 resulted in a two to three-
plied during fruit growth it decreased the physiological fold growth acceleration with an increasing number of
drop of fruits, causing an increased number of fruits per fruiting bodies from 1-2 to 4-7 in the first flush (Gartz et
plant, accompanied by an increase in the average fruit al., 1990; Adam et al., 1991).
weight and in the brix/acidity ratio. The increase in fruit Many other examples of brassinosteroid use for in-
setting due to decreasing physiological fruit dropping was creasing crop yield can be found in the literature (Kamuro
also observed in lemon, peach, pear, persimmon and apple. and Takatsuto, 1999; Khripach et al., 1999; Núñez Vázquez
In Citrus unshiu increased juice production and a higher and Robaina Rodríguez, 2000).
brix/acidity ratio was observed (Kuraishi et al., 1991). In The brassinosteroids can be mixed with solid excipi-
Citrus madurensis Lour. brassinolide (1) retarded fruit ents (such as talc, mica, diatomaceous earth, clay), pastes
abscission was observed (Iwaori et al., 1990). (such as lanolin) or liquids (usually water or hydroalcoholic
The application of the ethers 207 and 208 (figure 29), mixtures) for use as powders, pellets, tablets, pastes, sus-
active in the bean second internode assay, increased leaf pensions, solutions, in the presence or not of emulsifiers
width in sugar beet and the lateral diameter of the root that help homogenize the preparation. The application can
(Kerb et al., 1986). be made by spraying, spreading, coating or dipping the
Some brassinosteroid analogues, synthesized for long plants or their organs or the soil. The amount of
lasting activity in the field, at first showed some useful- brassinosteroid to be applied varies with the brassinosteroid
ness in agricultural practice. So the phenylbrassinosteroids structure, the formulation employed, the kind of plant to
209-211 (figure 29), applied to corn plants, increased their be treated and the effect desired. Usually the concentra-
sizes by 14-15 % and their weight by 23-36 % (Hayashi et tion of the brassinosteroid in the preparation ranges from
al., 1989). The lactone 212 (figure 29), an intermediate in 0.01 to 100 ppm, and it can be applied with other agro-
the synthesis of 28-homobrassinolide (2), increased rad- chemicals, such as other plant hormones or growth regu-
ish fresh weight by 13-22 %, wheat seed weight by 11-22 lators, fertilizers, herbicides, insecticides and other adju-
%, grapevine cluster weight by 9-18 %, onion fresh bulb vants (Mori, 1984).
weight by 11-18 % and rice plant weight by 21-22 % Although many brassinosteroids, such as 24-
(Kamuro et al., 1990). The mixture of epoxides 213 (fig- epibrassinolide (6), are commercially available and em-
ure 29) increased the size of soybean and corn plants and ployed in some countries, more accurate studies on dos-
the dry weight of corn seeds (Takatsuto et al., 1990c). age, method and time of application, fit brassinosteroid
It has been shown in studies with arborescent plant spe- suitability for the plant or cultivar, and association with
cies that pretreatment with 22,23-diepi-28-homobrassinolide other phytohormones are needed, since many of the re-
(143) induced increase in rooting and rooting quality in cut- sults were obtained by experiments performed in green-
tings taken from mature Norway spruce donor plants and im- houses or small fields. The preliminary results regarding
proved their viability (Rönsch et al., 1993). the increases of crop yield and antistress effects on sev-

eral plants at very low doses, and the fact of being easily second convention uses the sequence rules of Cahn, Ingold
metabolized, as seen for tomato and serradella (Adam and and Prelog (Cahn et al., 1966), according to which, briefly,
Schneider, 1999; Schneider, 2002), recommend when substituents of a saturated carbon atom, arranged in
brassinosteroids as ecologically safe plant growth promot- decreasing order of atomic number, are viewed so that the
ers (Kamuro and Takatsuko, 1999; Khripach et al., 2000) substituent of least precedence is on the remote side of the
with promising properties for practical use in agriculture carbon, are arranged clockwise, this carbon is designated
and horticulture. as R, and if they are arranged counterclockwise, this car-
bon atom is designated as S. The application of these rules
to the side chain of brassinolide (1) and 22,23-
CONCLUSION
diepibrassinolide are presented in formulae v and vi, re-
Even after more than 20 years of the isolation of
spectively. The second convention is adopted for the offi-
brassinolide (1) and other natural brassinosteroids, there
cial nomenclature of steroids (Joint Commission on Bio-
is a continued effort to isolate or detect these natural prod-
chemical Nomenclature, 1989). The symbol ξ is used for
ucts from or in many plant species, to improve the bio-
designating a stereocenter of unknown configuration.
logical or physical chemical methods for their detection,
to elucidate their biosynthesis and metabolism and to pros-
pect their bioactivity and agricultural uses.
The exploitation of brassinosteroids physiological ac-
tivity (Sasse, 1999), the comprehension of the molecular
mechanisms of their activity (Clouse, 1997; Altmann 1999)
and the synthesis of natural and artificial brassinosteroids
(Back et al., 1997; Khripach et al., 1999) are other areas
of intense activity that soon will allow the general em-
ployment of these substances in agricultural practice, due
to their peculiar characteristics in promoting plant growth,
increasing crop yield and resistance to biotic and abiotic
stress, and on being ecologically safe plant growth pro-
moters.
APPENDIX
Steroids are compounds containing the gonane skel-
eton, usually methylated at carbons 10 (C-19) and 13 (C-
18), and a side chain extending from carbon 17 (i). For
substituents above the steroidal nucleus, i.e., those point-
ing in the same direction as carbons 18 and 19 the β desig-
nation is given, and for those below the steroidal nucleus
the α notation is assigned (ii). There are two different con-
ventions for the designation of the configuration of the
substitutents in the side chain. The first is the Fieser-
Plattner convention, according to which the side chain is
placed so the longest chain extends upward from the ring
D and under the plane of the drawing (Fieser and Fieser,
1948; Plattner, 1951a, 1951b). The side chain substituents
project above this plane: those appearing at the right side
of the chain are designed as α, and those appearing at the
left as β (iii and iv, for the cholestane side chain). The Figure 30. Notations for the steroid system.

NOTE ADDED IN PROOF Arabidopsis seedlings of the det2-1 mutant resulted in 16 %

A recent publication describes the identification of the conversion to 22α-hydroxy-4-campesten-3-one (215) and 0.3
seven new brassinosteroid precursors 214-220 of the 22α- % conversion to 6-deoxocathasterone (54), accompanied by
hydroxy type (figure 31) in cultured cells of C. roseus and the accumulation of 22α-hydroxy-4-campesten-3-one (215).
in A. thaliana seedlings of wild phenotype and det2-1 Feeding C. roseus cultured cells or 7-day old Arabidopsis seed-
mutant [Fujioka S, Takatsuto S, Yoshida S (2002) An early lings of the wild phenotype with hexadeuterated 6-
C-22 oxidation branch in the brassinosteroid biosynthetic deoxocathasterone (54) resulted in the detection of 3-epi-6-
pathway. Plant Physiology 130:930-939]. deoxocathasterone (55) and 22α-hydroxy-5α-campestan-3-
one (216). Either labeled 6-deoxotyphasterol (42) or 6-
deoxoteasterone (43) were detected in these experiments.
It was also found that administration of 22α-
hydroxycampesterol (214), 22α-hydroxy-4-campesten-3-one
(215), 22α-hydroxycholesterol (217) or 22α-hydroxy-4-
cholesten-3-one (218) failed to rescue the wild phenotype when
applied to dark or light grown Arabidopsis det2-1 mutants.
On the other hand, 22α-hydroxy-5α-campestan-3-one (216),
6-deoxocathasterone (54), 22α-hydroxy-5α-cholestan-3-one
(219) and 6-deoxo-28-norcathasterone (54a) rescued the wild
phenotype when applied to dark or light grown Arabidopsis
det2-1 mutants, the last two less effectively.
These findings led the authors to propose the operation of
Figure 31. New brassinosteroid precursors detected in C.
an early C-22 oxidation subpathway in the biosynthesis of
roseus and A. thaliana.
brassinosteroids (figure 32), which could probably be linked
Feeding C. roseus cultured cells or 7-day old Arabidopsis to the late C-6 oxidation pathway via the 23α-hydroxylation
seedlings of the wild phenotype with hexadeuterated 22α- of either 3-epi-6-deoxocathasterone (55) or 6-
hydroxycampesterol (214) resulted in the detection of labeled deoxocathasterone (54) to, respectively, 6-deoxotyphasterol
22α-hydroxy-4-campesten-3-one (215), 22α-hydroxy-5α- (42) or 6-deoxoteasterone (43). These results suggest that
campestan-3-one (216), 6-deoxocathasterone (54) and 3-epibrassinosteroids are biosynthesized through a metabolic grid
6-deoxocathasterone (55), while administration of 214 to instead of two distinct or linked main pathways.
Figure 32. Early C-22 oxidation subpathway in brassinosteroid biosynthesis.

Acknowledgements: Support from Fundação de Amparo Adam G, Marquardt V (1986) Brassinosteroids. Phy-
à Pesquisa do Estado de São Paulo (grants FAPESP 1999/ tochemistry 25:1787-1799.
07907-2 and 2001/05711-5) is gratefully acknowledged. Adam G, Schneider B (1999) Uptake, transport and me-
tabolism. In: Sakurai A, Yokota T, Clouse SD (eds),
One of us (G.A.) thanks the Fonds der Chemischen
Brassinosteroids - Steroidal Plant Hormones, pp.113-
Industrie for support. 136. Springer Tokyo, Japan.
Adam G., Marquardt V, Vorbrodt HM, Hörhold C, Andreas
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BRASSINOSTEROIDS

Brassinosteroid phytohormones - structure, bioactivity

Marco António Teixeira Zullo1* and Günter Adam2

INTRODUCTION unexpected response, combining elongation (typical of

Braz. J. Plant Physiol., 14(3):143-181, 2002

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 1. Natural brassinosteroids.

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 1. Natural brassinosteroids (continued).

Figure 2. General formulae of natural brassinosteroids.

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Alnus glutinosa (L.) Gaertn. Betulaceae pollen 1, 9 Plattner et al., 1986

Cucurbita moschata Duchesne Cucurbitaceae seeds 1, 9 Jang et al., 2000

Braz. J. Plant Physiol., 14(3):143-181, 2002

Soeno et al., 2000b

Braz. J. Plant Physiol., 14(3):143-181, 2002

shoots 1, 9, 36 Nomura et al., 1997

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 4. Natural products related to brassinosteroids with incompletely elucidated structure.

Figure 5. Brassinosteroid derivatives for gas chromatography/mass spectrometry.

Braz. J. Plant Physiol., 14(3):143-181, 2002

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 7. Biosynthesis of brassinolide via the early C-6 oxidation pathway.

The 6-deoxobrassinosteroids, presenting weak there is no conversion of 6-deoxoteasterone (43), 3-

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 8. Biosynthesis of brassinolide via the late C-6 oxidation pathway.

Figure 9. Biosynthesis of campestanol (88) from (3S)-squalene-2,3-oxide (91).

Braz. J. Plant Physiol., 14(3):143-181, 2002

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 10. Biosynthesis of brassinolide (1) from campestanol (88).

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 11. Biosynthesis of 28-norcastasterone (13).

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 12. Metabolism of 3-dehydro-24-epiteasterone

In tomato, 24-epicastasterone (14) is hydroxylated and

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 14. Metabolism of 24-epibrassinolide (6) by cultured cells of L. esculentum.

Figure 15. Metabolism of 24-epicastasterone (14) by cell culture of O. sativus.

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 16. Metabolism of 24-epibrassinolide (6) by culture cells of O. sativus.

BIOLOGICAL ACTIVITY ON INSECTS

Braz. J. Plant Physiol., 14(3):143-181, 2002

II cell bioassay natural brassinosteroids showed no ago-

BIOLOGICAL ACTIVITY AND STRUCTURE-

Braz. J. Plant Physiol., 14(3):143-181, 2002

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 21. Structures of brassinosteroid analogues 147-157.

Figure 22. Structures of brassinosteroid analogues 158-166.

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 23. Structures of brassinosteroid analogues 167-170.

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 24. Structures of brassinosteroid analogues 171-178.

Figure 25. Structures of brassinosteroid analogues 179-184.

Braz. J. Plant Physiol., 14(3):143-181, 2002

Figure 26. Structures of brassinosteroid analogues 185-198.

Braz. J. Plant Physiol., 14(3):143-181, 2002

It is usually assumed that a brassinosteroid is linked

Braz. J. Plant Physiol., 14(3):143-181, 2002