1 Department of Cell and Developmental Biology, John Innes Centre, Norwich NR4
7UH, UK
2 Business Unit Bioscience, Plant Research International, PO Box 16, 6700 AA
Wageningen, The Netherlands
* Author for correspondence (e-mail: robert.sablowski{at}bbsrc.ac.uk)
Accepted 25 November 2004
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SUMMARY |
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Key words: Homeotic genes, Floral development, Transcription, AGAMOUS
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Introduction |
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The molecular basis for the combinatorial action of homeotic genes may be
that in each case, the corresponding proteins are assembled into a different
protein complex. For example, stamen development requires combination of the
homeotic genes AGAMOUS (AG), APETALA3
(AP3), PISTILLATA (PI) and at least one of the
SEPALLATA (SEP1, SEP2 and SEP3) genes, whereas
carpel development occurs when AG and SEP are expressed, but
not AP3/PI (Bowman and
Meyerowitz, 1991; Honma and
Goto, 2001
; Jack et al.,
1992
; Krizek and Meyerowitz,
1996
; Pelaz et al.,
2000
; Yanofsky et al.,
1990
). Based on protein-protein interactions in yeast and on
co-immunoprecipitation, it has been proposed that stamen development is
directed by a protein complex in which SEP3 bridges the interaction between AG
and the AP3/PI heterodimer; similarly, the direct interaction between SEP3 and
AG in yeast and suggests that these two proteins associate to control carpel
development (Honma and Goto,
2001
).
Presumably each of the complexes containing homeotic proteins selects a
different set of downstream target genes that participate in the development
of a specific organ type, although the exact composition of these complexes in
vivo, and how they select different target genes, remains unknown
(Jack, 2001). To understand
how the activity of homeotic genes is combined and translated into the
patterns of cell division and differentiation that actually shape the floral
organs, it is necessary to identify these downstream targets. However, very
little is known about the genes that function downstream of the floral
homeotic genes.
Genetic analysis has revealed some intermediate regulatory genes that
control specific aspects of floral organ development. For example, AG
activates SPOROCYTELESS (SPL), which controls sporogenesis
in both stamens and carpels (Ito et al.,
2004). SUPERMAN (SUP) controls cell
proliferation in stamen and carpel primordia and its expression depends on
AG, AP3 and PI (Sakai et
al., 2000
; Sakai et al.,
1995
). The SHATTERPROOF genes (SHP1 and
SHP2) are required in the carpel margins for differentiation of the
dehiscence zone, where later the fruit splits open to release the seeds
(Liljegren et al., 2000
).
SPATULA (SPT) controls cell differentiation at the carpel
margins and in the transmitting tract (the tissue that guides the growth of
pollen tubes towards the ovules) (Bowman
and Smyth, 1999
; Heisler et
al., 2001
), and CRABS CLAW (CRC) participates in
directing the development of tissues derived from the abaxial side of the
carpel primordium (e.g. the outer epidermis)
(Eshed et al., 1999
).
A more comprehensive view of gene expression in floral organs came from
transcript profiling experiments comparing wild type and homeotic mutants
(Wellmer et al., 2004;
Zik and Irish, 2003b
). These
experiments revealed hundreds of genes that are preferentially expressed in
different organs, but these were mostly expressed at late stages of
development and were probably only indirectly dependent on the floral homeotic
genes (Wellmer et al., 2004
).
To fully understand the program of gene expression controlled by the floral
homeotic genes, it is necessary to know how it unfolds from organ initiation
to maturity. Here, we report the results of a global analysis of the program
of gene expression triggered by AG, from the onset of organogenesis to early
stages of reproductive organ development.
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Materials and methods |
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Dexamethasone (Sigma, stock solution 10 mM in ethanol) was used at a final concentration of 10 µM in Silwet L-77 0.015%, applied directly on the inflorescence tips; for mock treatments, the solution contained the same amount of ethanol (0.1%) and Silwet L-77. After treatment, RNA was extracted from inflorescence apices and stored at -70°C until activation of AGGR was confirmed (2 weeks later).
Scanning electron microscopy (SEM)
Plants were fixed in 2.5% glutaraldehyde in phosphate-buffered saline (PBS)
at 4°C overnight, dehydrated in an ethanol series, critical-point dried in
liquid CO2, sputter-coated with gold palladium, analysed and
photographed with a Philips XL 30 FEG SEM.
RNA isolation
Total RNA was extracted using TRI reagent (Sigma) according to the
manufacturer's instructions. For array hybridisation, the RNA was cleaned up
with RNeasy columns (Qiagen) and precipitated to increase final
concentration.
Array hybridisation and analysis of expression data
Gene Chip arrays were hybridised as in the manufacturer's protocol
(Affymetrix). To calculate P-values for increase or decrease in
expression, the Wilcoxon signed-rank test
(Hubbell et al., 2002;
Liu et al., 2002
) was applied
to each pair of chips after normalisation across all probe sets, using Micro
Array Suite 5.0 (Affymetrix). To calculate fold differences in expression, raw
expression levels were imported from Micro Array Suite 5.0 into Gene Spring
5.1 (Silicon Genetics) and normalised first to the fiftieth percentile of each
chip, then across all chips before further analysis.
Reverse transcription-PCR (RT-PCR)
Total RNA (2 µg) was treated with RNase-free DNase, and first strand
cDNA was synthesised using oligo(dT) primer (Invitrogen) and Superscript RT
(Invitrogen). Aliquots of the cDNA were used as template for PCR with gene
specific primers (see Table S3 in supplementary material).
In situ hybridisation
RNA was hybridised in situ (Fobert et
al., 1996), using digoxigenin-labelled probes transcribed with T7
polymerase from linearised plasmid (pGEM-T easy, Promega) containing 3'
cDNA fragments. Colour detection was performed with BCIP/NBT according to the
manufacturer's instructions (Boehringer).
Production of recombinant AG protein
To produce AG protein, the AG ORF was PCR-amplified from pCIT1516 vector
(Yanofsky et al., 1990) and
cloned into pRSET-A (Invitrogen). BL21(DE3) pLysE cells were transformed with
the construct, and His-AG proteins were expressed under the control of the T7
promoter. To prepare recombinant His-AG, inclusion bodies were purified using
the BugBuster HT Protein Extraction Reagent (Novagen), according to the
manufacturer's instructions, dissolved in dialysis buffer (20 mM Tris, 100 mM
KCl, 1 mM EDTA, 1 mM DTT, 1 mM PMSF, 12% glycerol, pH 8.0) containing 6M urea,
dialysed overnight against the same buffer without urea and stored at
-20°C.
Electrophoretic mobility shift assays (EMSA)
Probes were made from complementary oligonucleotides (see Table S3 in
supplementary material), annealed in 20 mM Tris (pH 8.0), 50 mM NaCl, 1 mM
EDTA, labelled with 32P by filling in with DNA polymerase I (Klenow
fragment), and gel-purified prior to use. DNA-binding assays and gel
electrophoresis were essentially as described previously
(Riechmann et al., 1996).
Chromatin immunoprecipitation (ChIP)
The procedure was adapted from Ito et al. and Wang et al.
(Ito et al., 1997;
Wang et al., 2002
).
Inflorescence tissue (
1 g) of Col-0 plants was fixed with 1% formaldehyde
in MC buffer [10 mM sodium phosphate (pH 7.0), 50 mM NaCl, 0.1 M sucrose) for
1 hour under vacuum. Fixation was stopped with 0.125 M glycine, followed by
three washes with MC. The tissue was ground in liquid nitrogen, the powder was
suspended in M1 buffer [10 mM sodium phosphate (pH 7.0), 0.1 M NaCl, 1 M
2-methyl 2,4-pentanediol, 10 mM ß-mercaptoethanol, CompleteTM
Protease Inhibitor Cocktail (Roche Diagnostics GmbH, Mannhein, Germany)], the
slurry was filtrated through 55 µm mesh and centrifuged at 1000
g for 10 minutes. Subsequent steps were at 4°C unless
indicated otherwise. Filtration and centrifugation were repeated twice, then
the pellet was washed five times with M2 buffer (M1 buffer with 10 mM
MgCl2, 0.5% Triton X-100) and once with M3 buffer (M1 without
2-methyl 2,4-pentanediol). The nuclear pellet was resuspended in 1 ml Sonic
buffer [10 mM sodium phosphate (pH 7.0), 0.1 M NaCl, 0.5% Sarkosyl, 10 mM
EDTA, CompleteTM Protease Inhibitor Cocktail (Roche Diagnostics GmbH), 1
mM PMSF]. Chromatin was solubilised on ice with a probe sonicator (MSE,
Soniprep 150) by 25 cycles of 15-second pulses of half maximal power with 30
seconds cooling time between pulses. After sonication, the suspension was
centrifuged (microcentrifuge, top speed) for 5 minutes and the supernatant was
mixed with one volume of IP buffer [50 mM Hepes (pH 7.5), 150 mM KCl, 5 mM
MgCl2, 10 µM ZnSO4, 1% Triton X-100, 0.05% SDS]. The
solubilised chromatin was pre-adsorbed overnight with 7.5 µl antiserum
against CLAVATA3 (CLV3) (sc-12598, Santa Cruz Biotechnology, Santa
Cruz, CA) (used as AG-negative serum due to the lack of pre-immune serum).
After centrifugation, the supernatant was mixed with 40 µl of protein
G-Sepharose [Sigma, 50% slurry in 10 mM Tris (pH 7.5), 150 mM NaCl] and
incubated on a rotating wheel for 1 hour. After centrifuging, the supernatant
was equally divided over two tubes with 2.5 µl AG antiserum
(sc-12697, Santa Cruz Biotechnology) or 2.5 µl CLV3 serum
(control). After 1 hour on a rotating wheel and centrifugation, the
supernatant was mixed with 20 µl protein G-Sepharose (Sigma) before
incubation for another hour on the rotating wheel. The protein G-Sepharose
beads were washed five times with 1 ml IP buffer for 10 minutes at room
temperature. Elution with 0.1 M glycine, 0.5 M NaCl, 0.05% Tween-20 (pH 2.8)
was as described (Wang et al.,
2002
). The eluate was treated with 1 µl RNase A (10 mg/ml) and
proteinase K (to final of 0.5 mg/ml). After overnight incubation, a second
aliquot of proteinase K was added and incubated at 65°C for 6 hours. After
phenol/cloroform, then chloroform extraction, DNA was precipitated with 2.5
volumes of ethanol, one-tenth volume 3M NaAc (pH 5.4) and 1 µl glycogen,
and resuspended in 10 µl of 10 mM Tris (pH 8.0).
ChIP PCR was performed to reveal if a specific DNA fragment was enriched in the immunoprecipitated DNA sample compared with the pre-immune DNA sample. Primers were designed around the consensus AG binding sites and control primers were made for regions lacking the consensus AG binding site. Template ChIP DNA was diluted, amplified for 35 to 40 cycles (see Fig. 5), and analysed on a 1.5% agarose gel, followed by scanning with a Molecular Imager FX-PRO Plus (Bio-Rad Laboratories, Hercules, CA). The primer sequences were (5' to 3'):
|
AP3, CGGAGCTCCGTTAATAAATTGACG and TTTGGTGGAGAGGACAAGAGA;
AP3 exon 7, AACATGTTTTGGTGAATTAGGAA and GCACCAGCAAACCTTTTAGC;
GA4, TTGTCCCTTTATATACGCATTAATCA and GAGACCAAGAGGAGGCAAAA;
AG, TGGTCTGCCTTCTACGATCC and CAACAACCCATTAACACATTGG;
SEP3, CGGCCATATCCACTTTTACG and TTTTTGGGATAATTTTACTTTCCAC; and
EIF4A1 control, TCTTGGTGAAGCGTGATGAG and GCTGAGTTGGGAGATCGAAG.
![]() |
Results |
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In plants treated in parallel with solution lacking DEX, organogenesis was not seen, whereas the frequency of DEX-induced organogenesis after a single DEX treatment ranged from between 30% and 100% of plants in different experiments. Individual DEX-treated plants showed an all-or-nothing response (i.e. either robust organ induction in all treated inflorescence apices, or no induction). AGGR was still expressed in plants that failed to initiate organs in response to DEX (not shown), so transgene silencing was unlikely to be the cause of the variable organ induction. The all-or-nothing response suggested that organ induction was a bistable switch (see Discussion).
Global analysis of gene expression during AG-induced organogenesis
To screen for genes whose expression changed in ap1-1, cal-1
meristems after AG activation, we used the Arabidopsis ATH1
high-density oligonucleotide array (Affymetrix). Three time points were chosen
after a single DEX treatment of 35S::AG-GR, cal-1, ap1-1 meristems:
one day, when no morphological changes were visible, three days, when the
earliest signs of organ primordia were seen, and seven days, when stamen and
carpel primordia were recognisable (Fig.
1G-I). To ensure that the samples came from plants in which
organogenesis had been induced, a few treated meristems were left on each
plant and organ development was checked after two weeks. For each time point,
two independent samples with AGGR activated were compared with two
mock-treated controls, giving four possible combinations of treatment versus
control. Genes up- or downregulated were defined independently for each time
point as those with a statistically significant change in all four
treatment/control pairs (Wilcoxon signed-rank test, P<0.05)
(Hubbell et al., 2002;
Liu et al., 2002
) and a mean
change of at least twofold.
Using these filtering criteria, 149 of the 22,810 genes represented on the
array were upregulated in at least one of the three time points
(Fig. 2A, and Tables S1 and S2
in supplementary material). Based on their predicted molecular function, the
majority of these genes fell into three classes: unknown function (50),
DNA-binding proteins (38) and metabolic enzymes (30)
(Fig. 2C). The set of
upregulated genes contained most of the known genes with a specific role in
stamen and/or carpel development, including AG itself
(Yanofsky et al., 1990),
AP3 (Jack et al.,
1992
), PI (Goto and
Meyerowitz, 1994
), SEP1, SEP2 and SEP3
(Pelaz et al., 2000
),
SUP (Sakai et al.,
1995
), CRC (Bowman and
Smyth, 1999
) and SHP1, SHP2
(Liljegren et al., 2000
).
JAGGED (JAG) (Dinneny et
al., 2004
; Ohno et al.,
2004
), which controls the development of leaves and floral organs,
was also activated, in addition to four (At2g01520, At3g04960, At4g21590,
At5g57720) out of ten uncharacterised genes whose expression correlated with
that of floral homeotic genes during floral induction
(Schmid et al., 2003
). Thus
our array experiment independently detected many of the genes expected to
function downstream of AG, based on previous genetic and array-based
experiments.
|
A set of 1453 genes expressed mostly at relatively late stages in specific
floral organs has been identified by comparing the transcripts in wild-type
and homeotic mutant flowers (Wellmer et
al., 2004). The overlap between these genes and our list of
AG-regulated genes is relatively small (20 of the 149 AG-activated
genes and six of the AG-repressed genes; see Tables S1 and S2 in supplementary
material), suggesting that the transcriptional program in early organogenesis
is distinct from that in late organs.
Genes that showed sustained activation are expressed in wild-type carpel and stamen development
To confirm independently of the array data that we have identified genes
controlled by AG, we focused on genes that were activated at multiple time
points after AG induction. A set of twelve genes were upregulated on day 1 or
3 and then remained activated until day 7
(Fig. 3A). This set includes
four well-known regulators of stamen or carpel development (AP3, CRC, AG,
SEP3), and two genes implicated in the biosynthesis of the growth
regulator, gibberellin: GA4 encodes an enzyme that catalyses the
production of bioactive gibberellin
(William et al., 2004;
Williams et al., 1998
) and
ATH1 encodes a homeodomain protein proposed to regulate gibberellin
biosynthetic genes (Garcia-Martinez and
Gil, 2001
). The remaining six genes encode a B3 domain protein
(At3g17010), a zinc-finger protein (At1g13400) related to SUP, a
homologue (At3g11000) of a protein implicated in somatic embryogenesis in
carrot (Schrader et al.,
1997
), a predicted bifunctional nuclease (At4g21590), a WD-domain
protein (At1g47610) and a protein (At1g02190) similar to CER1, which is
involved in the synthesis of epicuticular wax and in pollen development
(Aarts et al., 1995
).
|
|
One caveat of detecting AG binding sites is that the frequency of CArG boxes in Arabidopsis genes is high: our search criteria detected at least one match in 49% of 27,186 upstream 3 kb sequences (www.arabidopsis.org/cgi-bin/patmatch/nph-patmatch.pl). The likelihood of finding a match in eight out of ten genes, however, is relatively low (4.8%, assuming binomial distribution and 49% likelihood for any single gene). Thus our subset of 12 genes was enriched for AG binding sites. A comparable enrichment was not seen for the complete set of AG-activated or repressed genes (matches were found in the upstream 3 kb sequences for 61% of the upregulated genes and 56% of downregulated genes), possibly because the complete set includes indirect AG targets.
Another caveat of the in vitro binding results is that multiple MADS domain
proteins recognise similar sequences in vitro
(Riechmann et al., 1996), so
the CArG boxes might be targeted in vivo by MADS domain proteins other than
AG. To confirm binding to AG in vivo, we used chromatin immunoprecipitation
(ChIP) for a subset of genes of particular interest: AG, AP3, SEP3
and CRC (which suggested that AG activated itself and most
of the other regulators of stamen and carpel identity); and GA4
(which suggested that another role of AG is to promote gibberellin
biosynthesis). Fragments of these genes containing the in vitro-detected AG
binding sites were enriched in immunoprecipitates obtained with antibodies
against AG, but not with an unrelated antibody
(Fig. 5C). By contrast,
fragments that lacked AG binding sequences, such exon 4 of EIF4A1
(Fig. 5C) and exon 7 of
AP3 (not shown), were detected to the same background levels with
both antibodies. Thus AG interacted in vivo with predicted regulatory
sequences of AG, AP3, CRC, SEP3 and GA4.
AG and AP1 maintain AP3 expression during organogenesis
The activation of AP3 by AG was not predicted by previous genetic and
molecular analysis, particularly because AP3 is expressed normally in
ag mutants (Jack et al.,
1992) (Fig. 6B).
This, however, could be due to redundant activation by AP1
(Lamb et al., 2002
;
Ng and Yanofsky, 2001
), which
is normally repressed by AG in the centre of the floral bud
(Gustafson-Brown et al., 1994
)
and could take over AP3 activation in the innermost organs of
ag mutant flowers. To test this idea, we compared AP3
expression in the ag-3 mutant, in the ap1-1 mutant and in
the double mutant (Fig. 6). In
the ag-3 mutant, stamens and carpels are replaced by additional
whorls of sepals and petals (Yanofsky et
al., 1990
) (Fig.
6A). As expected, AP3 expression was readily detected in
stage 3 buds and persisted throughout the development of both normal and
ectopic petals of the ag-3 mutant
(Jack et al., 1992
)
(Fig. 6B). In the
ap1-1 mutant, petals are mostly absent and sepals are replaced by
leaf-like organs that often subtend ectopic flowers
(Mandel et al., 1992
)
(Fig. 6C). In this mutant,
AP3 expression was normal in stage 3 and continued throughout stamen
development (Fig. 6D). Like
ag-3, the ag3-3, ap1-1 double mutant flower produced an
indeterminate number of organs, which were leaf-like and subtended secondary
flowers (Fig. 6E), similar to
the first whorl organs of ap1-1. In the double mutant, early
AP3 expression showed the normal pattern in both the primary and
secondary flowers, while expression in later organ development was abolished
(Fig. 6F).
|
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Discussion |
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Previously, auto-regulation of floral homeotic genes was known only for
AP3 and PI, and their orthologues in snapdragon,
DEF/GLO. In early buds, these genes are activated
independently of each other, and, where they overlap, a positive-feedback loop
is established that maintains their expression during petal and stamen
development (Jack et al.,
1994; Schwarz-Sommer et al.,
1992
). Activation of AP3 by AP3/PI is likely to be
direct, whereas activation of PI requires an intermediate protein
synthesis step (Honma and Goto,
2000
). In the case of AP3, the auto-regulatory loop is
required only in stamens: AP3 expression is still maintained in the
sepal-like organs that replace petals in the pi-1 mutant
(Jack et al., 1992
). This is
an important point, because it shows that AP3 expression can be
uncoupled from the organ identity directed by AP3/PI, and
therefore the absence of AP3 expression in the developing organs of
ag-3, ap1-1 double mutants was not a trivial consequence of the fact
that these organs were neither petals nor stamens. The requirement of
AG to maintain AP3 expression when this role cannot be
fulfilled by AP1 supported the idea that AG also
participates in AP3/PI regulation.
The co-ordinated regulation of AG, AP3, PI and SEP3 would
be expected if, as proposed by recent models, these proteins function together
in the same protein complexes (Honma and
Goto, 2001; Jack,
2001
). In particular, if the predicted protein complexes are
correct, the AP3/PI auto-regulatory loop should also require
either AG or AP1 (which has also been proposed to form a
complex with AP3/PI and SEP during petal development)
(Honma and Goto, 2001
). Our
results confirmed this prediction.
However, if AG can only function when complexed with other MADS-domain proteins, then initiation of organogenesis by AGGR must have relied on partner proteins already present in the cal-1, ap1-1 meristems. One possibility is that a low level of AG-independent expression of SEP, AP3 and PI genes provided the required partners. This initial expression could be controlled by the same mechanism that activates these genes independently of each other in early wild-type buds. The need to establish a regulatory loop to amplify initially limiting levels of its partners may be the reason why a single activation of AGGR in cal-1, ap1-1 meristems resulted either in no response, or in robust organogenesis in an apparently random fashion.
In addition to AP3, CRC was strongly activated by AG.
This was not expected because of the genetic evidence that CRC can
function in the absence of AG. In the ag-1, ap2-2, pi-1
triple mutant, in spite of the loss of AG function, the floral organs
develop several carpelloid features, such as stigmatic cells and ectopic
ovules. In this background, loss of CRC function caused a clear
reduction of these carpelloid features, showing that CRC does not
require AG to direct carpel development
(Alvarez and Smyth, 1999). Our
results suggest that although independently activated, CRC expression
is reinforced by AG. Previous genetic results suggest that this
reinforcement may be mutual: loss of CRC weakens AG
function, causing the heterozygous ag-1/AG plants, which normally
have a wild-type phenotype, to show a partial ag loss-of-function
phenotype (Alvarez and Smyth,
1999
). It remains to be tested whether this occurs because
CRC also activates AG, participating in the auto-regulatory
loop.
We also saw that, at least in the cal-1, ap1-1 backgound,
AG activated its own transcription. This could be inferred
independently of the array experiments, from the fact that the endogenous
AG was required for organogenesis in cal-1, ap1-1 plants
after transient activation of AGGR, and was supported by the
chromatin immunoprecipitation results. One difficulty with the idea that
AG auto-regulates, however, is that AG is still expressed in
the inner organs of ag-1 mutant flowers
(Gustafson-Brown et al.,
1994). Thus if AG activates itself during normal
development, this activity must be redundant. As discussed above, if
CRC participates in the AG regulatory loop, then
CRC activity might account for the continued AG expression
in ag flowers.
Combined with the published data, our results suggest a model for how AG
and other floral organ identity genes are coordinately regulated
(Fig. 7). In stamen
development, AG, AP3, SEP3 and PI are initially expressed independently of
each other. Where their expression overlaps, the predicted AG/SEP3/AP3/PI MADS
protein complex (Honma and Goto,
2001; Jack, 2001
)
maintains and amplifies their expression. In carpel development, the predicted
AG/SEP3 complex may establish a similar feedback loop, which also reinforces
CRC expression.
|
Activation of GA4 by AG may be another branch of the
homeotic gene autoregulatory loop. There may be, however, additional functions
for gibberellin in floral organogenesis. Another gibberellin biosynthetic
gene, encoding GA20-oxidase, is repressed by genes that maintain
undifferentiated cells in the meristem, and activated in the leaf primordia
that emerge from the meristem (Hay et al.,
2002; Sakamoto et al.,
2001
). This suggests that gibberellin may have a more general role
in the transition from meristem identity to organogenesis. This idea seems
inconsistent with the fact that organ emergence is normal in
gibberellin-deficient mutants, both during the vegetative phase and in flowers
(the floral defects in ga1-3 become visible only at later stages of
development) (Goto and Pharis,
1999
; Wilson et al.,
1992
). However, even severe mutants such as ga1-3, still
produce low levels of gibberellin (Hedden
and Phillips, 2000
), which might be sufficient for the proposed
functions in early organ development. Although it is not clear what these
functions might be, the known role of gibberellin in controlling cell growth
and division (Yang et al.,
1996
) suggests that it might play a role in the localised changes
in growth that drive the emergence of organ primordia from the meristem. If
this is true in the floral organ primordia, then gibberellin could be part of
the link between homeotic genes and the cellular behaviour that shapes floral
organs.
Global view of gene expression in stamen and carpel primordia
From the predicted protein functions of the 149 genes that were upregulated
by AG, two prominent features emerged (Fig.
2). First, genes expected to function in transcriptional control
were over-represented (26%), compared with their total frequency in the genome
(5.9%) (Riechmann and Ratcliffe,
2000). The fraction of regulatory genes increased over the time
course from 13% (day 1) to 28% (day 3) to 34% (day 7). This contrasts with
more mature organs, where the frequency of regulatory genes was 5.5%, similar
to their representation in the genome
(Wellmer et al., 2004
), and
suggests that up to 7 days after organ initiation much of the program of gene
expression downstream of AG was concerned with refining patterns of gene
expression. Complex cascades of transcription factors, as seen in early
development of Drosophila and sea urchin, also cause delayed
responses to initial inputs and have been proposed to function as timing
devices during development (Rosenfeld and
Alon, 2003
).
Second, of the 36 predicted DNA-binding proteins that were upregulated at
day 7, 53% belonged to two transcription factor families (10 B3 domain, PFAM
profile PF02362, and 9 MADS domain, PF00319). MADS domain proteins play a
prominent role in floral development and the diversification of this family
correlates with the evolution of plant reproductive structures
(Theissen et al., 2000). Our
data suggests that the B3 domain family has undergone a comparable
diversification of roles in reproductive development.
Developmental genetics has identified many regulatory genes whose
expression determines where and when a specific structure or organ develops.
The problem of understanding how regulatory gene expression is translated into
complex multicellular structures is universal, and has led to a number of
attempts to describe the gene expression programs controlled by these
regulators (Furlong et al.,
2001; Livesey et al.,
2000
; Michaut et al.,
2003
). Like other global descriptions of changes in gene
expression during development, however, our view of gene expression under AG
has two limitations. First, it is unlikely to be complete, because we cannot
guarantee that genes with very low or localised expression were not missed,
and because of the difficulties associated with detecting downregulation if it
occurs only in a subset of the cells. Second, the set of genes controlled by
AG probably cannot be organised within a single network of interactions,
because they may represent the overlap of multiple programs of gene expression
that run in parallel in different regions and cell types of organ
primordia.
In spite of these limitations, the list of genes controlled by AG will provide a basis for the functional analysis of intermediate regulators of early organogenesis, and will provide target promoters that are needed to test current models for the molecular basis of how homeotic genes act combinatorially.
Supplementary material
Supplementary material for this article is available at
http://dev.biologists.org/cgi/content/full/132/3/429/DC1
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ACKNOWLEDGMENTS |
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