Download Inhibition of Transdifferentiation into Tracheary Elements by Polar

Plant Cell Physiol. 46(12): 2019–2028 (2005)
doi:10.1093/pcp/pci217, available online at www.pcp.oupjournals.org
JSPP © 2005
Inhibition of Transdifferentiation into Tracheary Elements by Polar Auxin
Transport Inhibitors Through Intracellular Auxin Depletion
Saiko Yoshida *, Hideo Kuriyama and Hiroo Fukuda *
Department of Biological Sciences, Graduate School of Science, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo, 113-0033 Japan
;
Polar auxin transport is essential for the formation of
continuous vascular strands in the plant body. To understand its mechanism, polar auxin transport inhibitors have
often been used. However, the role of auxin in vascular differentiation at the unicellular level has remained elusive.
Using a Zinnia elegans cell culture system, in which single
mesophyll cells transdifferentiate into tracheary elements
(TEs), we demonstrated that auxin transport inhibitors
prevented TE differentiation and that high concentrations
of 1-naphthaleneacetic acid (NAA) and IAA overcame the
repression of TE differentiation. Measurements of NAA
accumulation with 3H-labeled NAA in the presence or
absence of 1-N-naphthylphthalamic acid (NPA) revealed
enhanced NAA accumulation within the cell. In the NPAtreated cells, intracellular free NAA decreased, while its
metabolites increased. Therefore, the polar auxin transport
inhibitors may prevent auxin efflux and consequently promote NAA accumulation in Zinnia cells. The excess intracellular NAA may also activate NAA metabolism, resulting
in a decrease in free NAA levels. This depletion of free NAA
may prevent TE differentiation. The decreased auxin activity in NPA-treated cells was confirmed by the fact that the
DR5 (a synthetic auxin-inducible promoter)-mediated
expression of a reporter protein was suppressed in such
cells. Gene expression analysis indicated that NPA suppressed TE differentiation at an early process of transdifferentiation into TEs. Based on these results, the interrelationship between auxin and vascular cell development
at a cellular level is discussed.
Keywords: Auxin metabolism — NAA — Polar auxin transport — Tracheary element differentiation — Zinnia elegans.
Abbreviations: BA, benzyladenine; CaMV, cauliflower mosaic
virus; CFP, cyan fluorescent protein; DMSO, dimethylsulfoxide; DR5,
synthetic auxin-inducible promoter; GH3, auxin-inducible promoter of
the soybean gh3 gene; HFCA, 9-hydroxyfluorene-9-carboxylic acid;
NAA, 1-naphthaleneacetic acid; NLS, nuclear localization signal;
NPA, 1-N-naphthylphthalamic acid; TE, tracheary element; TIBA,
2,3,5-triiodobenzoic acid; RT–PCR, reverse transcription–PCR; TLC,
thin-layer chromatography; YFP, yellow fluorescent protein
*
Introduction
Polar auxin transport has been implicated in the control of
various physiological phenomena such as apical dominance
and gravitropism (Klee and Estelle 1991). Apical–basal auxin
transport also plays a pivotal role in the formation of a continuous vascular strand (Jacobs 1952, Berleth et al. 2000, Sachs
2000, Aloni 2001, Dengler 2001, Kuriyama and Fukuda 2001,
Turner and Sieburth 2002, Fukuda 2004). Mutants with
alterations in vascular patterns have been characterized in
Arabidopsis (Sieburth 1999, Berleth et al. 2000, Ye 2002).
PIN-FORMED1 (PIN1) encodes an auxin efflux carrier
involved in polar auxin transport. PIN1 is conspicuously localized on the plasma membrane where it contacts the basal end
wall in procambial cells and xylem parenchyma cells. The pin1
mutant exhibits fused cotyledons, abnormal phyllotaxy, pinshaped inflorescences and excess vascularization. GNOM/
EMB30 (GN) encodes a guanine nucleotide exchange factor of
ADP-ribosylation factor GTPase (ARF-GEF), which is responsible for targeted recycling of PIN1 (Steinmann et al. 1999,
Geldner et al. 2003). The gn mutation causes disconnected vasculature with the formation of clustered or scattered tracheary
elements (TEs) (Mayer et al. 1991, Koizumi et al. 2000).
Therefore, well-arranged auxin efflux carriers are necessary for
normal vascular development (Gälweiler et al. 1998, Berleth
and Mattsson 2000).
Auxin-response mutants have also revealed the importance of auxin perception and signaling in vascular formation.
The MONOPTEROS (MP) gene encodes a transcription factor
that belongs to a family of auxin-response factors that respond
and act in auxin signaling. The mp mutants display the formation of discontinuous and reduced vasculatures in mature plants
(Hardtke and Berleth 1998). Arabidopsis mutants with a defect
in perceiving auxin such as axr6 and bodenlos show reduced
vasculature (Hobbie et al. 2000, Hamann et al. 2002).
Polar auxin transport is prevented by reagents that block
auxin efflux such as 1-N-naphthylphthalamic acid (NPA), 9hydroxyfluorene-9-carboxylic acid (HFCA) and 2,3,5-triiodobenzonic acid (TIBA). NPA is the most widely used auxin
transport inhibitor and is classed as a phytotropin. Polar auxin
transport inhibitors cause the enhancement of vascular cell formation in leaf margins and parallel vascular strands in the central regions in a similar fashion to the phenotype of pin1
Corresponding authors: Saiko Yoshida, E-mail, [email protected]; Fax, +81-3-5841-4462; Hiroo Fukuda, E-mail, [email protected]; Fax, +81-3-5841-4461.
2019
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Intracellular auxin affecting TE differentiation
mutants (Mattsson et al. 1999, Sieburth 1999, Berleth and
Mattsson 2000, Berleth et al. 2000, Mattsson et al. 2003).
These results suggest that polar auxin transport inhibitors disrupt auxin flow and accumulate auxin in nearby tissues. This
promotes the abnormal formation of vascular cells (Mattsson et
al. 1999). Most of these data are consistent with the auxin flow
canalization hypothesis proposed by Sachs (1991): auxin flow,
starting initially with diffusion, induces the formation of the
polar auxin transport cell system, which in turn promotes auxin
transport and leads to canalization of auxin flow along a narrow file of cells. This file of cells differentiates into a strand of
procambial cells, and eventually into strands of various vascular cells. However, at the cellular level, this hypothesis remains
elusive. For instance, what happens in cells after changes in
auxin flow? How does auxin induce the differentiation of vascular cells such as procambial cells?
Fukuda and Komamine (1980) noted that auxin and cytokinin were prerequisites for the induction of transdifferentiation
of mesophyll cells into TEs in an in vitro Zinnia culture system. In this system, mechanically isolated single mesophyll
cells transdifferentiate into TEs via procambial-like cells without cell division. This is an excellent system for the analysis of
auxin function in vascular differentiation at the cellular level. A
detailed analysis of the transdifferentiation process revealed
three distinctive stages (Fukuda 1997, Fukuda 2004). Stage 1
corresponds to a functional dedifferentiation process from isolated mesophyll cells. At stage 2, dedifferentiated cells may
restrict their potency, change into procambial cells, and then
change into xylem cell precursors. During this stage, a proteoglycan, xylogen, regulates the local and inductive cell-cell
interactions required for xylem differentiation (Motose et al.
2004). Brassinosteroids, which are synthesized actively during
stage 2, initiate stage 3. At stage 3, the differentiation of TEs
occurs from xylem precursor cells (Yamamoto et al. 1997,
Yamamoto et al. 2001).
To elucidate auxin transport and function in vascular differentiation at the cellular level, we analyzed the effects of
auxin transport inhibitors on vascular cell differentiation, auxin
metabolism and gene expression using a Zinnia culture sys-
Table 1
ation
Effects of auxin transport inhibitors on TE differentiConcentra- TE differenti- Cell division Cell viability
tion (µM)
ation (%)
(%)
(%)
NPA
TIBA
HFCA
0
10
20
50
0
0.5
1
10
0
10
20
50
29.1 ± 2.1
2.2 ± 0.1
0.6 ± 0.2
0.0 ± 0.1
30.9 ± 1.3
6.7 ± 1.2
0.8 ± 1.1
0.0 ± 0.0
30.9 ± 1.3
16.1 ± 4.5
5.1 ± 3.5
0.2 ± 0.3
15.7 ± 2.5
7.8 ± 1.6
7.7 ± 2.0
3.3 ± 1.3
32.3 ± 2.6
21.4 ± 1.5
10.9 ± 3.9
3.8 ± 2.3
32.3 ± 2.6
20.9 ± 1.7
28.3 ± 1.4
21.0 ± 4.5
66.5 ± 1.2
69.5 ± 1.0
66.2 ± 3.2
55.4 ± 4.3
76.2 ± 0.5
75.2 ± 1.1
73.4 ± 0.6
49.2 ± 2.8
76.2 ± 0.5
71.7 ± 3.0
69.8 ± 3.2
72.9 ± 2.0
Auxin transport inhibitors were added at the start of culture, and the
frequencies of TE differentiation and cell division were determined
after 96 h of culture. Data are mean values ± SDs of the results from
three replicates.
tem. The obtained results indicated that auxin transport inhibitors affected intracellular auxin metabolism and consequently
suppressed TE differentiation at the transition from stage 1 to
stage 2.
Results
Effects of auxin transport inhibitors on TE differentiation and
cell shape
We examined the effects of various auxin transport inhibitors with different chemical properties, NPA, TIBA and HFCA,
on TE differentiation. TE differentiation was efficiently
induced in the presence of 0.54 µM 1-naphthaleneacetic acid
(NAA) and 0.89 µM 6-benzyladenine (BA). The addition of
NPA at 20 µM, TIBA at 1 µM or HFCA at 50 µM to the
medium containing 0.54 µM NAA and 0.89 µM BA com-
Fig. 1 NPA-treated cells. Isolated Zinnia cells were cultured for 96 h without (A) or with 20 µM NPA (B). NPA-induced suppression of TE formation was overcome by 10.7 µM NAA (C). Scale bars indicate 100 µm.
Intracellular auxin affecting TE differentiation
Fig. 2 Effects of NPA on cell shape. Cells were cultured for 96 h
with or without 20 µM NPA. The length and width of 50 cells in three
independent samples under each condition were measured. Similar
experiments were preformed twice. Bars represent SDs.
pletely prevented TE differentiation without affecting cell viability (Table 1). These reagents also prevented cell division.
Because all of the three inhibitors examined showed similar
effects, we used NPA for further analysis.
Cells treated with 20 µM NPA became larger and more
rounded than untreated cells (Fig. 1A, B). Measurements of
cell width and length confirmed a significant increase in width,
but not in length, of NPA-treated cells (Fig. 2).
Overcoming auxin transport inhibitor-induced suppression of
TE formation by auxins
The inhibition of TE differentiation caused by NPA may
be considered simply as a result of the accumulation of excess
amounts of auxin within individual cells from the inhibition of
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auxin efflux by NPA. In order to examine this possibility, lower
concentrations than 0.54 µM NAA were applied to the NPA
(20 µM)-treated cells at the initiation of culture. However, such
lower concentrations of NAA did not restore TE differentiation. Next, additional NAA at various concentrations was
applied to cell cultures containing 20 µM NPA and 0.54 µM
NAA at the initiation of culture. As a result, high concentrations of additional NAA such as 4.8–26.3 µM almost completely overcame the NPA-dependent inhibition of TE formation
(Fig. 1C, 3A). These high concentrations of NAA also overcame the NPA-dependent inhibition of cell division (Fig. 3B).
High concentrations of NAA also reversed the TIBA- or
HFCA-dependent inhibition of TE formation (data not shown).
Similarly, we examined the restorative effects of 2,4-D and
IAA. The addition of as high a concentration as 5–10 µM IAA
(Fig. 3C), which is similar to concentrations of NAA necessary for the recovery of TE differentiation, overcame the suppression of TE formation by NPA. In contrast, as little as 0.05
or 0.5 µM 2,4-D was enough to reverse NPA-dependent inhibition of TE formation (Fig. 3E). Conversely, in the presence of
1 µM 2,4-D, cells were cultured with various concentrations of
NPA. NPA did not prevent TE differentiation strongly in the
presence of 2,4-D (Fig. 4), compared with the case of NAA.
These concentrations of IAA and 2,4-D also reversed the
TIBA- or HFCA-dependent inhibition of TE formation (data
not shown). Therefore, high concentrations of NAA and IAA,
and low concentrations of 2,4-D could overcome the suppression of TE differentiation caused by auxin transport inhibitors.
Auxin concentrations necessary for the restoration of cell division were a little lower than those for TE differentiation (Fig.
Fig. 3 Overcoming the NPA-induced inhibition of TE differentiation and cell division
with various concentrations of auxins. When
Zinnia mesophyll cells were cultured for 96 h
in D medium, which contains 0.54 µM NAA
and 0.89 µM BA, 36.1% (A) and 47.4% (C,
E) of cells differentiated into TEs, and 22.9%
(B) and 19.0% (D, F) of cells divided. Addition of 20 µM NPA to D medium completely
inhibited TE differentiation (A, C, E) and
partly inhibited cell division to 4.0% (B) and
7.3% (D, F). Additional NAA (A, B), IAA
(C, D) or 2, 4-D (E, F) at various concentrations was applied to cell cultures containing
20 µM NPA, 0.54 µM NAA and 0.89 µM
BA. The frequencies of TEs (A, C, E) and
cell division (B, D, F) were determined after
96 h in culture. Data are mean values ± SDs
of the results from three replicates.
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Intracellular auxin affecting TE differentiation
Fig. 4 Effects of NPA on TE differentiation in the presence of 2,4-D.
Various concentrations of NPA were added at the initiation of culture
in the presence of 1 µM 2.4-D. The frequencies of TE differentiation
(open circles), cell division (filled circles) and cell viability (open triangles) were determined after 96 h in culture. Data are mean values ±
SDs of the results from three replicates.
3B, D, F). In contrast, phytosulfokine-α, brassinosteroids and
BA, which induce or promote TE differentiation, did not overcome the NPA-induced inhibition (data not shown).
Intracellular levels of NAA and its metabolites
To determine why high concentrations of NAA overcame
NPA-dependent suppression of TE formation, we investigated
the levels of NAA accumulated in Zinnia cells using 3H-labeled
NAA. Cells were exposed to [3H]NAA with or without NPA at
the start of culture. When examined after 48 h of culture, total
NAA was higher in NPA-treated cells than in untreated cells
(Fig. 5A). This result is consistent with the fact that NPA prevents auxin efflux. Free and metabolized [3H]NAA in cells was
then separated by thin-layer chromatography (TLC) (Fig. 5B–
D). Similarly to previous experiments, labeled NAA was
metabolized into several derivatives (Cohen and Bandurski
1982, Smulders et al. 1990, Bartel et al. 2001). The pattern of
these metabolites did not differ between NPA-treated and
untreated cells. However, surprisingly, free NAA was
decreased by the NPA treatment, although levels of most
metabolites were higher in NPA-treated cells than in untreated
cells. These results indicated that NPA elevated total NAA
amounts, but decreased free NAA in cells.
NPA prevented auxin-responsive DR5::YFP expression in Zinnia cell cultures
To examine the effect of NPA on intracellular auxin activity, we monitored the expression of DR5::YFPn (yellow fluorescent protein with a nuclear targeting signal) in Zinnia cells
that had been cultured for 48 h in the presence or absence of
20 µM NPA (Fig. 6). Plasmids having DR5::YFPn and cauliflower mosaic virus (CaMV) 35S promoter::CFPn (cyan fluorescent protein with a nuclear targeting signal), which is a
Fig. 5 A TLC autoradiograph of NAA and its metabolites. Isolated Zinnia mesophyll cells were cultured for 48 h with 0.54 µM NAA in the
presence (NPA) or absence (control) of 20 µM NPA. After culture, cells were separated from the medium, and NAA and its metabolites were
extracted with methanol. After the radioactivity of each extract was measured using a liquid scintillation counter, the standard solution of 1-NAA
and the extracts were separated by TLC and autoradiographed. (A) Calculated total NAA concentration. (B) Calculated free NAA concentration.
(C) Radioactivity corresponding to bands 1–10 in (D). PSL (photo-stimulated luminescence) indicates the amount of radioactive signals on TLC
plate. (D) Autoradiography of a TLC plate. The values are means ± SDs of the results from 3–4 samples. The experiment was repeated twice and
yielded similar results.
Intracellular auxin affecting TE differentiation
Fig. 6 Observations of DR5 promoter activity in NPA-treated cells.
Plasmids carrying pCaMV35S::CFPn and pDR5::YFPn were co-bombarded into cultured Zinnia cells. Bright field (A, B), CFP fluorescence (C, D) and YFP fluorescence (E, F) images of cells cultured in
the D medium (A, C, E), and in the D medium containing 20 µM NPA
(B, D, F) are indicated. Note that a cell cultured in NPA-containing D
medium shows negligible levels of YFP fluorescence (F) mediated by
the DR5 promoter activity compared with that cultured in D medium
alone (E). The scale bar represents 30 µm.
marker of transformed cells, were co-bombarded into the cultured cells. Then the cells were cultured for another 6 h. YFP
fluorescence in cells expressing CFP fluorescence was
observed. We performed this experiment three times with fluorescence microscopy or confocal microscopy. In each experiment, the nucleus of all the cells expressing CFP had strong
YFP fluorescence when cultured in the absence of NPA. In
contrast, the expression level of DR5::YFP in the CFP-expressing cells in NPA-treated culture was much lower than those in
untreated culture (Fig. 6E, F). Although NPA treatment
appeared to reduce CFP expression, the reduction was much
less than the NPA-induced reduction of YFP expression (Fig.
6C, D). This result suggests that NPA reduces intracellular
auxin activity, which is consistent with the decrease in the
intracellular free NAA level caused by NPA.
Inhibition of stage 2 of TE differentiation by NPA
Pulse experiments with 20 µM NPA indicated that NPA
prevented TE differentiation completely when added before
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36 h of culture (Fig. 7A), and prevented cell division when
added before 12 h (Fig. 7B). An experiment with a combination of NPA and NAA revealed that NAA overcame NPAinduced suppression of TE formation with a delay whenever
NAA was added before 96 h. An exception was that there was
no delay when it was added at 12 h of culture (Fig. 7C). The
delay corresponded roughly to the time when NAA was added
after NPA treatment. These results indicate that NPA inhibits
the early process of TE differentiation after 12 h of culture. To
confirm this, we performed a reverse transcription–PCR (RT–
PCR) expression analysis of the following stage-specific
marker genes: ZePI-1 as a stage 1 marker, ZC4H as a stage 1
and stage 3 marker, TED 2 and TED 3 as stage 2 markers, and
ZCP4 as a stage 3 marker (Fig. 8). These markers were chosen
based on Yamamoto et al. (1997), Demura et al. (2002) and
Pyo et al. (2004). In normal culture, stages 1, 2 and 3 occur
roughly between the start of culture and 24 h, between 24 and
48 h, and between 48 and 72–96 h, respectively. NPA did not
suppress mRNA accumulation of stage 1 marker genes until
24 h, but rather increased it between 36 and 48 h. However, at
that time, the accumulation of stage 1 marker gene mRNAs
decreased in untreated cells (Fig. 8A, B). In contrast, NPA
severely suppressed the accumulation of mRNAs for the stage
2 markers TED2 and TED3 (Fig. 8C, D), and completely suppressed accumulation of the stage 3 marker, ZCP4 (Fig. 8E).
Overall, NPA seemed to suppress the termination of stage 1
and the progression of stage 2, resulting in the prevention of
TE differentiation.
Discussion
Polar transport inhibitors prevent TE differentiation by
decreasing intracellular levels of auxin
Polar auxin transport inhibitors are known to prevent
auxin efflux. We noted that all of the auxin transport inhibitors
with different structures we tested prevented TE differentiation
at physiological concentrations, which strongly suggests that
the inhibition of auxin transport represses TE differentiation at
the cellular level. This observation is consistent with previous
reports showing that TIBA treatment prevents the formation of
wall thickening of TEs in a Zinnia culture system (Burgess and
Linstead 1984, Church and Galston 1988). We first expected
that excess accumulation of intracellular auxin caused by transport inhibitors prevented TE differentiation because higher, as
well as lower, concentrations of NAA prevent TE differentiation (Fukuda and Komamine 1980). However, an exogenous
supply of decreased NAA concentrations did not cause the differentiation of TEs. Instead, quite high concentrations of NAA
overcame the repression of TE differentiation.
Recent reports demonstrated that auxin is perceived by the
component, TIR1, of ubiquitin ligase SCFTIR1 within the cell
(Dharmasiri et al. 2005, Kepinski and Leyser 2005). Based on
this information, our results may be explainable by the following two hypotheses. The first is that polar auxin transport
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Intracellular auxin affecting TE differentiation
Fig. 7 Effects of NPA pulse treatments on TE differentiation and cell division. (A and B) NPA at 20 µM was added to cell cultures at various
times during culture. After 96 h in culture, the frequencies of TE differentiation (A) and cell division (B) were determined. (C) NPA at 20 µM was
added to cells at the initiation of culture, and the amount of NAA was increased to 10.7 µM at various times during culture. The frequencies of TE
differentiation were determined at the indicated time points until 240 h of culture. Data are mean values ± SDs of results from three replicates.
Fig. 8 Changes in the accumulation of mRNAs
for various stage marker genes in cells cultured
with (open symbols) or without NPA (filled
symbols). Total RNA was isolated from Zinnia
mesophyll cells that had been cultured for the
indicated time periods with or without 20 µM
NPA. mRNA accumulation for ZePI-1 (A),
ZC4H (B), TED2 (C), TED3 (D) and ZCP4 (E)
was examined by RT–PCR. The data represent
results from two experiments.
inhibitors artificially block the uptake of extracellular auxin
into cells, leading to a depletion of intracellular auxin concentration. The second is that auxin transport inhibitors prevent
auxin efflux as their original function, which leads indirectly to
the depletion of intracellular active auxin, probably through a
disturbance in auxin metabolism, or a reduction in auxin sign-
aling activity. Several (putative) NPA-binding proteins (NBPs)
such as aminopeptidases (Murphy et al. 2002) and multidrug
resistance proteins (MDR; Noh et al. 2001, Murphy et al. 2002,
Noh et al. 2003) and BIG (Ruegger et al. 1997, Gil et al. 2001)
on the plasma membrane have been identified through genetic
and biochemical studies. Indeed, in the mdr mutants, localiza-
Intracellular auxin affecting TE differentiation
tion of the efflux carrier, PIN1, was perturbed (Noh et al.
2003). Such previous observations strongly suggest that NPA
functions specifically in auxin efflux. The fact that NPA did not
completely prevent TE differentiation in the presence of 2,4-D,
which can be transported outside cells by diffusion without any
help from efflux carriers (Imhoff et al. 2000), supports the specific function of NPA in auxin efflux. The measurements of
NAA accumulation with 3H-labeled NAA in the presence or
absence of NPA determined that NPA enhanced NAA accumulation within the cell. Almost all exogenously supplied NAA at
0.54 µM accumulated in cells after culture for 48 h in the presence of NPA, although without NPA, substantial amounts of
NAA still existed in the medium (data not shown). This result
is consistent with the blockage of auxin efflux by NPA. However, surprisingly, free intracellular NAA decreases and its
metabolites increase in NPA-treated cells.
It is widely known that auxins are rapidly metabolized to
oxidized forms or conjugates by sugars, amino acids or small
peptides in plant cells (Cohen and Bandurski 1982, Bartel et al.
2001). The profile of NAA metabolites obtained in this study
was similar to that reported in previous literature (Caboche et
al. 1984, Vijayaraghavan and Pengelly 1986, Smulders et al.
1990, Delbarre et al. 1994). This suggests that NAA metabolites in Zinnia cells might contain known metabolites such as
NAA-aspartate and NAA-glucoside. The function of auxin
metabolites is generally thought to be the regulation of auxin
transport, auxin storage, the protection of auxins from enzymatic destruction, and in the homeostatic control of free auxin
levels (Cohen and Bandurski 1982, Bartel et al. 2001). Production of such auxin metabolites is promoted by the auxins themselves (Goren and Bucovac 1973, Riov et al. 1979, Caboche et
al. 1984, Smulders et al. 1990). Because recently Staswick et
al. (2005) reported that auxin-inducible promoter of the soybean gh3 genes (GH3s) encoded IAA-amide synthetases, some
of such conjugates might be synthesized by Zinnia GH3-like
enzymes which are induced by auxin. Induction of NAA
metabolites by NAA was concentration dependent, and high
concentrations of NAA promoted NAA conjugation probably
to detoxify NAA (Caboche et al. 1984, Smulders et al. 1990).
As such, polar auxin transport inhibitors may prevent auxin
efflux to promote NAA accumulation in Zinnia cells, and the
excess intracellular NAA activates NAA metabolism, resulting
in a decrease in free NAA levels. Although high concentrations of IAA, similar to those of NAA, were needed to reverse
the NPA-dependent inhibition of TE differentiation, a 100
times lower concentration of 2,4-D was enough for the restoration (Fig. 3A, C, E). Generally, 2,4-D is known to be much
more resistant to metabolism than IAA and NAA (Ribnicky et
al. 1996). Indeed, in a Zinnia culture system, 36% of free 2,4-D
was retained even after 48 h of culture when 0.25 µM 2,4-D
was added at the start of culture (N. Shinohara personal communication). Therefore, this high resistance of 2,4-D against
cellular metabolic activity within the cell may be a reason why
low concentrations of 2,4-D overcame the NPA effect, in addi-
2025
tion to the fact that 2,4-D is released from the cell by diffusion,
not by efflux.
Furthermore, NPA compromised the auxin-responsive
DR5 promoter activity in Zinnia cells at 54 h of culture, indicating that NPA reduces intracellular auxin activity. This result
is explainable by the fact that the NPA-induced depletion of
cellular free auxin levels leads to the weak response of the DR5
promoter. This depletion of free, active NAA may prevent TE
differentiation.
Auxin depletion prevents the termination of stage 1 and the progression of stage 2 during TE differentiation
The pulse experiments with NPA indicated that NPA
effectively suppressed TE differentiation when added at 12–
36 h of culture. This time period corresponds to late stage 1 and
early stage 2 (Fukuda 1997, Fukuda 2004). A gene expression
analysis with real-time PCR indicated that NPA did not prevent the expression of stage 1 marker genes but rather promoted it (for marker genes, see Yamamoto et al. 1997). In
contrast, expression of stage 2 marker genes was substantially
repressed, and a stage 3 marker gene was completely suppressed by NPA. This result suggested that NPA prevented the
termination of stage 1 and the progression of stage 2. This
result was confirmed by expression patterns of many more
genes in a microarray (S. Yoshida, unpublished). Generally,
exogenously supplied NAA is incorporated into cells and converted into its metabolites quite rapidly. For example, half of
NAA is converted within 15 min in tobacco cells (Delbarre et
al. 1996). If this is also true in Zinnia cells, NAA depletion
may occur rapidly after NPA treatment. As such, auxin depletion does not suppress stage 1 but prevents the progression of
stage 2, resulting in the inhibition of TE differentiation. In
other words, auxin is necessary for the progression of stage 2.
A few factors such as brassinosteroids (Yamamoto et al.
1997) and xylogen (Motose et al. 2004) play pivotal roles in
the transition from stage 2 to stage 3. Therefore, inhibitors of
brassinosteroid biosynthesis have been used to prevent the
transition from stage 2 to stage 3 (Yamamoto et al. 1997,
Yamamoto et al. 2001). This provides an insight into the
molecular mechanism underlying the transition (Ohashi-Ito et
al. 2002, Ohashi-Ito and Fukuda 2003, Ohashi-Ito et al. 2005).
However, there has been no other drug that prevents the progression of a specific stage, particularly in early processes. In
this study, we found a new inhibitor that specifically affected
the termination of stage 1 and the progression of stage 2. Moreover, the inhibitory effect of NPA was reversibly overcome by
NAA. Therefore, the use of both NPA and NAA will allow a
detailed analysis of the transition mechanism from stage 1 to
stage 2 at a molecular level. Indeed, efforts to identify key
genes regulating the transition are under way using NPA and
NAA treatments.
It is well known that polar auxin transport inhibitors
prevent various physiological events such as gravitropic
responses, lateral root formation, root elongation, floral bud
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Intracellular auxin affecting TE differentiation
formation, vascular patterning and embryonic patterning
(Okada et al. 1991, Liu et al. 1993, Luschnig et al. 1998,
Mattsson et al. 1999, Sieburth 1999, Morris 2000). These inhibitions have been considered a consequence of the inhibition of
auxin transport. We demonstrated here that transport inhibitors
also affect cellular events such as auxin metabolism in addition to preventing auxin transport directly. This finding will
result in caution in interpretations of effects caused by auxin
transport inhibitors. Finally, it is still unknown why excess
NAA accumulation in cells reduces the intracellular free NAA
level. Further studies are necessary to answer this question.
Materials and Methods
Reagents
NPA (Tokyo Kasei Kogyo Ltd. Tokyo, Japan), TIBA (ICN Biomedicals Inc., CA, USA), HFCA (ICN Biomedicals Inc.), NAA
(Kanto Chemical, Tokyo, Japan), 2,4-D (Wako Pure Chemical Industry, Osaka, Japan), IAA (Wako Pure Chemical Industry), and Brassinolide (Fuji Chemical Industry, Toyama, Japan) were dissolved in
dimethylsulfoxide (DMSO; Wako Pure Chemical Industry) as stock
solutions, stored at –20°C, and added to cultures at appropriate concentrations. Unless otherwise specified, they were added to the
medium at the initiation of culture. In the medium, final concentrations of DMSO were <0.5% (v/v) in all samples. At this level, it has no
effect on TE differentiation or cell division (Fukuda and Komamine
1981).
Cell culture
The first leaves of 14-day-old seedlings of Zinnia elegans L. cv.
Canary Bird (Takii Shubyo, Kyoto, Japan) were used as source material for the isolation of mesophyll cells in suspension culture according to the method of Fukuda and Komamine (1980). Cell culture was
carried out according to Fukuda and Komamine (1982) and cells were
suspended at a density of approximately 8×104 cells ml–1. The culture
medium for the induction of TE differentiation (D medium) contained
0.54 µM NAA and 0.89 µM BA. For control cultures, in which no TEs
were differentiated, CB medium that contained only BA as a phytohormone was used. Cells were cultured in the dark at 27°C while
rotated at 10 rpm on a revolving drum.
The frequencies of TE formation (%) were determined as the
number of TEs per number of living cells plus TEs. The frequencies of
cell division (%) were determined as the number of divided cells per
number of living cells plus TEs. The number of cells was counted in
three samples under a microscope, and at least 500 cells were examined in each sample.
Cell length and width were measured using a Java-based image
analysis program, ImageJ, which was developed at the US National
Institute of Health. Measurements were made at 96 h of culture. Three
independent samples were performed for each condition. Fifty cells
from three independent cell cultures were taken to measure length and
width. This experiment was performed twice. Mean values of the
length and width of cells were compared between control and NPAtreated cells.
Measurements of auxin accumulation in cultured Zinnia cells
Cells were cultured for 48 h in a medium that contained 0.13 µM
(25 TBq mmol–1) [3H]NAA (American Radiolabeled Chemicals Inc.,
MO, USA) and 0.40 µM unlabeled NAA with or without 20 µM NPA.
After culture, cells were rinsed thoroughly to wash out contaminations
of NAA that bind to the cell walls, and then collected with Handee™
Spin Cup Columns (Pierce Biotechnology Inc., IL, USA). Cell cakes
were weighed for estimation of their volume and subjected to NAA
extraction for 18 h at room temperature with 4-fold volumes of
methanol. The radioactivity in each extract was counted using a liquid
scintillation counter (Beckman LS-6000, Beckman Coulter, Inc., CA,
USA) and dpm was calculated automatically. The total NAA concentration was estimated based on an internal standard. An 8 µl aliquot of
each extract and a standard solution of 10 µM free NAA were spotted
on silica gel TLC plates and developed in a solvent (chloroform :
methanol : acetic acid; 75 : 20 : 5 by vol.) for 1 h. Migration profiles
were monitored by scanning radioactivity on TLC plates with a BASTR2025 (Fuji Photo Film, Tokyo, Japan) and visualized by FLA2000
(Fuji Photo Film). Radioactive counts of the free and metabolized
NAA spots were measured, and the free NAA concentration was calculated by multiplying the total NAA concentration by the ratio of free
NAA to total NAA metabolites on the TLC plates.
Bombardment experiments and fluorescence microscopic observations
A nucleotide sequence including the 7×DR5 (Ulmasov et al.
1997) was fused with a YFP cDNA with a nuclear localization
sequence (NLS; BD Biosciences Clontech, CA, USA). The CaMV
35S promoter was fused to a CFP cDNA with an NLS (BD Biosciences Clontech). After 48 h of culture, plasmids containing these
sequences were co-bombarded into cells as described in Ito and
Fukuda (2002) with some modifications. Cells were cultured for
another 6 h and observed using epifluorescence (model BX-50-FLA,
Olympus, Tokyo, Japan) and confocal laser scanning (FV500, Olympus) microscopes equipped with differential interference contrast
optics. For epifluorescence microscopy, CFP and YFP signals were
visualized with standard CFP (excitation filter BP425–445, dichroic
mirror DM450 and barrier filter BA460–510) and YFP (excitation filter BP490–500, dichroic mirror DM505 and barrier filter BA515–560)
filter sets (Olympus), respectively. More than five transformed cells
were observed in each sample.
RNA preparation and reverse transcription
Total RNA was isolated by a modification of the phenol–SDS
method (Ozeki et al. 1990) from cells cultured for 0, 12, 24, 36, 48, 60,
72, 84 and 96 h. RNA samples (10 µg) were treated with 10 U of
RNase-free DNase I (Roche Diagnostics, Basel, Switzerland) in the
presence of 50 U of RNase inhibitor (Wako Pure Chemical Industry)
for 30 min at 37°C. The quality of total RNA was checked by gel
electrophoresis. Reverse transcription was performed according to the
manufacturer’s instructions with 250 ng of total RNA using oligo(dT)
as a primer. The reaction mixture included 1 pmol oligo(dT), 10 pmol
dNTP, 4 µl of first strand buffer, 0.2 µmol dithiothreitol (DTT), 40 U
of RNase inhibitor (Wako Pure Chemical Industry), 10 pg of rabbit βglobin mRNA (Novagen, Madison, WI, USA), and 200 U of SuperScript™ III (Invitrogen, Tokyo, Japan) in a total reaction volume of
20 µl. The reaction was incubated at 65°C for 5 min, 42°C for 50 min,
and then inactivated at 70°C for 15 min. The mixture was incubated at
37°C for 20 min together with 2 U of RNase H (Takara, Tokyo, Japan),
and then cDNA was purified with the microcon-PCR (Millipore,
Billerica, MA, USA).
Real-time PCR with a LightCycler
Quantitative real-time PCR was performed with a LightCycler
(Roche Diagnostics) using the LightCycler–FastStart DNA Master
SYBR Green I Kit (Roche Diagnostics) according to the manufacturer’s protocol. The following primers were used for amplification by
PCR: ZePI-1, 5′-cgattactcttgaggtggag-3′ (forward) and 5′-agaccaagtgaagtgtggac-3′ (reverse); ZC4H, 5′-tagttcctttgagaagcggagg-3′ (forward) and 5′-gtattgtagtactaagtagtaggtggg-3′ (reverse); ZeCAD2, 5′-
Intracellular auxin affecting TE differentiation
cccgtgtgaactaccgtagtaa-3′ (forward) and 5′-attggagtctgagagtgaaggc-3′
(reverse); TED2, 5′-aagtcccaagctacctcacgaa-3′ (forward) and 5′-ccgttagttttgttgttgaatgacactg-3′ (reverse); TED3, 5′-cccttctccattgtcact-3′ (forward) and 5′- caaccgatatacaagccct-3′ (reverse); and ZCP4, 5′acaccagaaatgtactaccgaagaatg-3′ (forward) and 5′-aagtaaggttcccatgttgtttccttg-3′ (reverse). The gene expression values obtained were normalized using a rabbit β-globin gene in each sample. Standard curves were
obtained using serial dilutions of plasmids that contained the amplification target gene, and the relative values of gene expression were calculated. The experiment was carried out twice with each gene.
Acknowledgments
We acknowledge Dr. Naoki Shinohara (The University of Tokyo)
for technical advice on measurements of [3H]NAA accumulation in
Zinnia cells. We thank Professor Tom J. Guilfoyle (University of Missouri, Columbia, MO, USA) for providing us with a vector having the
DR5 promoter, and Dr. Yoshikatsu Matsubayashi (Nagoya University,
Nagoya, Japan) for supplying PSK-α. We are very grateful to Dr.
Shinichiro Sawa (The University of Tokyo) for technical assistance.
This work was supported in part by Grants-in-Aid to H.F. from the
Ministry of Education, Science, Sports and Culture of Japan (No.
14036205), from the Mitsubishi Foundation and from the Japan
Society for the Promotion of Science (No. 17207004).
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(Received September 8, 2005; Accepted October 11, 2005)