Department of Plant and Microbial Biology, 111 Koshland Hall, University of California, Berkeley, CA 94720, USA
* Author for correspondence (e-mail: feldman{at}nature.berkeley.edu)
Accepted 23 December 2002
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SUMMARY |
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Key words: Auxin, Root, Quiescent center, Redox regulation, Maize
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INTRODUCTION |
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How a QC forms is not known, but it is suggested that it may arise as a
consequence of polar auxin transport (Kerk
and Feldman, 1995; Kerk et
al., 2000
). The availability of many Arabidopsis mutants
perturbed in their auxin signaling, such as mp, ett, aux1 and
pin1, all point to a central role for auxin in the establishment and
maintenance of organization in root meristems
(Sabatini et al., 1999
). The
recent characterization of AtPIN4, a member of the PIN family of putative
auxin efflux carriers, suggests a model in which QC establishment occurs
because of the development of an auxin sink
(Friml et al., 2002
).
According to this model, root meristems organize because of a `sink-driven
morphogenetic auxin gradient'. Thus, QC formation can be viewed as a
consequence of a sink-dependent gradient in auxin. What is not certain,
however, is the mechanism (intermediate steps) by which auxin specifies QC
formation. In this regard, it has recently been proposed that ascorbic acid
(AA) may serve as an intermediate linking auxin and QC establishment
(Kerk and Feldman, 1995
).
Since this suggestion, a number of investigators have provided data supporting
a role for AA and/or glutathione (GSH) in root meristem establishment. As in
the case of ascorbate, which was shown to be at low levels in the QC of maize
(Kerk and Feldman, 1995
),
measurements of glutathione in intact roots of Arabidopsis similarly
showed reduced levels in the QC (Fricker
et al., 2000
;
Sánchez-Fernández et al.,
1997
). Additional evidence for a role for GSH in root meristem
organization comes from Arabidopsis mutated in the gene ROOT
MERISTEMLESS (RML1), which encodes for the first enzyme of
glutathione biosynthesis (Vernoux et al.,
2000
). Mutations in this gene lead to a reduction in adequate
levels of GSH and plants are unable to form an active postembryonic root
meristem. But these mutants can be rescued (an organized root meristem
reforms) by providing seedlings with GSH
(Vernoux et al., 2000
). The
mechanism(s) by which AA and/or GSH influence the development of root
meristems is not certain, though attention has focused on the possible roles
of these compounds as redox intermediates. The absence of detectable AA in the
QC led to experiments in which roots were treated with AA, which stimulated QC
cells to divide, and resulted in the suggestion that the QC arises, and/or is
maintained, because of a localized depletion of AA
(Kerk and Feldman, 1995
;
Liso et al., 1988
).
A link between polarly transported auxin and AA was proposed based on the
observation that: (1) auxin stimulates the synthesis of ascorbic acid oxidase
(AAO), the enzyme that converts AA from the reduced to the oxidized form; and
(2) that AAO is able to degrade auxin
(Kerk et al., 2000). Taken
together, these two observations formed the basis for the hypothesis that the
QC develops as a consequence of the root's requirement to lower endogenous
auxin levels (Kerk et al.,
2000
). In this scenario polarly transported auxin increases AAO
levels, resulting in a turnover of auxin and a localized depletion of AA, with
a concomitant formation of a QC. Thus, in mutants with perturbed auxin
transport, one would predict that alterations in the distribution of redox
intermediates, such as AA and GSH, would be associated with changes in the QC.
In this paper we explore further the proposed role of redox intermediates in
linking auxin and QC formation.
We make measurements in the QC and adjacent meristem of the distribution of
auxin (indole-3-acetic acid, IAA), of redox intermediates (glutathione,
ascorbic acid and NADPH), and also of the enzymes associated with their
synthesis and metabolism. We focused on these particular intermediates because
they are generally believed to be major players in the control of the redox
status in cells (Noctor and Foyer,
1998). In addition, by using several approaches to perturb polar
auxin transport we were able to alter the redox environment of the QC, thereby
providing insight into the mechanism by which auxin, via redox intermediates,
may mediate the organization of root meristems.
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MATERIALS AND METHODS |
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Ascorbate and glutathione extraction and assay
For ascorbate or glutathione analysis 60-70 QCs (0.42 mg) and 20 PMs (11
mg) were homogenized in 70 µl or 150 µl, respectively, of the
appropriate extraction buffer. Extraction and analysis of ascorbate and
glutathione were carried out essentially following the protocol of Zhang and
Kirkham (Zhang and Kirkham,
1996), except that the total volume of the reactions was 100
µl. Spectrophotmetric measurements were made on a Shimadzu UV160U
spectrophotometer using Eppendorf Uvette microcuvettes. For each assay
standard curves were run simultaneously. To determine whether the extraction
procedure could result in an artifactual loss (oxidation) of reduced ascorbic
acid (AA) or of reduced glutathione (GSH), converting the compounds to their
oxidized forms [dehydroascorbate (DHA) or oxidized glutathione (GSSG),
respectively] we `spiked' a sample of tissue at the beginning of the
homogenization with a known amount of reduced AA or reduced GSH. In general,
we were always able to recover 75-90% of the ascorbic acid or the glutathione
in the reduced forms (data not shown). The values for endogenous AA and GSH
were corrected accordingly. Each extraction was repeated at least three times
with the variation indicated.
Enzyme extraction and assay
The activities of four AA/GSH cycle-associated enzymes were also measured:
ascorbate oxidase (EC 1.10.3.3; AAO), dehydroascorbate reductase (EC 1.8.5.1;
DHAR), ascorbate free radical reductase [EC 1.6.5.4; AFR (monodehydroascorbate
reductase)] and glutathione reductase (ED 1.6.4.2; GR). They were assayed
according to procedures described previously: AAO
(Kerk and Feldman, 1995), DHAR
and AFR (Arrigoni et al., 1997
)
and GR (Zhang and Kirkham,
1996
). The final volumes for each assay were 100-150 µl.
Measurement of reactive oxygen intermediates
Measurements of reactive oxygen species (ROS)
O2·- and H2O2 were carried
out following the protocol of Schopfer et al.
(Schopfer et al., 2001).
H2O2 measurement was accomplished using a Bio-Tek FL600
microplate fluorescence reader, and readings compared to those from a
simultaneously run standard curve. At least three replicates were averaged for
each experiment.
Measurement of NADPH/NADP+
NADPH/NADP+, key regulators of the ascorbic acid/glutathione
cycles, were measured following the protocol of Zhang et al.
(Zhang et al., 2000). For each
experiment 60 QCs and 20 PMs were extracted and a standard curve was prepared
in a range from 0.02 to 0.1 mM NADPH. At least three replicates were run for
each measurement. Using this protocol the limit of detectability is
10-3 mM NADPH.
Immunolocalization of auxin (IAA)
Tissue for immunolocalization and binding of the auxin antibody were
carried out essentially as described previously
(Kerk and Feldman, 1995) using
alkaline phosphatase for detection (anti-mouse IgG AP conjugate; Promega,
Madison, Wisc.). For visualization of the IAA distribution we used NBT/BCIP
tablets for alkaline phosphatase (Roche). The monoclonal antibody used for
this immunolocalization has been shown to be highly specific to free auxin in
Zea root tips (Shi et al.,
1993
), and was used previously on maize roots
(Kerk and Feldman, 1995
).
Controls were again carried out to confirm specificity.
NPA treatment
Roots were treated with NPA (sodium salt of naptalan; N-1-napthylphthalamic
acid; Uniroyal Chemical Company), which is believed to inhibit polar auxin
transport by binding to the auxin efflux carrier
(Nemhauser et al., 2000).
Using intact seedlings (with roots, 2-3 cm in length), the root was inserted
through the center of a 1% agar `collar' (1 cm2 x 3 mm thick)
containing 10-5 M NPA, and the collar positioned at the junction of
the root with the seed. The NPA-treated roots were then returned to a tray
lined with moistened filter paper, with the NPA collar resting on a glass
slide, and not directly in contact with the filter paper. The trays were
tightly covered and the roots returned to darkness. At the end of 24 hours of
NPA treatment the roots' gravity response was visibly perturbed, with roots
orienting randomly (data not shown). For immunolocalization, biochemical and
histological studies, the NPA collar was removed from the root after 24 hours
of treatment (or for some roots, after 48 hours of NPA treatment) and
subsequently these roots used as described more fully in the section detailing
the respective experiments. In order to assess more quantitatively the effects
of NPA on auxin transport, we used roots previously treated for 24 hours with
NPA, then excised the root from the seed (root now about 3 cm in length), and
at the basal cut surface of the root (proximal to the NPA collar) placed a 1%
agar block (approximately 1 mm2) containing 1x10-8
M 5[3H]IAA (specific activity 16.7 Ci/mM; Amersham). Roots with the
attached agar `donor' blocks were returned to a moistened chamber and
incubated for 12 hours in the dark. For control roots we used a plain agar
collar. For most experiments 40 roots were used. Following incubation, the
terminal 1 mm section (the root apex) and basal 1 mm section (the portion of
the root in contact with the donor block) of the roots were excised, pooled,
and then dissolved in 0.5 ml of tissue solubilizer (Amersham) for 15 hours at
room temperature. Three mls of scintillation fluid (ScintiVerse, Fisher
Scientific, Co.) were added to each vial and the samples counted in a
scintillation counter (Beckman).
BrdU Incorporation
BrdU incorporation and detection were carried out as described previously
(Kerk and Feldman, 1995).
Assaying oxidative activity (oxidative stress) in living tissues
Assaying oxidative activity (oxidative stress) in living cells was
accomplished using a dye that is colorless when chemically reduced (when
freshly obtained), but when oxidized, fluoresces green in UV (340 nm
irradiation; 530 nm emission) light. For this work we used carboxy-H2DCFDA
(C-400) dye (Molecular Probes, Eugene Oregon, catalogue no. C-6827). This dye
is oxidized by a wide variety of oxidants, and hence gives a general picture
of the redox environment, rather than indicating the status of a specific
oxidant. At physiological pH this dye has two negative charges facilitating it
passive movement through membranes during loading. Upon oxidation the
fluorescent product is reportedly trapped inside the cell facilitating long
term observation (Collins et al.,
2000; Ha et al.,
1998
; Xie et al.,
1999
). A 10 µM solution of dye was freshly prepared in water
(pH 6.8) just prior to loading into the maize root tissue. In some cases root
tissues had been previously treated with NPA for 24 or 48 hours and then
allowed to grow for an additional 24 hours (no NPA) prior to being exposed to
the dye. In order to facilitate entrance of the dye, root caps were removed
just before loading. Seedlings with decapped roots were placed in Petri dishes
on moistened filter paper and the tips of the roots immersed in 50 µl of
the aqueous dye. Loading typically occurred for a period of 2-3 hours, which
was followed by several washes of the roots in plain water. The QCs and the
adjacent meristem (PM) were then dissected free, placed in water on a
microscope slide, and immediately observed and photographed (in both white and
UV light) with a Leica DM microscope. It was important to photograph the
tissue immediately after exposing to UV light, since prolonged illumination
(in excess of 45 seconds) induced the oxidation of the dye. Although this
limited our observation time, in instances where the tissues initially showed
no fluorescence, this UV-inducible fluorescence allowed us to determine that
dye had indeed entered the cells, but had not been oxidized, under our
experimental conditions.
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RESULTS |
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NPA affects auxin distribution
In order to ascertain the effectiveness of the NPA treatment in perturbing
polar auxin flow, auxin transport was assayed using radiolabelled auxin
([3H]IAA), as previously described
(Kerk et al., 2000). NPA
treatment resulted in a 50% reduction in the amount of radioactivity
transported to the tip (Table
1). A 24-hour NPA treatment resulted in an increase in auxin in
the outer cortical and epidermal regions and a decrease in the amount of auxin
in the columella initials. However, the QC still showed high levels of auxin
(Fig. 2E). After 48 hours of
NPA treatment both the QC and columella initials stained much less intensely
for auxin, but the extent of staining was increased in the proximal cortical
and procambial files (Fig. 2I).
Finally, in roots treated with NPA for 24 hours, and then allowed to grow for
an additional 24 hours without NPA (24+24), the region of most intense
staining included cortical and procambial initials
(Fig. 2M), though this staining
was not as extensive as in roots treated for 48 hours with NPA (compare
Fig. 2I with 2M). In the
24+24-hour treated roots the QC and columella initials showed relatively
little staining (Fig. 2M).
Therefore, at the cellular level, the most pronounced effects of the NPA
treatments are in lowering the levels of auxin in the QC and root cap
initials, and in increasing auxin proximally, in the cells constituting the
typically mitotically active proximal root meristem (PM)
(Fig. 1D,
Fig. 2B). Thus, NPA treatments
lead to a proximal relocation of the auxin maximum. Monitoring of these same
treatment times with BrdU revealed that a 24-hour NPA treatment caused a
diminution in QC size (Fig.
2F,G), but a small QC was still evident. At 24 hours the distal
region of the QC showed less organization, as a consequence of cell
enlargement and a limited amount of cell division
(Fig. 2H). Treating roots for
48 hours with NPA results in an activation of the distal region of the QC
(Fig. 2J,K) and cells of the QC
grew into the root cap (Fig.
2L). However, a small, uncharacteristically elongated QC still
remained (Fig. 2J). In roots
treated with NPA for 24 hours, and then allowed to grow for an additional 24
hours without NPA, the distal end of the QC activated, and showed BrdU
labeling, and this was accompanied by the production of several new (atypical)
layers of cells in the region between the root cap and the tip of the
procambial cylinder (Fig.
2N-P). A small QC was still evident
(Fig. 2N).
|
Root cap excision affects auxin distribution
Excising the root cap causes changes in auxin distribution. Twenty-four
hours after RC removal a large amount of auxin has accumulated throughout the
root tip (Fig. 2Q). Forty-eight
hours after excision a new, rudimentary root cap has formed, indicating that a
new root cap initial layer has formed and is functioning
(Fig. 2S,T). Auxin staining
remains high in the procambial files and is also detected in the columella
region of the regenerating cap (Fig.
2S, arrow). Excision of the root cap also activates the QC.
Twenty-four hours after cap excision most cells of the root tip have
incorporated BrdU and a QC is not detected
(Fig. 2R). By 48 hours after
excision a very small QC is beginning to reform
(Fig. 2T), and by 72 hours a
distinct QC is evident (Fig.
2V).
Ascorbate, glutathione, associated enzymes and NADPH/NADP+
in the QC and PM
Measurements were made of the reduced and oxidized forms of ascorbate and
glutathione in the QC and in adjacent meristem tissue (PM) (Tables
2,
3), and ratios were calculated.
The PM tissue showed about 10x more ascorbate in the reduced form (AA)
than found in the QC, but about 1000x less of the oxidized form (DHA)
compared to the QC. In the QC, there is less AA than DHA (a ratio of 1:18),
compared with a ratio of 300:1 (reduced:oxidized) in the PM. With regard to
glutathione, here too the reduced form (GSH) is about 10x more
concentrated in the PM than in the QC, whereas the oxidized form (GSSG) is
about 10x higher in the QC than in the PM. For the PM, the ratio of
reduced to oxidized is 23:1 and for the QC, 1:8.6. We also measured the
activities of 4 key enzymes involved with regulating ascorbate and glutathione
levels (Table 4). For ascorbic
acid oxidase (AAO) we found approximately 4x as much activity in the QC
as in the adjacent meristem. For glutathione reductase (GR) approximately
9x more activity was found in the PM than in the QC. The activity of
ascorbate free radical reductase (AFR or monodehydroascorbate reductase) was
about the same for both tissues. No dehydroascorbate reductase activity (DHAR)
was detected in either tissue. The concentrations of NADPH and
NADP+ were measured for the PM and were 0.05±0.01 mM and
0.09±0.02 mM, respectively, but neither of these two species was
detected in the QC (limits of assay detectabilty >10-3 mM)
(Table 5).
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O2·- and H2O2
levels in the QC and PM
We also measured the levels of O2·- and
H2O2 since these two species can represent major
reactive oxygen species intermediates (ROS)
(Table 6). While it is not
possible to obtain a specific number for the concentration of
O2·-, following the protocol of Schopfer et al.
(Schopfer et al., 2001) we
find 15.6x more O2·- (on a per mg fresh
weight basis) in the QC than in the PM
(Table 6) (using a 25-minute
incubation). For H2O2 34x more was measured per mg
QC tissue, than for PM tissue.
|
NPA affects levels of ascorbate, glutathione and the activities of
associated enzymes
We also measured the levels and state of ascorbate and associated enzymes
in NPA-treated roots. For the QC, NPA treatment increases 10x the levels
of reduced AA, and decreases by a factor of 4 the levels of the oxidized
species (Table 2), resulting in
a change in the ratio of reduced to oxidized forms, from 1:18 to 2:1. For the
meristem (PM) tissue, NPA causes a marked shift in the amounts and ratios of
AA and DHA. AA decreases about 2.5x, but DHA increases about 50x,
resulting in a change in the ratio of AA to DHA from 300:1 in the untreated,
to 2.3:1 in the NPA-treated roots. Thus NPA treatment results both in a change
in the absolute amounts and in the ratios of the two forms of ascorbate. A
similar trend holds for glutathione; NPA treatments alter the reduced:oxidized
ratios in both the QC and PM (Table
3). NPA treatment also changes the activity of AAO and GR
(Table 4). In general, AAO
activity increases 3-4x in both the QC and meristem. In contrast, GR
activity increases 60x in the QC, while at the same time decreasing
about 3x in the adjacent meristem tissue. Thus, NPA treatments cause the
redox status of the QC to become less oxidizing.
Visualization of the oxidized redox status of the QC
Since the biochemical data indicated that the QC and PM were possibly very
different with regard to their overall redox status, we attempted to visualize
these suggested differences using a redox-sensitive dye, fluorescing in an
oxidizing environment and not fluorescing, or fluorescing less in relatively
less oxidizing environments. We found that the QC was highly fluorescent,
indicating a relatively oxidizing environment
(Fig. 3A,B). This is in
contrast to the adjacent PM tissue, which showed no fluorescence
(Fig. 3C,D) thereby suggesting
that the overall redox status of the PM is relatively less oxidizing than that
in the adjacent QC. These results with the dye are consistent with our
measurements of higher levels of specific reactive oxygen species (e.g.,
O2·- and H2O2) in the QC,
compared to the PM.
|
NPA and decapping change the redox status in the QC
Using the redox-sensitive dye we also monitored the overall redox status of
NPA-treated root meristems (Fig.
4A-J). Twenty-four hours after commencing NPA treatment
fluorescence was noticeably less in QCs
(Fig. 4C,D). Thirty-six to
forty-eight hours after commencing the NPA treatment the extent of
fluorescence was considerably diminished, or nearly undetectable, in QCs
(Fig. 4E-H). Forty-eight to
seventy-two hours after beginning NPA treatment the QC showed reduced or
little fluorescence, whereas the adjacent meristem was highly fluorescent
(Fig. 4I,J). This is the
reverse of what was observed in untreated roots (compare to
Fig. 3C,D). Thus, the
NPA-induced shift in the auxin maximum from the QC to the PM is paralleled by
the development of a more oxidized status in the PM, a more reduced status in
the QC, and the activation of cell division in the QC. Decapping also causes
changes in the redox status of the QC and meristem. Twenty-four hours after RC
removal the redox status of the now activated `QC'
(Fig. 2R) was relatively more
reduced (compare Fig. 5B,D).
Forty-eight hours after RC removal the central portion of the regenerating
root apex showed a small, reforming QC
(Fig. 2T) and a central region
of fluorescence, indicating the development of a region with a relatively more
oxidizing status (Fig. 5F). By
72 hours a new root apex was almost completely reformed and the zone of
fluorescence had enlarged (Fig.
5H).
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DISCUSSION |
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Because the AA:DHA and GSH:GSSG ratios in the QC are skewed in favor of the oxidized forms, this suggests that the QC would be less able to rid itself of ROS and hence would have a relatively more oxidized redox state. Use of a redox-sensitive dye revealed that the QC is, relatively, under more oxidative stress than the adjacent PM cells (Fig. 3). Although we cannot precisely quantify the degree of oxidative stress in the QC, compared to the PM, it is clear from the measurements of ROS (O2·- and H2O2), that there are differences in the oxidative intermediates in these two tissues (15.6x more O2·- and 34x more H2O2 were found in the QC, compared to the PM) (Table 6). Taken together, these data point to a general inhibition in the QC of mechanisms to generate and maintain a reduced redox status.
Activating the QC correlates with changes in the redox status
Data thus far presented suggest that the QC is under oxidative stress. We
therefore reasoned that if cells in the QC can be stimulated to divide, QC
activation should be associated with changes in the redox status of the QC.
Excision of the cap or treatment of roots with NPA activates the QC. Excision
of the cap both alters QC redox status
(Fig. 5D) and causes the QC to
disappear (Fig. 2R).
Reappearance of a QC in decapped roots
(Fig. 2T) correlates with the
redevelopment of an oxidized state in cells of the presumptive QC
(Fig. 5F). As with root cap
excision, NPA treatments not only activate the QC but also result in changes
in the redox status of the QC (Fig.
2F,J,N; Fig. 4;
Tables
2,3,4).
Twenty-four-hour NPA-treated roots have a small QC (compare
(Fig. 2B and 2F) and also, the
zone of fluorescence (indicating a relatively oxidizing status) is diminished
(Fig. 4D). After 48 hours of
NPA treatment, when the QC is maximally activated
(Fig. 2J,K), no fluorescence is
detected (Fig. 4G,H), thereby
indicating a relatively more reduced redox status. Interestingly, after 48
hours of NPA treatment, it is the PM that now fluoresces
(Fig. 4I,J). Thus, as with root
cap removal, NPA treatments cause both an activation of the QC and a changed
redox status in both the QC and PM. Taken together, these results demonstrate
that not only is an oxidized redox state characteristic of the QC, but
moreover, that activation of the QC correlates with the development of a less
oxidized status in the QC, and the development of a new redox status (more
oxidized) in the adjacent PM. Given that mild oxidative stress has been shown
to impair the G1/S transition in plants
(Chen and Ames, 1994;
Bijur et al., 1999
;
Chen et al., 1995
;
Logemann et al., 1995
;
Reichheld et al., 1999
) and in
animals (Russo et al., 1995
)
and that redox status can be a modulator of the balance between renewal and
differentiation in animal cells (Smith et
al., 2000
), we conclude that the lengthened cell cycle times in
the QC are also likely a consequence of the relatively oxidized redox status
of the QC. This correlation between redox status and the degree of quiescence
further supports the suggestion that "intracellular redox homeostatis
could affect cell-cycle progression by regulating key components of the
G1/S transition" (den Boer
and Murray, 2000
), and leads us to conclude that the establishment
and maintenance of an oxidizing environment may be central to the development
and elaboration of a QC.
Auxin regulates the QC via redox
Excision of the cap or treatment of roots with NPA not only causes changes
in the redox status of the QC and activation of the QC, but also brings about
changes in auxin levels and distribution in the root tip. These observations
suggest a linkage between auxin distribution, QC formation and redox status at
the root tip. In an untreated, control maize root the auxin maximum is located
in the root cap columella initials/QC (Fig.
2C), mirroring very closely the auxin patterns observed in
Arabidopsis roots (Friml et al.,
2002; Sabatini et al.,
1999
). We show that NPA causes a shift in the auxin maximum from
the columella/QC zone to the region of the PM (compare
Fig. 2C and 2I), thereby
confirming in maize what has earlier been reported for the effects of NPA on
the position of the auxin maximum in Arabidopsis
(Sabatini et al., 1999
). But
in addition, we extend this work and show that a shift in the auxin maximum
correlates with a change in the redox status, which becomes more oxidizing in
the region to which the auxin maximum has been shifted (the PM), and less
oxidizing in the region from which the auxin maximum has been displaced (the
QC) (Fig. 4G,H). Of particular
note is the observation that shifting the auxin maximum to the PM results in a
decrease in BrdU incorporation in the PM (compare
Fig. 2B and 2J), indicating a
decrease in the rate of cell division in the PM. Taken together these results
provide strong evidence that QC activation and changes in the redox status are
associated with a shift of the auxin maximum.
As with the NPA treatments, excising the cap leads to marked changes in
auxin distribution in the remaining root tissues. Of particular significance
is the appearance of auxin staining (development of a new auxin maximum) in
the columella region of the new, regenerating root cap
(Fig. 2S; arrow). While it is
not now certain whether the increase in auxin in this region occurs before the
start of reforming a new QC, it is clear that the development of a new auxin
maximum in the regenerating cap occurs at least at the very earliest stages of
QC reformation (Fig. 2S,T).
These results thus imply a role for the root cap initial layer/columella (and
perhaps the whole cap) in the development and maintenance of the QC, and
further suggest that positioning of the auxin maximum in the (new) root cap
may be central to QC redevelopment. Support for an hypothesized role for the
auxin maximum in `specifying' the QC comes from Arabidopsis that was
treated for extended periods with NPA; a shift of the auxin maximum to new,
more proximally located cells is accompanied by the atypical expression of
QC-specific markers in these cells
(Sabatini et al., 1999).
Auxin is widely involved in patterning
(Friml et al., 2002;
Sabatini et al., 1999
), acting
as a positional signal (Uggla et al.,
1996
). Based on our findings that location of the auxin maximum
correlates with oxidative stress in the QC, we suggest that auxin can provide
positional cues by virtue of its ability to influence, on a localized scale,
the redox status of tissues. The challenge now is to elucidate the
hypothesized link between auxin and ROS formation.
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ACKNOWLEDGMENTS |
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