Development 121, 2825-2833 (1995)
Printed in Great Britain © The Company of Biologists Limited 1995
2825
A biochemical model for the initiation and maintenance of the quiescent
center: implications for organization of root meristems
Nancy M. Kerk* and Lewis J. Feldman
Department of Plant Biology, 111 Koshland Hall, University of California, Berkeley, CA 94720-3102, USA
*Author for correspondence (e-mail: kerk@nature.berkeley.edu)
SUMMARY
A new hypothesis for the formation of the quiescent center
is presented. Reported data support a mechanism for the
establishment and maintenance of the quiescent center.
The quiescent center is located at the most distal part of
the root proper, the most terminal location in the root
proper on the path of polar transport from the shoot. Of
the many substances polarly transported in the root, auxin
is one of the best studied and has been shown to affect root
meristem organization. In our mechanism, polar auxin is
directly linked to quiescence through the action of
ascorbate oxidase and ascorbic acid. Immunolocalization
of auxin in the root tip of Zea mays showed that auxin levels
in the quiescent center were high compared to the levels in
the immediately surrounding meristematic cells. Isolated
quiescent centers were shown to have high levels of
ascorbate oxidase mRNA and ascorbate oxidase activity
relative to proximal meristem tissue. Exogenous auxin
caused an increase in ascorbate oxidase mRNA levels and
ascorbate oxidase enzyme activity in cultured root tissue.
Immunolocalization of ascorbate oxidase in Zea root tips
showed high levels of the protein in the quiescent center
relative to surrounding cells. This is the first report of a
positive marker and activity for the quiescent center. Histochemical detection of ascorbic acid in Zea root tips
showed that quiescent center cells have low or undetectable
levels of ascorbic acid, presumably due to the high levels of
ascorbate oxidase in the quiescent center. As ascorbic acid
is a compound known to be necessary for the transition
from G1 to S in the cell cycle, its low levels in the quiescent
center may be directly responsible for holding these rarely
dividing cells in the extended G1 state in which they are
mainly found. We propose that our mechanism complements published mathematical modeling of the anatomical
structure of root apices, and further propose that the
control of relative growth rates in this focal region of the
root apex by this mechanism is a determining aspect in generating anatomical patterning in the root apex.
Key words: quiescent center, auxin, ascorbic acid, Zea mays, root
development
INTRODUCTION
Apical meristems give rise to all tissue and organ systems of
the postembryonic plant. Not only do they generate the cells
from which the plant is constructed, but apical meristems also
function as the organizing centers for postembryonic morphogenesis. The evidence for this conclusion has emerged
gradually from many different studies addressing the initiation,
organization, maintenance and function of apical meristems,
especially root apical meristems (Steeves and Sussex, 1989).
The organization of root meristems has been studied through
analyses of cell lineages, based on histological sections.
Because cell lineages, or files, converge at the root pole,
Hanstein (1868) postulated that a few cells, located at the pole
could serve as the initials for the root, and through high mitotic
activity, generate all the cells that make up the root. In an effort
to define these cells precisely, Clowes (1953, 1954) performed
surgical experiments on Zea mays root apices and showed that
if these cells were damaged or removed, the remaining surrounding cells could directly reconstitute a complete apex. This
led Clowes to conclude that the functional initials were actually
located peripherally to the very central cells, in a region of the
meristem later named the proximal meristem (Feldman and
Torrey, 1975). Clowes’s (1956) use of thymidine labeling
showed for the first time that the most central cells of the
meristem actually divide infrequently, or not at all, and he
named this population of cells the quiescent center (QC).
Since its discovery, much work has contributed to the characterization of the QC. While it is believed to be a feature of
all angiosperm root apices, the most extensive analysis of the
QC has been done on roots of maize, in which the QC can
attain a size of 1000-1500 cells (Feldman and Torrey, 1976).
Dolan et al. (1993) have shown that in Arabidopsis the QC
comprises only four central cells derived from the hypophysis
and is surrounded by cells that act as the initials for the files
of cells that make up the root. Average cell cycle times within
the QC of Zea are in the range of 170 hours. In contrast, the
cell cycle time of the root cap initials, the most rapidly dividing
cells in the root, is only 10-16 hours, and for the proximal
meristem, 18-25 hours (Clowes, 1961).
Cells within the QC can be distinguished from surrounding
2826 N. M. Kerk and L. J. Feldman
meristem cells by their fainter histochemical staining (Goyal
and Pillai, 1986), lower RNA content (Clowes, 1972), lower
protein content (Jensen, 1958), and low RNA polymerase
activity (Fisher, 1968). In situ hybridizations with a 3H-labeled
histone gene probe and with [3H]polyuridylic acid (poly U) to
root tissue sections have revealed populations of unlabeled
cells which correspond precisely to the area defined as the
developing QC in rice and Capsella for each probe respectively
(Raghavan and Olmedilla, 1989; Raghavan, 1990).
Prior to discernible apical organization, cells at the presumptive root pole (the future root meristem) show uniformly
high rates of mitosis, and stain densely with a variety of histochemical stains. Establishment and elaboration of a QC
occurs gradually and precedes stable histological patterning in
the meristem. From experiments using uptake of radiolabelled
precursors, Clowes (1978) concluded that establishment of the
QC preceded all histogenic events except establishment of a
root cap meristem layer. More recently, using in situ hybridization of radiolabelled poly U to Capsella, Raghavan (1990)
was able to show that very early during embryogenesis, before
any root apical organization was evident, one cell and subsequently several were clearly distinguished by their relative lack
of bound probe. He then demonstrated that these unlabelled
cells were the origin of the QC. His results, in an elegant way,
supported previous hypotheses that the organization of apical
meristems in roots follows the establishment and elaboration
of the QC. Other work concerning the establishment of histological patterning in developing lateral root primordia and in
adventitious root primordia has also provided evidence for the
establishment of a QC prior to histological organization of a
root meristem (Rondet, 1961; Feldman, 1977).
Using microsurgical techniques, earlier workers had shown
that it was possible to excise defined regions of the root apical
meristem in maize, including the QC itself (Feldman, 1977).
Results of these efforts showed that roots are able to regenerate a new apical meristem following surgical excisions, but that
prior to the initiation of distinctive histological zonation, a QC
reformed, appearing initially as a group of mitotically relatively inactive cells surrounded by the rapidly dividing cells
remaining at the cut root stump (Feldman, 1977; Rost and
Jones, 1988). From this work it was also concluded that the
reformation of a QC precedes and is requisite for organization
of root meristems.
Despite these many studies we do not have definitive information about the factors that initiate and maintain quiescence
in these cells nor do we have a convincing functional role for
these cells. Past workers have suggested that the QC and the
‘ultimate initials’ that it contains are equivalent to ‘stem’ cells
in animals (Barlow, 1978). Torrey (1972), and later Feldman
(1975, 1979), proposed that the QC may be a site of hormone
biosynthesis in the root and as a consequence of these
localized, enhanced metabolic processes these cells were
inhibited with regard to many other physiological activities.
Other possible explanations for the quiescent state have
focused either on the supposed nutritional status of the QC or
on the possibility that physical constraints prevent cells of the
QC from dividing (Clowes, 1972).
In this paper we present a new hypothesis for the formation
of the quiescent center and provide data that address the cause
and maintenance of the QC. We have used the perspective of
examining the QC with regard to its position in a whole plant
context. The QC is located at the most distal part of the root
proper, the most terminal location on the path of polar transport
from the shoot. Of the many substances polarly transported in
the plant, auxin is one of the best studied and has been shown
to affect root meristem organization. Here we provide a
detailed mechanism linking polar auxin transport with the
establishment and maintenance of the QC (Fig. 1). In this
mechanism polar auxin is directly linked to quiescence through
the action of ascorbate oxidase and ascorbic acid. Briefly, we
report that auxin and ascorbate oxidase levels are high in the
QC relative to surrounding cells and that the QC cells have low
or undetectable levels of ascorbic acid, a compound known to
be necessary for the transition from G1 to S in the cell cycle.
Having discussed the mechanism imposing quiescence we
discuss the implications that this mechanism has for the establishment of pattern at the root apex.
MATERIALS AND METHODS
Plant growth conditions and tissue collection
Corn caryopses (var. Merit, Asgrow Seed Co., Kalamazoo, MI) were
imbibed and germinated in the dark at 25˚C for 2 days. Tissue was
collected by surgical removal of the cap and excision of the QC,
(Feldman and Torrey, 1976). In this cultivar, the QC is separated from
the proximal meristem by a weak, thin-walled junction making
possible routine, clean dissections of isolated QCs. QCs were
collected in a moist environment, quick frozen on dry ice and stored
at −80˚C for extractions as described below. Approximately 2 mm of
the remaining root stump, the proximal meristem region, was also
collected and stored in this manner. Where mature root tissue was
used, 1 cm segments located approximately 1 cm behind the tip were
collected. For root tissue cultured with or without exogenous auxin,
mature root segments (1 cm) were cultured on Murashige and Skoog
(MS) medium, 3% sucrose, 0.8% agar in the presence of 0 or 1.0 mg/l
IAA
AAO
Polar IAA
AA
Fig. 1. Autoradiograph of a median longitudinal section of a maize
root that had incorporated [3H]thymidine. The silver grain deposits in
the darkly colored nuclei indicate those cells that were undergoing
DNA synthesis during the labeling period. Note the prominent
quiescent center at the apex. Superimposed on the section is an
overlay indicating relative amounts of the elements of the proposed
mechanism for maintenance of the quiescent center at locations in
the meristematic and quiescent regions in the root tip. Auxin and
ascorbate oxidase are at relatively high levels in the quiescent center
while levels of ascorbic acid are relatively low. (IAA, auxin; AAO,
ascorbate oxidase; AA, ascorbic acid). Magnification, ×110.
Maintenance of the quiescent center 2827
of 2,4-D. Material was incubated in the dark at 25˚C and was
harvested and collected as above after 2 and 4 days (Esaka et al.,
1992).
(Oberbacher and Vines 1963). Protein concentration was determined
using the Bio-Rad protein assay and units of activity were normalized
to this amount.
Indoleacetic acid transport
Caryopses were germinated as above and grown for 3 days. Roots were
severed at the base, below the scutellar node and the cut end was placed
in a 0.5 ml microfuge tube containing half-strength MS medium,
10−8 M nonradioactive indoleacetic acid (IAA) and [14C]IAA (1
µCi/ml, specific activity = 4.8 Ci/mM) in 0.8% low melting temperature agarose (Feldman, 1981). After 24 hours of transport at room temperature in the dark, roots were removed from tubes and squashed
between plastic wrap and Whatman blotting paper. A glass plate was
placed over the plastic wrap and pressure was applied squashing the
root firmly and uniformly onto the paper. Squashed roots were exposed
to X-ray film for approximately 2 weeks.
Ascorbic acid localization
This procedure uses the unique ability of ascorbic acid to reduce silver
nitrate to silver in acidic conditions (Chayen, 1953). Root tips were
quick-frozen in isopentane, cooled in a dry ice bath and dehydrated
in several changes of absolute ethanol at −30˚C. The ethanol was
replaced with toluene and the tissue infiltrated with paraffin and
sectioned at 10 µm (Jensen, 1962). Ascorbic acid was localized
according to the methods of Jensen and Kavaljian (1956). Black
deposits of metallic silver indicate regions where ascorbic acid is
present. The control for this procedure is to expose the sectioned
material to a copper sulfate solution that oxidizes all the ascorbic acid
to dehydroascorbic acid, which does not react with the silver nitrate.
RNA isolation, northern blotting and in situ hybridization
RNA was isolated from frozen tissues collected as described above.
RNA was isolated from material using a modification of the method
of Puissant and Houdebine (1990). Approximately 150 individual
frozen QCs were ground in a small grinding tube in 25 µl guanidinium
buffer on ice. 2.5 µl 2 M sodium acetate, pH 4.0, and 25 µl water
saturated phenol/chloroform was added. The solution was transferred
to a microfuge tube, vortexed and centrifuged at 12,000 g for 10
minutes at 4˚C. The upper phase was recovered and precipitated with
isopropanol. The pellet was resuspended in 25 µl 4 M LiCl, then centrifuged at 3000 g for 10 minutes at 4˚C. The resulting pellet was
redissolved in Dep-treated H20, 0.1% SDS, extracted with an equal
volume of chloroform, and the aqueous phase was adjusted to 0.2 M
sodium acetate, pH 5.0, and precipitated again with isopropanol. The
resulting pellet was resuspended in Dep-treated H2O, and RNA was
quantitated and used for northern blot analysis. RNA was extracted
from proximal meristems and other root sources by the same method
except the volumes were scaled up to process the tissues which were
more easily collected in larger amounts.
RNA electrophoresis, blotting and hybridization were performed
essentially as described previously (Maniatis et al., 1989). 5 or 10 µg
of total RNA was electrophoresed in a 1% agarose gel containing
formaldehyde and blotted to Nytran (Schleicher & Schuell). Equal
loading of RNA was confirmed by ethidium bromide staining of the
gel before transfer to the membrane. RNAs were probed with the near
full length cDNA clone for cucumber AAO, pASO11 (Ohkawa et al.,
1989). The probe was radiolabeled using random hexamer priming
with the Prime-a-Gene method (Promega).
In situ hybridizations were done according to the method of
Jackson (1991). Maize root tips were hybridized with 35S-labeled
sense and antisense riboprobes (synthesized with Ribo Probe kit,
Stratagene) coding for elongation factor-α cloned from radish (Kerk,
1990). Slides were hybridized overnight, washed in 2× SSC, 50%
formamide at 50˚C, dried and exposed to Kodak NTB-2 emulsion.
Immunolocalization
Tissue for immunolocalization was fixed in freshly prepared 2%
aqueous ethyl-3-(-3-dimethylaminopropyl) carbodiimide hydrochloride (EDC) (Sigma) on ice for 30 minutes under vacuum (Shi et al.,
1993), followed by postfixation in 2.5% paraformaldehyde/0.25%
glutaraldehyde in 0.1 M phosphate buffer, pH 7.0, at 4˚C overnight.
Binding of antibody (Ab) was carried out essentially as described by
Chichiricco et al. (1989) using alkaline phosphatase for detection. The
monoclonal antibody (mAb) to auxin has been shown to be specific
to free auxin in Zea root tips (Shi et al., 1993). Several controls were
carried out to confirm specificity. The controls included: (1) omitting
prefixation with EDC, (2) omitting incubation with the primary mAb
and (3) incubation with the mAb previously exposed to an excess of
auxin in incubation solution. The AAO antibody was a polyclonal Ab
(Esaka et al., 1988) and was used for localizations in tissue sections
as above, with the omission of the prefixation in EDC.
Assay of ascorbic acid oxidase activity
Ascorbic acid oxidase (AAO) activity was assayed by following the
decrease in the spectrophotometric absorbance of ascorbic acid over
time in the presence of proteins extracted from QCs, proximal
meristems, and other root tissues using the method of Oberbacher and
Vines (1963). Tissue homogenates were prepared from freshly
collected quiescent centers, proximal meristems and cultured root
segments. Tissues were ground in 5 parts (w/v) 0.1 M potassium
phosphate buffer, pH 7.0, on ice and centrifuged at 10,000 g for 15
minutes at 4˚C. 10 µl samples of these homogenates were added to
the reaction mixture containing 0.05 M potassium phosphate buffer
(pH 7.0), 0.5 mM EDTA, and 0.15 mM L-ascorbic acid in a volume
of 1.0 ml (Esaka et al., 1988). One unit of activity was defined as the
amount of enzyme which oxidizes 1.0 µmol of L-ascorbic acid per
minute and was converted from the change in A265 at 25˚C over time
BrdU incorporation
Corn caryopses were imbibed and germinated for 3 days as described
above. Caryopses were then placed upon parafilm covered deep Petri
dishes such that the root extended through holes punched in the film
into solution contained in the dish. Solutions were either water or 0.1
mM ascorbic acid both maintained at pH 5.9. Roots were incubated
at 25˚C in the dark for 24 hours with gentle agitation. Bromodeoxyuridine (BrdU) was added to 10 µM and roots were incubated
for a further 24 hours. Root tips were then excised, fixed and sectioned
as described above.
Hydrolysis and immunofluorescent staining
A modification of the procedure recommended by Boehringer
Mannheim for use with their Anti-BrdU antibody was used. Sections
were hydrolyzed in 1 N HCl for 1.5 hours at 37˚C, then neutralized by
immersion in 0.1 M borate buffer, pH 8.5, washed with PBS, and
incubated with anti-BrdU antibody (Developmental Studies
Hybridoma Bank, NICHD) for 2 hours. Slides were washed in PBS
and incubated with rabbit anti-mouse IgG-FITC (Southern Biotechnology Associates, Inc.) overnight in the dark at room temperature in
a humidified chamber. Slides were washed and covered with a drop of
50% glycerol, 0.15% N-propyl gallate in PBS and a coverslip. The
stained material was observed with a Zeiss Axiophot microscope
equipped with a Zeiss ZVS-47DEC video camera. Video frames from
the ZVS-47DEC were digitized and displayed on a Macintosh Power
Mac 8100/80AV and arranged using Adobe Photoshop v. 3.0 software.
RESULTS
Localization of auxin
We established that auxin accumulates in the region of the QC
2828 N. M. Kerk and L. J. Feldman
Fig. 2. Autoradiograph of a primary maize seedling root after 24
hours of incubating the basal end with [14C]IAA. Arrow shows the
accumulation of IAA in the root tip. Also note the distribution of
label in the vascular tissue and developing lateral root primordia
(small arrowheads). Bar, 1 cm.
by using the following two methods: (1) by demonstrating
regions of accumulation of polarly transported [14C]IAA using
autoradiography and (2) immunolocalization of IAA to root tip
tissue sections. The first approach allowed visualization of the
path of polar IAA transport in 3-day old maize seedling roots
using [14C]IAA. Roots were severed from the hypocotyl and
exposed to [14C]IAA at the cut surface. Transport was allowed
to proceed for 24 hours and the roots were processed and
exposed to X-ray film. Fig. 2 shows a representative root.
Radioactivity was localized in the vascular tissue and in the
root tip. A characteristic feature was the low level of signal in
the region behind the tip, the region of cell elongation. Signal
can also be seen in vascular traces leading to developing lateral
root primordia.
Higher resolution of auxin localization in the root tip was
obtained using a monoclonal antibody to auxin. This antibody
has previously been shown to have high specificity for free
auxin (Shi et al., 1993). Fig. 3A shows the alkaline phosphatase staining pattern indicating antibody binding to auxin.
There was distinctive dark staining in the region corresponding to the quiescent center. The root cap also showed high
levels of auxin, as did the outer cortical cells of the root
proper. The vascular tissue showed significant staining as
well. The root cap meristem region between the QC and the
cap had much less auxin as did the inner cortex and epidermis.
Two controls are also shown. First was the pattern seen when
tissue sections were treated as above but without the primary
antibody in the dilution buffer (Fig. 3B). There was little
detectable staining in the QC. The other control shows the
result of incubating the antibody with an excess of auxin in
solution prior to exposure of the antibody to the tissue section
(Fig. 3C). With this control, sites antigenic for auxin should
become saturated prior to exposure to auxin in the tissue
sections, and hence should not be able to bind auxin during
immunolocalization. This pattern showed some generalized
background staining but the marked differential distribution
seen with the antibody alone was not detectable. The staining
in the quiescent center, root cap, outer cortical cells and
central cylinder was very reduced when the antibody was pretreated with auxin.
Effect of auxin on ascorbate oxidase levels
Auxin has an effect on AAO levels in the root. AAO activity
Fig. 3. Auxin localization in longitudinal sections of maize root apices. (A) Section incubated with monoclonal antibody to auxin. Note dark
staining in region of the quiescent center. In addition, staining is intense in the root cap and outer cortical files and in maturing vascular
elements. B and C are controls. B treated as A but without incubation with the primary antibody; C as A but antibody pretreated with a molar
excess of auxin before incubation with tissue sections. Magnification ×90.
Maintenance of the quiescent center 2829
Fig. 5. Expression of ascorbate oxidase and p34cdc2 in various
root tissues (Q, quiescent center; P, proximal meristem; R, mature
root tissue). (A) Ethidium bromide stained gel to show RNA
loadings of the blotted gel (M = molecular mass (×10−3) markers).
(B) Northern blot of gel hybridized with ascorbate oxidase cDNA
probe. (C) Same filter as in B, hybridized with p34cdc2 cDNA
probe.
Fig. 4. Effects of auxin (2,4-D) on ascorbate oxidase activity and
expression in cultured roots. (A) Ascorbate oxidase activity after
culture with (m) or without (v)2,4-D. (B) Northern blot of total
mRNA from root tissues cultured in the presence, +, or absence, −,
of 2,4-D after 0, 2, and 4 days of culture. Note message level is
highest after 2 days of culture with auxin.
proximal meristem, making it possible to remove the root cap
and collect individual QCs. Northern blots of total RNA
probed with a full-length cDNA for AAO show high levels of
mRNA in the QC and lower levels in the proximal meristem
and mature root region (an ethidium bromide stained gel is
shown as a control for equal loading of the blotted gel, Fig.
5A,B). The same filter was reprobed with a cDNA for a cell
increases in tissue cultured with auxin.
Roots cultured with auxin for 4 days
showed a ten-fold increase in activity
compared to control roots (Fig. 4A).
Northern blots of RNA prepared from
portions of the same root tissue as that
used for the activity assays show
message levels of AAO peak at day 2
and decrease by day 4 (Fig. 4B). Esaka
et al. (1992) also observed this same
mRNA profile in pumpkin fruit tissue
cultured with auxin. These data provide
evidence that culture with auxin results
in an increased level of AAO mRNA
and enzyme activity in root tissue, and
that a continued increase in the mRNA
level does not underlie the increasing
levels of enzyme activity, since AAO
activity levels continue to increase even
though mRNA levels decrease before
day 4.
Ascorbate oxidase localization in
the quiescent center
The following experiments examined
AAO mRNA and protein distribution,
and AAO activity in the different regions
of the root. The cultivar of maize used for
this work has a slightly weakened cell
wall zone between the QC region and the
Fig. 6. Characterization of the quiescent center region of maize root apices.
(A) Immunolocalization of ascorbate oxidase in the root apex. Note the dark staining in the
quiescent center. (B) In situ hybridization of elongation factor-α to the root apex. Notice the
probe does not bind strongly to the region of the quiescent center, and in the root cap, binds
only to the root cap meristem. Magnification, ×90.
2830 N. M. Kerk and L. J. Feldman
in the QC, root cap, central vascular cylinder and outer cortical
cells, AAO protein is relatively high in these tissues. The AAO
pattern is also very similar to the pattern of auxin distribution
previously shown with the auxin antibody (compare with Fig.
3A). Spectrophotometric assays of AAO activity in protein
extracts made from QCs and proximal meristems show that this
increased level of AAO protein in the QC corresponds to much
higher activity levels of AAO in the QC (Fig. 7). Thus we have
shown that the QC has high levels of AAO mRNA, AAO
protein and AAO activity, compared to the immediately surrounding cells of the proximal and root cap meristems.
Units of AAO/mg of total protein
0.06
0.05
0.04
0.03
0.02
0.01
0.00
Proximal Quiescent
Meristem Center
Fig. 7. Ascorbate oxidase activity in two regions of the root tip.
cycle gene, p34cdc2, (Colasanti, J. et al., 1988) to demonstrate
the different levels of these two messages in the total mRNA
extracted from these tissues (Fig. 5C). No detectable mRNA
was present in the QC, while high levels were detected in the
proximal meristem where a high rate of cell division occurs.
There is lower signal in the mature root as would be expected
in tissues with fewer dividing cells.
The distribution of AAO protein in these regions of the tip
is similar to the pattern of mRNA distribution. AAO is
localized very distinctly in the QC but occurs at much lower
levels in the proximal meristem (Fig. 6A). The root cap
meristem region has relatively low levels of AAO protein.
When this pattern is compared to the in situ hybridization
pattern of elongation factor-α, a translation factor that has been
shown to be a marker for cells undergoing active protein
synthesis, the two patterns can be seen as the inverse of each
other (Fig. 6B). While elongation factor-α is at very low levels
Fig. 8. Histochemical localization of ascorbic
acid in the root tip of maize. (A) Region of the
proximal meristem; note black dots of silver
denoting ascorbic acid and the mitotic figure
indicating this is a region of high mitotic
activity. (B) The control for A; notice mitotic
figures but the absence of black dots in the
section. (C) Section from the same root as A,
but from the region of the quiescent center.
Note the absence of black dots and mitotic
figures. (D) Low power view of a longitudinal
section indicating the regions from which A
and C were photographed (QC, quiescent
center; C, mitotically active cortex).
Magnification, (A-C) ×450; (D) ×80.
A
Ascorbic acid localization
The primary substrate for AAO is ascorbic acid, which is
utilized in several metabolic processes, and has been reported
to be necessary for the transition from G1 to S in the cell cycle
of several plants (Chinoy, 1984). Others have also suggested
that ascorbic acid may have a regulatory role in cell proliferation (Innocenti et al., 1989; Arrigoni et al., 1989; Liso et al.,
1984; Citterio et al., 1994). Moreover, it is known that cells in
the quiescent center have extended G1 phases and divide rarely
(Clowes, 1975). Localizations of ascorbic acid show it to be
absent or present at undetectable levels in quiescent center cells
(Fig. 8). At high magnification, the proximal meristem region
shows orthogonal cell files with a clearly visible mitotic figure.
The silver deposits that indicate the presence of ascorbic acid
in these cells appear as black dots. Cells in the QC region are
much less regularly shaped, show no mitotic figures and
contain no silver deposits.
Effect of ascorbic acid on the quiescent center
Roots incubated for 48 hours in 0.1 mM ascorbic acid showed
increased levels of cell division activity compared to roots
incubated in water. Most striking was the activation of all cells
in the quiescent center as viewed in median sections through
the root apex (Fig. 9). The immunofluorescent signal over the
B
C
QC
C
D
Maintenance of the quiescent center 2831
nuclei indicates cells that had incorporated BrdU during
nuclear DNA replication and thus marked those that had passed
from G1 through S of the cell cycle during the period of the
experimental treatment. The control roots in water showed a
prominent quiescent center and low levels of signal in the area
of the stele. The root cap meristem was evident and appeared
to be composed of two clearly labeled cell tiers. In contrast,
the ascorbic acid-treated roots had an overall higher level of
BrdU incorporation, which is reflected not only by the brighter
fluorescence of the majority of nuclei, but also by enhanced
cytoplasmic fluorescence, perhaps indicative of increased rates
of organelle DNA replication (Fujie et al., 1993). No QC could
be distinguished, but just distal to the root cap juncture, the
root cap meristem showed at least 4 prominent tiers of dividing
cells. Thus, applying ascorbic acid directly to roots activates
cell division in the majority if not all quiescent center cells.
In summary, the QC has relatively higher levels of auxin,
AAO mRNA, AAO protein, and AAO activity, and lower or
undetectable levels of ascorbic acid compared to the more
rapidly dividing cells surrounding it. In addition, the general
pattern of auxin distribution in the root tip is coincident with
the pattern of AAO protein localization; both of which appear
to be the inverse of the pattern of elongation factor-α, which
has been used as an indicator of high levels protein synthesis
and as a negative delineator of the quiescent center (Kerk,
1990).
DISCUSSION
Establishment and maintenance of the quiescent
center
We have proposed a new model for the establishment and
maintenance of the quiescent center that is derived from consideration of its position and possible function in a whole plant
context. As a terminal region with regard to the transport of
many substances, the QC is located in a potentially dynamic
region of the plant root. Recent studies of phloem transport and
unloading in Arabidopsis roots have shown that fluorescent
dye tracers can accumulate in the quiescent center after phloem
unloading in the elongation zone and symplastic transport to
the tip (Oparka et al., 1994). Our studies of transport of
[14C]IAA in maize seedling roots showed accumulation of
radioactivity at the root tip and also in the region of cell
elongation. This latter area corresponds anatomically to the
region of phloem unloading in Arabidopsis roots.
Antibody localization of auxin revealed that the root cap and
quiescent center contained relatively higher levels of IAA than
the immediately surrounding proximal meristem and root cap
meristem cells. The vascular tissue, pericycle and outer cortical
cells also showed high levels of antibody binding and these
tissues also correlate anatomically with the regions in which
Oparka et al. (1994) observed phloem transport and unloading.
Salamatova (1993) has also reported similar patterns of auxin
distribution in Monstera roots, using color reagents.
The effects on growth and development of the balance of
hormone levels in plant tissues is well established. Localized
accumulation of a hormone in a small population of cells could
cause them to have significantly different metabolic properties
than the cells surrounding them. Indeed, other investigators of
the quiescent center have hypothesized that the hormone
balance in the quiescent center may be different from that in
adjacent regions, but speculations as to the exact difference
have varied widely.
Many auxin responsive genes have now been identified and
promoters for some of these have been shown to drive transcription of reporter genes in response to auxin exposure. Here
we report that exogenous auxin can increase the level of AAO
mRNA in root segments and that the mRNA level of AAO is
significantly higher in quiescent center cells than in proximal
meristem cells or mature cells of the root. Auxin was previously shown to increase levels of ascorbate oxidase activity in
Fig. 9. Cell division activity shown in median longitudinal sections of maize root apices supplied with 10 µM BrdU for the last 24 hours of a 48
hour experimental treatment. Immunofluorescence indicates nuclei that had incorporated BrdU during DNA synthesis. (A) Control root kept in
water. Note the prominent quiescent center. (B) Root was treated with 0.1 mM ascorbic acid for 48 hours. No quiescent center is evident.
Magnification, ×140.
2832 N. M. Kerk and L. J. Feldman
tobacco pith tissue and cultured pumpkin fruit (Newcomb,
1951; Esaka et al., 1992) and the present results show the same
effect in segments of maize root tissue. It would be of interest
now to determine the mechanism for these higher levels of
AAO in the quiescent center, and to determine if this gene has
an auxin-inducible or auxin-sensitive promoter.
AAO is a plant-specific copper-binding protein that has been
localized mainly to the cell wall but there are reports of localization to other cellular compartments in many different cell
types. It is very effective at oxidizing ascorbic acid, a
compound necessary for many metabolic reactions and for the
transition from G1 to S in the cell cycle. We have shown high
levels of AAO protein and AAO activity in QC cells relative
to proximal meristem cells, and we suggest this causes the lack
of detectable ascorbic acid in the QC, and as a consequence,
the reduced mitotic activity there. These results are consistent
with other reports of the involvement of ascorbic acid in the
progression of G1 phases in the cell cycle as well as in other
aspects of plant cell growth (Cordoba et al., 1994). Moreover,
as a redox reaction, the oxidation and ratio of ascorbic acid to
dehydroascorbic acid in regions of the root suggests that the
ascorbate system could be part of a larger redox regulatory
system. There is a growing body of data that show redox
potential can influence the regulation of gene expression and
that modulation of the redox potential in cells is indeed an
important regulatory system for cellular functions (Allen,
1993; Crane, 1994). When root tips are cultured in the presence
of ascorbic acid, cells in the QC are activated to divide, and
when root tips are treated with an inhibitor of ascorbic acid
biosynthesis, cell division is inhibited and cells arrest in G1
(Liso et al., 1984, 1988). Hence we conclude that the localized
depletion of a substance that is essential for many cellular
processes, and especially for the completion of the cell cycle,
should be viewed as an important factor in maintaining the
quiescent state of these cells.
plants (Schiavone and Cooke, 1987; Liu et al., 1993; Hinchee
and Rost, 1992; Stange, 1988; Kerk and Feldman, 1994).
Hence alterations of other components of the model that relate
to the growth rate at the quiescent center should also disrupt
normal patterning of the root apex.
We suggest that our proposed biochemical model complements this biophysical growth model by providing a
mechanism for regulating the relative growth rate in this focal
region of a developing root meristem. Our model (Fig. 1)
accounts for the localized depletion of ascorbic acid in the QC,
and hence for the rate of cell growth and division in this
specific region of the root tip. In addition our work supports
the models of Hejnowicz and Hejnowicz which predict that
localized control of growth rates here is a determining aspect
in generating the pattern of cell walls in a given root tip. These
growth rates may be different in different species and at
different times during development, and thus underlie the
diversity of anatomical patterns that have been reported in root
tips. A similar type of mechanism may even underlie the organization of other kinds of meristems.
Implications of the model for meristem organization
How can this biochemical model be used to answer questions
of meristem organization? Hejnowicz and Hejnowicz (1991)
have modeled the formation of root apical patterning using a
biophysical perspective based on growth tensor analysis. One
aspect of their modeling predicts the cellular pattern
generated when a focus of slow growth is imposed in an
otherwise actively growing uniform growth field. Using
given rules for cell wall placement with respect to the
principal directions of growth, the model quite strikingly
predicts the formation of a closed meristem cell pattern as
shown in maize root apices (Hejnowicz and Hejnowicz,
1991). When a different tensor is applied, such that the focus
region becomes the region of maximal growth, the model
predicts the pattern in a pteridophyte root with a single apical
cell. This suggests that root apical patterning may be controlled by regulation of relative growth rates at this focal
point in the developing root meristem.
Our model is essentially a mechanism for controlling the
growth rate at this focal point. Therefore, experimental changes
that alter the steady state would be expected to have an effect
on apical patterning. For instance, inhibition of polar auxin
transport severely disrupts organization of the root meristem.
This has been shown for embryonic roots, lateral roots, intact
primary roots, and regenerating roots in many families of
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We are very grateful to Dr Ian Sussex for his critical reading of the
manuscript and helpful discussions and suggestions. We are also very
grateful to Dr Steven Ruzin who was so generous with his help with
microscopy and image processing in the NSF Center for Plant Development at Berkeley. Thanks also to Dr A. Shinmyo, Department of
Fermentation Technology, Osaka University, Japan, for his generous
gift of the ascorbate oxidase cDNA clone and antibodies that we used
in this study. We thank Prof. E. W. Weiler, Ruhr-Universität Bochum,
for the monoclonal antibody to auxin, and we also thank Dr Sundaresan, Cold Spring Harbor Labs for the p34cdc2 cDNA clone. This
work was supported by a USDA post-doctoral fellowship to N. Kerk,
and an NSF grant to L. Feldman.
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(Accepted 16 May 1995)