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Plant Physiol, December 2000, Vol. 124, pp. 1493-1506
New Techniques Enable Comparative Analysis of Microtubule
Orientation, Wall Texture, and Growth Rate in Intact Roots of
Arabidopsis
Keiko
Sugimoto,1
Richard E.
Williamson, and
Geoffrey O.
Wasteneys*
Plant Cell Biology Group, Research School of Biological Sciences,
Australian National University, Canberra, Australian Capital Territory
2601, Australia
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ABSTRACT |
This article explores root epidermal cell elongation and its
dependence on two structural elements of cells, cortical microtubules and cellulose microfibrils. The recent identification of Arabidopsis morphology mutants with putative cell wall or cytoskeletal defects demands a procedure for examining and comparing wall architecture and
microtubule organization patterns in this species. We developed methods
to examine cellulose microfibrils by field emission scanning electron
microscopy and microtubules by immunofluorescence in essentially intact
roots. We were able to compare cellulose microfibril and microtubule
alignment patterns at equivalent stages of cell expansion. Field
emission scanning electron microscopy revealed that Arabidopsis root
epidermal cells have typical dicot primary cell wall structure with
prominent transverse cellulose microfibrils embedded in pectic
substances. Our analysis showed that microtubules and microfibrils have
similar orientation only during the initial phase of elongation growth.
Microtubule patterns deviate from a predominantly transverse
orientation while cells are still expanding, whereas cellulose
microfibrils remain transverse until well after expansion finishes. We
also observed microtubule-microfibril alignment discord before cells
enter their elongation phase. This study and the new technology it
presents provide a starting point for further investigations on the
physical properties of cell walls and their mechanisms of assembly.
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INTRODUCTION |
Cell elongation and the direction of
organ expansion are linked processes of functional importance in plant
development. The structural features of the plant cell wall along with
its physiological activity dictate the mechanical properties that
regulate the rate and direction of cell expansion. The arrangement of
newly synthesized cellulose microfibrils (CMFs) is thought to play a
prominent role in determining these properties (Preston, 1974 ; Taiz,
1984 ; Carpita and Gibeaut, 1993 ) and cortical microtubules (CMTs) are
usually implicated in the alignment of CMFs. Two broad questions remain to be answered: (a) How does wall structure relate to the rate and/or
direction of expansion? (b) What role do CMTs play in determining wall
structure, growth rates, and other features of cell expansion?
The most favored but as yet unproven model for CMF alignment in
diffusely expanding cells suggests that movement of cellulose synthase
complexes (rosettes) along plasma membranes is driven by cellulose
crystallization but guided by CMTs (Herth, 1980 ; Giddings and
Staehelin, 1991 ). CMTs fit quite nicely as the likely modulator of
cellulose alignment but many lingering doubts remain. Previous
descriptions by transmission electron microscopy of CMTs in similar
orientations to CMFs are limited to very tiny areas of cells
(Palevitz and Hepler, 1976 ; Hardham et al., 1980 ; Seagull and Heath,
1980 ; Vesk et al., 1996 ). Furthermore, the striations seen in
glutaraldehyde-fixed, uranyl acetate/lead citrate stained thin sections
have been shown to not be CMFs in Equisetum hyemale root
hairs (Emons, 1988 ), raising some doubt as to their assumed identity in
earlier studies (Palevitz and Hepler, 1976 ; Hardham et al., 1980 ;
Seagull and Heath, 1980 ). Fluorescent probes that localize CMFs
have been used for observing CMF-CMT co-alignment during xylogenesis
(Falconer and Seagull, 1986 ), protoplast regeneration (Galway and
Hardham, 1989 ; Hasezawa and Nozaki, 1999 ), and cotton fiber development
(Seagull, 1986 ). The method, however, has low resolution and lacks
clarity when applied to analyze primary walls in multicellular tissues
(Sauter et al., 1993 ).
An indirect but more compelling proof of CMT function is that CMT
disruption leads to isotropic cell expansion (Green, 1962 ) or that it
prevents the establishment of a dominant axis of polarity (Galway and
Hardham, 1986 ). Some studies have demonstrated that CMT destabilizing
drugs cause disordered CMFs (Green, 1962 ; Hogetsu and Shibaoka, 1978 ;
Marchant and Hines, 1979 ; Robinson and Quader, 1982 ; Richmond, 1983 )
but in most cases the alignment of CMFs has been determined by
birefringence patterns and not confirmed at the ultrastructural level.
Other observations of inconsistent or nonexistent CMT-CMF co-alignment
have led to questioning of the role CMTs play in aligning CMFs (Itoh,
1976 ; Srivastava et al., 1976 ; Preston, 1988 ; Kimura and Mizuta,
1994 ).
The plethora of Arabidopsis root mutants (Baskin et al., 1992a ; Benfey
and Schiefelbein, 1994 ; Hauser et al., 1995 ) is helping to identify key
root morphology genes (Arioli et al., 1998 ), many of which have broader
roles in plant development. Understanding the role of wall structure in
cell growth and morphogenesis is an essential adjunct to the
molecular-genetic studies. Fortunately, Arabidopsis roots are also
ideal experimental systems with a predictable and simple anatomy (Dolan
et al., 1993 ) that remains uniform throughout root growth (Esau, 1977 ).
They are highly anisotropic; once cells leave the cell division zone,
there is no appreciable radial expansion (Baskin et al., 1992a ), and
individual cell growth is completed in a matter of hours. In this
article we present methods for examining CMF and CMT patterns in
Arabidopsis root epidermal cells. We then describe and compare CMT and
CMF alignment patterns in relation to growth kinetics of the wild-type
root under standard culture conditions.
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RESULTS |
Our goal was to describe and compare CMF and CMT patterns in
relation to the growth and development of epidermal cells in the
wild-type Arabidopsis root. To achieve this, we required methods enabling analysis of wall texture and CMT patterns in root tips that
remained essentially intact so that the stage of development of each
cell could be unequivocally determined. In addition, we wanted methods
that were sufficiently reliable and simple to allow their routine
application in experimental studies and in mutant characterization.
Root Growth and Elongation Profile
The consistent nature of Arabidopsis root growth on vertical agar
plates was critical to the correlative nature of this study. Under
standardized culture conditions, root elongation rates increased gradually with time, from 0.15 ± 0.02 mm
h 1 at d 4 to 0.24 ± 0.04 mm
h 1 at d 8, while root diameter remained
relatively constant at 0.14 ± 0.004 mm. Baskin et al. (1992a)
reported identical results. For all experiments, plants were analyzed
precisely 5 d after planting to avoid age-dependent differences in
growth profiles. Spatial distribution of root growth along this axis
was determined by marking the surface of roots with carbon grains and
then measuring the separation of two consecutive grains after a set
time interval (Fig. 1, A and B). Figure
1C depicts relative elemental growth rate (REGR) at set points along
the root axis. Roots exhibited a characteristic bell-shaped axial
growth curve with growth accelerating in the region 0.25 to 0.375 mm
from the root apex, peaking around 0.625 mm, and then declining until
it was undetectable around the 1.25-mm point. This growth profile
accords with the recent measurements of Mullen et al. (1998) , who used
a similar marker movement method, and Baskin et al. (1995) , who deduced
REGRs from kinematic measurements.

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Figure 1.
Spatial distribution of root elongation 5 d
after germination under standard growth conditions. A, Root tip labeled
with carbon grains showing positions of carbon grains at set
increments. B, One hour later, the separation of carbon grains
identifies the elongation zone. Bar = 100 µm. C, The growth
profile of 5-d root tips (n = 30), calculated by
measuring carbon grain separation, indicates that growth accelerates
between 0.25 and 0.375 mm from the tip, reaches a maximum at
approximately 0.625 mm, and declines to zero at 1.25 mm.
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Field Emission Scanning Electron Microscopy (FESEM) Examination of
CMF Patterns throughout Root Development
To accurately analyze CMF patterns at precise stages of cell
growth, we needed to first develop a method for examining cell walls in
roots that retained continuous files of epidermal cells. The best way
to achieve this was to remove one or more longitudinal cryosections to
expose an internal surface in the fixed root and then extract
sufficient cytoplasm to reveal wall texture in the exposed cells
without compromising the root tip's integrity (Fig. 2). In favorably oriented roots this
enabled analysis of wall structures in cells at every stage of cellular
development from the root cap to beyond the elongation zone. Epidermal
cells were identified most reliably along the edge of the cut (those in
the center were often cortex or endodermal cells) so most images were collected from the radial rather than inner tangential wall surfaces. As shown in Figure 3A, the cell walls
were occluded by cytoplasm if no extraction were attempted. Several
extraction procedures were compared. Chloroform/methanol, used to
solubilize lipids from crude cell wall fractions, removed relatively
little material and only occasionally revealed underlying cell wall
structures (Fig. 3B). Two detergents, saponin and Triton X-100 at 1%
(v/v), both removed substantial material and revealed wall
structures in some places but only after vigorous shaking for
48 h at 30°C (Fig. 3, C and D, respectively). By comparison,
10-min treatment with 0.1% (v/v) sodium hypochlorite removed
cytoplasmic material effectively and clearly (Fig.
4). This gave the most reliable results
without altering the integrity of the specimen and was used for all
subsequent analysis of wall structure.

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Figure 2.
Procedure for FESEM observation of walls from
Arabidopsis root tips. Longitudinal cryosectioning removes the outer
layer of the root so that after extraction, critical-point drying, and
coating with platinum, the microfibrillar texture of nearly every cell
can be examined.
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Figure 3.
Comparison of
extraction methods for preparing root tissues for FESEM analysis
following fixation and cryosectioning away the outer layer. After
extraction, all specimens were osmicated, dehydrated, critical point
dried, and platinum-coated. A, With no treatment, cytoplasmic debris
occludes cell wall. B, Chloroform-methanol (1:1) treatment for 1 h
at 40°C removes only some cytoplasmic material. C, One percent
(v/v) saponin for 48 h at 30°C reveals patches of wall
material. D, One percent (v/v) Triton X-100 for 48 h at
30°C is quite effective but takes considerable time. Bar = 300 nm.
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Figure 4.
Wall texture differs on inner (A) and outer (B)
face of a root epidermal cell. FESEM specimens were prepared by
extracting with 0.1% (v/v) sodium hypochlorite for 10 min, a
method that is fast and causes moderate but consistent extraction.
Images are oriented so that the transverse cell axis is parallel to the
short axis of the page. A, The most recently deposited layer, i.e. the
layer closest to the plasma membrane, has prominent transverse fibrils
across which lie short fibers (arrows). Pit fields (asterisk) were
frequently seen. B, The texture differed considerably on the outer face
of epidermal cells. Microfibrils have no preferred orientation and vary
greatly in apparent thickness. The more open texture and ability to
distinguish several lamina suggest matrix material is less abundant
than at the inner wall surface. Bar = 600 nm.
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Characteristics of Primary Walls of Root Epidermal
Cells
As shown in Figure 4A, the innermost layer of the root epidermal
cell wall was mainly composed of transversely aligned fibrous structures. The fibers had fairly uniform diameter, between 10 and 20 nm (including the 2.5-nm thickness of platinum coating) but thin fibers
around 5 nm or thicker bundles around 30 nm in diameter were observed
occasionally. Other very short fibrous structures around 60 nm in
length and 5 nm in diameter were also abundant (Fig. 4A, arrowheads).
These shorter fibers lay at right angles across a few of the longer
transverse fibers. Both fibrous structures were generally embedded in
amorphous material that was absent after extensive acid extraction
(Fig. 5A). Pit fields, from which all
fibrous structures were excluded, were also common (Fig. 4A, asterisk).
In contrast, examining the external face of the outer tangential wall
of the root epidermal cells revealed several layers of loosely and
randomly aligned fibers 30 to 40 nm in diameter (Fig. 4B). Compared
with the internal face, short fibrous components were not detected and,
as judged by the clarity of the fibrous structures, matrix material was
less abundant.

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Figure 5.
Comparative chemical
and enzymatic treatments helped identify the chemical nature of cell
wall components. A, Boiling in acetic acid:nitric acid:distilled water
(8:1:2) for 1 h thoroughly extracted the cytoplasm and wall matrix
but CMFs are retained. B, Cellulysin (0.1%, w/v) severely
degraded and ruptured the wall, leaving an open meshwork of residual
material, consistent with the predominant fibrous material being
cellulose. C, Pectolyase (0.1%, w/v) removed non-fibrous
material, modified the arrangement of fibrils, and created numerous
holes in the wall. In contrast, treatment with 0.125 units
mL 1 endo-1,4- -glucanase (D) or protease (E)
had no detectable effect on wall texture. Bars (bar in B is for
B-E) = 300 nm.
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Cellulosic Identity of Microfibrillar Structures
Several extraction analyses with acids,
polysaccharide-degrading enzymes, and proteases confirmed that the
dominant fibrous structures depicted in samples extracted with sodium
hypochlorite were CMFs. Boiling seedlings in an acetic acid-nitric acid
mixture extracts everything except crystalline cellulose (McCann et
al., 1990 ). On our cryo-sectioned samples, this thorough extraction procedure revealed a very well-defined wall texture (Fig. 5A), although
the fibrous structures were somewhat rearranged and aggregated. Figure
5B shows how a relatively mild, 1-h digestion with 0.1% (w/v)
Cellulysin, which degrades crystalline cellulose, reduced the
proportion of fibrous material. Longer treatments rendered samples
useless for examination. Pectolyase, which contains mainly endo-polygalacturonase and endo-pectin lyase, removed non-fibrous material and created numerous large pores when
applied at 0.1% (Fig. 5C) but left the long transverse fibrous
structures essentially intact. Neither endo-1,4- -glucanase (Fig.
5D), which degrades xyloglucan and non-crystalline cellulose but not
crystalline cellulose, nor protease (Fig. 5E) caused any change to the
overall wall structure or removed any specific components. This
diagnosis is consistent with observations reported for other species
(McCann et al., 1990 ; Carpita and Gibeaut, 1993 ), depicting transverse
CMFs embedded in pectic polysaccharides.
CMFs Remain Transverse throughout Elongation
Figure 6 shows a low magnification
micrograph of one example of a root from which cryosections have been
removed, along with high magnification images depicting CMF patterns
from specific cells, as indicated, along this root's length (Fig. 6,
A-F). CMFs were predominantly transversely aligned throughout the cell
division zone (Fig. 6A). The presence of pit fields, which occupy
relatively large proportions of the smaller cells of the division zone,
caused localized deviation of CMFs. Apart from this, CMFs deviated very little from the transverse axis, as revealed by quantitative analysis (Fig. 7C). CMFs remained transverse
throughout (Fig. 6, B-D) and even beyond the elongation zone (Fig.
6E). The only detectable difference during this period was that CMFs in
younger cells, i.e. those in mitotic and early elongation zones (Fig.
6, A-C), were less distinct than those in older, non-expanding cells
(Fig. 6, D and E), perhaps because of changes in matrix quantity in their walls. The occurrence of non-transverse, short fibers varied from
sample to sample but there was no clear trend with respect to their
abundance in relation to the stage of cell expansion.

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Figure 6.
Developmental progression of wall texture in root
tip from cell division zone to differentiation zone. The low-
magnification image of the root indicates the extracted cells from
which the higher magnification FESEM images (A-F) were taken.
Micrographs are oriented to show microfibril alignment relative to the
cell and root long axis, not the orientation of the root, which warped
slightly during processing. In the cell division zone (A) microfibril
alignment is already predominantly transverse. Note numerous pit
fields, undulating texture, and predominantly transverse microfibril
alignment. Microfibril orientation remains predominantly transverse
throughout the elongation zone (B and C), at the time of (D) and after
growth cessation (E). F, Deviation of CMFs from the transverse axis was
first detected approximately 2 mm from the tip where root hairs are
well developed. Bar for whole root image = 250 µm. Bar for A
through E = 500 nm.
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Figure 7.
CMT and CMF orientation relative to the cell long
axis plotted against distance from root apex in d 5 root tips. REGR,
replotted from Figure 4C. Mean microtubule angle (degrees) deviates
from the transverse axis about the time REGR begins to decline (B). In
contrast, CMFs remain predominantly transverse until well after growth
has ceased (C). Values are means from six roots (microtubules), three
roots (microfibrils), and 30 roots (REGR) with SD indicated
by error bars. All data points differing significantly from the
measured mean orientation at 0.3 mm from the root apex (#), as
determined by the independent Student's t test, are
indicated as (*).
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Groups of obliquely aligned CMFs were detected 1.9 mm or further from
the root apex, where cell expansion was complete (Fig. 6F). These oblique CMFs generally ran in groups of 10 to 20 roughly parallel fibrils. In addition, their alignment seemed to shift abruptly
from transverse to nearly 45 degrees to the long axis without any
obvious transition region. Quantitative analysis of CMF angular
dispersion revealed that they only deviated significantly from
transverse alignment in the region further than 1.5 mm from the root
apex (Fig. 7C). CMF deviation from the transverse axis conformed to a
right handed or Z-form helix as conventionally determined by viewing
from outside the cell.
Whole Mount Immunofluorescence and Confocal Microscopy
CMT patterns were recorded from immunolabeled intact root tips
using confocal laser scanning microscopy. The method included a mild
enzymatic digestion (see Fig. 5C for pectolyase treatment) and cold
methanol treatment, and labeled CMTs in nearly all epidermal root tip
cells, except those beyond the elongation zone, which were
permeabilized unreliably. This enabled CMT orientation patterns to be
analyzed in cells at known REGRs.
Changes in CMT Orientation Preceded Growth Cessation and
Changes in CMF Patterns
Figure 8A shows a typical image of
anti-tubulin labeling of CMTs in root epidermal cells at d 5. CMTs were
detected in cells from the root cap and in epidermal cells in the late
cell division and elongation zones, which emerge from beneath the root
cap. In the root cap cells, CMTs were predominantly transverse in
orientation. In the distal cell division zone, where growth rates were
still low, CMTs deviated considerably about the transverse axis (Fig. 8B; see also large SD of CMT alignment at 0.2 mm in Fig.
7B). Once the longitudinal axis of the cell exceeded the radial axis, however, CMTs were consistently transversely aligned in parallel arrays, and such alignment was maintained while growth rates were increasing (Figs. 7, A and B and 8, C-E). In the region where growth
was starting to decline, CMTs were far less numerous and, in some
cells, were obliquely oriented (Fig. 8F). CMTs in these cells formed
shallow helices that were consistently right-handed (of Z-helix form,
when viewed from outside the cell). Nearing completion of elongation,
CMTs varied from transverse through oblique to longitudinal within
individual cells (Fig. 8G). There was no obvious difference between CMT
patterns in trichoblasts versus atrichoblasts. (Trichoblasts could be
identified by the emergence of root hairs at the apical end of cells in
the same files.) Adjacent cells, however, frequently showed
considerable variation in CMT orientation patterns despite having the
same REGR (compare CMT patterns in adjacent cells in Fig. 8G).

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Figure 8.
Confocal micrographs of immunolabelled
microtubules in an intact root tip (A) and at higher magnification in
selected cells from the same root tip (B-G). Root cap cells obscure
epidermal cells in much of the cell division zone (A). CMTs were
variably oriented in the cell division zone (B), predominantly
transverse during the phase of increasing REGR (C-E), and oblique to
longitudinal as REGRs declined (F and G). Rotation of a confocal
optical series projection from region F reveals that oblique
microtubule orientation is consistent around the circumference of the
cell (H-J). The original projection (I) is rotated 40° to the left
(H) and right (J) to indicate microtubule alignment on the radial wall
surfaces. Bar in A = 25 µm. Bar for B through G in G = 5 µm. Bar for H through J = 5 µm.
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One potential drawback of our method was that CMF images were collected
from the radial or, less often, the inner tangential wall surfaces,
whereas the confocal scans of CMTs were mainly projected from the
perspective of the outer tangential wall. Lack of CMT-CMF congruence
during the phase of declining REGR could conceivably be attributed to
CMTs changing orientation on the outer tangential surface but remaining
transverse along the radial and inner tangential walls. We disproved
this possibility. Figure 8F is projected so that optical sections from
the entire cells are displayed. The criss-crossing of CMTs indicates
that the inner tangential wall CMTs continue the right-handed spiral
configuration and U-shaped bends show that CMTs also deviate from the
transverse axis to the same extent on all longitudinal surfaces. We
also performed digital rotations of Z-series projections. As
demonstrated in Figure 8, H-J, CMTs are obliquely oriented on the
radial cell surfaces with the same handedness as on the outer
tangential surface.
These results clearly indicate that CMTs are predominantly transverse
during the early phase of elongation but gradually shift to oblique and
then longitudinal orientations as REGR declines. Quantitative analysis
of CMT alignment along the major root axis revealed that the shift from
transverse to oblique orientation starts around 0.7 mm from the root
apex and gradually proceeds until CMTs are predominantly longitudinally
aligned at around 0.9 mm (Fig. 7B).
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DISCUSSION |
This study provides a detailed comparison of CMT and CMF
orientation patterns in epidermal cells throughout cell growth.
Elongation rates (REGR) were measured directly from living roots the
same age as and grown under identical conditions to the specimens
sampled for wall or CMT analysis. By keeping root tips intact while
preparing specimens for immunolabeling or FESEM analysis, we could
identify the growth status of any cell with reasonable accuracy. We
found that CMT and CMF orientation patterns were similar only during the period of accelerating elongation. CMFs were transverse to the
root's long axis from the cell division zone through to and beyond the
elongation zone. CMT orientation, however, was variable in the cell
division zone, transverse in the first half of the elongation zone, and
began to deviate from the transverse axis once REGR began to decline.
Despite this change in CMT orientation during the later stages of
growth, cell expansion remained anisotropic, with no appreciable
increase in root diameter throughout or beyond the elongation zone.
CMTs consistently formed right-handed helices as they deviated from the
transverse axis. Well after cell expansion stopped, CMFs were observed
sporadically in oblique orientations that also deviated in a
right-handed manner though by this stage, most CMTs were oriented
nearly longitudinally. The ensuing discussion addresses the technical
aspects of this work along with the implications of our findings for
current theory on plant cell wall construction.
FESEM Observations of the Walls of in Situ Cells
High resolution methods for accurately imaging recently deposited
CMFs are critical for elucidating the role these elements play in cell
expansion. Ultrathin sections examined by transmission electron
microscopy can reveal striations in the wall, assumed to be CMFs
(Preston, 1974 ; Sawhney and Srivastava, 1975 ; Hardham et al., 1980 ;
Lang et al., 1982 ) but, as Emons (1988) has demonstrated, these
striations do not always represent CMFs. Freeze-etching (Preston, 1974 ;
Mueller and Brown, 1980 ), dry-cleaving (Sassen et al., 1985 ; Emons,
1988 ), deep etching (McCann et al., 1990 ), and replica methods (Green,
1958 ) are more suitable but limited. Generally only small portions of
cell walls are visible and the lack of control over where observations
can be made make it difficult to correlate wall texture to growth
kinetics and/or CMT patterns. Our protocol, involving cryo-sectioning
the root tip followed by cytoplasmic extraction and shadow-casting,
overcomes these problems. It was possible to view the innermost layer
of cell walls at any position along roots while maintaining the intact root structure so that CMF orientation could be measured with respect
to the axis of expansion.
Field emission electron sources produce scanning electron micrographic
images at outstandingly high spatial resolution by providing small
diameter beams of high brightness (Crewe et al., 1968 ) and low
accelerating voltages (Pawley and Erlandsen, 1989 ). In plants, FESEMs
use so far has been limited to simultaneously viewing CMTs and cell
walls in partially extracted onion root tips (Vesk et al., 1994 ) and
tobacco BY-2 cells (Vesk et al., 1996 ) and determining the alignment of
CMFs in some woody species (Abe et al., 1995 ; Prodhan et al., 1995 ). To
view the innermost layer of primary cell walls, we planed a thin layer
from the frozen root surface and gently extracted cytoplasmic material
from the exposed cells with sodium hypochlorite. Conventional sliding
microtomes are often used for cryo-sectioning (Takeda and Shibaoka,
1978 ; Hogetsu, 1986 ; Abe et al., 1995 ; Prodhan et al., 1995 ) but in this study, a cryo-ultramicrotome cooled to 120°C with liquid N2 was used to prevent structural deformation by
ice crystals. Dimethyl sulfoxide (DMSO) was used as a cryo-protectant
for the same reason. The innermost layer of cell wall was covered with cytoplasm, in contrast to reports for other species using replica techniques (Takeda and Shibaoka, 1978 ; Hogetsu, 1986 ), and treatment with sodium hypochlorite was found to be most effective for removing this material. What hypochlorite extracts from cells is not fully understood, although it has been used to extract cytoplasm from woody
tissue (Abe et al., 1995 ). From its chemical structure it is likely
that hypochlorite only interacts with proteins and not polysaccharides.
Furthermore, the wall structure observed with this rapid extraction
procedure was similar to that obtained after extraction by milder but
longer detergent treatments. From these observations it is very likely
we were able to observe a good representation of nascent cell wall structure.
Primary Cell Walls of Arabidopsis Root Cells
FESEM revealed several fine structural details of Arabidopsis
primary cell walls. The transverse fibrous structures are most likely
to be CMFs based on the following evidence. (a) The diameter of these
fibrils was 10 to 20 nm. Taking approximately 2.5-nm thickness of
platinum coating into account, this is in good agreement with the
reported diameter of cellulose, 5 to 12 nm, in higher plants (Brown,
1982 ; McCann et al., 1990 ). (b) They were only digested by Cellulysin
but not by other tested enzymes. (c) They were aligned predominantly
transversely in expanding root cells as previously noted in many
studies (e.g. Gunning and Hardham, 1982 ). (d) Although modified, they
survived boiling in nitric-acetic acid, a treatment that crystalline
cellulose but not other polysaccharides can withstand (McCann et al.,
1990 ). There were additional short but still fibrous structures
observed in the primary walls. These were approximately 60 nm in
length, approximately 5 nm in diameter, and ran at right angles across
a few adjacent CMFs. These fibers were most likely hemicellulosic even
though endo-1,4- -glucanase did not appear to remove them. Similar
structures were observed in onion cells (McCann et al., 1990 ) and were
suggested to be hemicellulosic based on the fact that they were
extractable with alkali. The non-fibrous structures observed on these
walls were probably pectins as they were only extractable with
pectolyase. In general, our observations demonstrated that Arabidopsis
root epidermal cells have typical dicot primary cell wall architecture (McCann and Roberts, 1991 ; Carpita and Gibeaut, 1993 ). Further confirmation of the chemical nature of each structure will be achieved
in future work by labeling cellulose with gold-conjugated cellulases,
immunolabelling pectins (Knox, 1992 ; Willats and Knox, 1999 ), and
chemical extraction of hemicelluloses.
CMF Alignment Corresponds to Growth Anisotropy But Not Growth
Rate
FESEM revealed that CMFs remained predominantly transverse well
after growth rates started to decline and that they only became sporadically oblique long after cells ceased elongating. These observations support the view that CMF alignment regulates the direction but not the rate of cell expansion (Baskin et al., 1999 ). Instead, factors that control the rigidification of cell walls regulate
the decline and cessation of cell elongation (Pritchard et al., 1993 ;
Cosgrove, 1997 ), whereas turgor remains more or less constant
(Pritchard et al., 1993 ). These factors probably act in processes such
as reducing wall loosening, increasing wall cross-linking activities,
and/or altering the composition of the other wall polysaccharides to
make the wall more rigid or less susceptible to wall loosening.
Assessment of Immunofluorescence Protocol
To study developmental changes in CMT organization patterns, it is
essential to label the full range of cells within a developing tissue.
This has been achieved with epidermal peels (Roberts et al., 1985 ),
internodal strips (Flanders et al., 1990 ), partially digested shoot
apical meristems (Marc and Hackett, 1989 ), and wax or plastic resin
sections of roots (Baluska et al., 1992 ; Baskin et al., 1992b ; Liang et
al., 1996 ). We combined mild enzymatic digestion of cell walls with
cold methanol treatment to obtain consistent CMT labeling in nearly all
epidermal root tip cells. Unlike the laborious processes of embedding
and sectioning, this protocol allows us to process large numbers of
samples. Furthermore, since the whole structure of each cell is very
well preserved, one can reconstruct accurate three-dimensional patterns
from confocal Z-series. This method is also quite useful to study the
effects of drugs and/or mutations on the arrangement of CMTs in cells at various developmental stages and could probably be adapted for wide
ranging applications including in situ hybridization, for the analysis
of gene expression patterns.
Establishing Transverse CMT Orientation in the Early Elongation
Zone
In this study, we demonstrated that CMTs in Arabidopsis root
epidermal cells undergo a series of conformational changes according to
the developmental stages of cells. Although CMT orientation is
relatively variable in the cell division zone, CMTs become consistently
transverse as soon as elongation growth is detected. This is consistent
with observations made in a wide range of cell types (Clayton et al.,
1985 ; Wick, 1985 ; Flanders et al., 1990 ; Nagata and Hasezawa, 1993 ;
Baskin et al., 1999 ; Bichet et al., 2000 ; Wenzel et al., 2000 ).
Arabidopsis mutants with apparently normal division patterns but random
cortical arrays including fass (McClinton and Sung, 1997 )
and botero1 (Bichet et al., 2000 ) may help identify the
mechanism for establishing transverse CMTs. In the botero1
mutant, CMTs fail to consolidate into transverse arrays as cells enter
the elongation zone and growth anisotropy is impaired (Bichet et al.,
2000 ).
CMTs and Growth Rate Decline
This study revealed that CMTs start to become oblique as soon as
elongation rates start to decline (Fig. 7, A and B). The following
observations, however, strongly suggest that mechanisms controlling
growth rates are independent of CMT orientation. First, CMT patterns
varied considerably between adjacent cells that necessarily had
identical growth rates (note the large SD for CMT alignment at 0.8 and 0.9 mm in Fig. 7B). Traas et al. (1984) and Baluska et al.
(1992) also reported that oblique CMTs did not appear simultaneously in
all cells, suggesting that these shifts are only incidental features
accompanying the decline of elongation. The pattern of CMT deviation
from the transverse axis is also clearly variable from one cell type to
another. In pea (Hogetsu and Oshima, 1986 ) and maize (Baskin et al.,
1999 ) roots, re-orientation of CMTs begins only well after cell
elongation starts to slow down. In barley leaves, CMTs remain
predominantly transverse throughout the elongation zone but in
GA-deficient mutants, CMTs lose transverse order in the distal
elongation zone (Wenzel et al., 2000 ). Wasteneys and Williamson (1987 ,
1993 ) reported that internodal cells of two characean genera show
remarkable differences in the CMT transition patterns during the
cessation of their growth, yet these do not result in any obvious
differences in growth dynamics or morphology.
Whether the decline in the elongation rate causes shifts in CMT
orientation has also been addressed. Earlier work suggested that strain
might play a role in determining CMT orientation (Wasteneys and
Williamson, 1987 ), but four studies prove otherwise. When organ growth
was perturbed by experimental conditions such as irradiating pea stems
with blue light (Laskowski, 1990 ), treating Nitella
internodes with pH band-inducing medium (Kropf et al., 1997 ), and
keeping auxin-treated azuki bean epicotyls in anaerobic conditions
(Takesue and Shibaoka, 1999 ), CMTs remained or became transverse after
cell elongation declined or even ceased altogether. In maize pulvinal
cells, CMTs remain transverse for many days despite zero growth
activity (Collings et al., 1998 ), suggesting that CMT orientation is
not influenced by elongation rate.
Role of CMTs in Aligning CMFs
We determined that CMTs and CMFs are both transversely aligned
with relatively small standard deviations in the region of the root
where REGR is rising. This agrees with previous reports for roots of
other taxa (Gunning and Hardham, 1982 ; Hogetsu, 1986 ; Baskin et al.,
1999 ) but we also found that CMTs are not as uniformly transverse as
CMFs in the cell division-elongation transition zone and that CMTs and
CMFs have contrasting orientations in the distal elongation zone.
Because cell shapes in the cell division zone are disc shaped to nearly
isodiametric, CMFs were predominantly transverse to the root's long
axis but generally not to the cell's. This suggests either that CMF
orientation is not responsible for determining all aspects of cell
expansion in this region or that cells establish transverse CMF
patterns some time prior to the phase of highly anisotropic expansion.
The finding that tightly transverse CMF alignment is established before
CMTs are tightly transverse, together with data that indicate that CMTs
become transverse only after elongation commences (Bichet et al., 2000 ; Wenzel et al., 2000 ), questions whether CMTs control CMF alignment at
this stage of development. Similar doubts were raised in a recent study
that showed CMT randomization when cellulose synthesis was inhibited
(Fisher and Cyr, 1998 ).
In the distal elongation zone, CMTs were oblique to longitudinal, while
CMFs remained transverse. Lack of CMT/CMF parallelism in post
elongation zones has previously been reported (Hogetsu, 1986 ; Traas and
Derksen, 1989 ; Baskin et al., 1999 ). A time lag before cellulose
synthase complexes change their direction in response to changes in CMT
orientation could explain the disconnection, but by plotting the
x axis of Figure 7 as time instead of distance, we calculate
this lag time at 4.5 h, which is more than half of the estimated
8-h cell elongation period. Alternatively, CMT-dependent alignment
mechanisms could cease well before growth arrest, but CMFs might
continue to use already deposited cell wall layers as templates. It is
also conceivable that cellulose synthesis might be reduced as cell
expansion slows so that the observed CMFs in older cells might have
been laid down prior to changes in CMT orientation. Examining the
spatial distribution of cellulose synthesis along the long axis of
roots will help test this third possibility.
 |
CONCLUSION |
Our results indicate that close CMT/CMF co-alignment occurs only
in the proximal elongation zone when REGRs are increasing. Throughout
development, CMF orientation is more consistently transverse than CMT
orientation, a result that questions the nature of the relationship
between CMT and CMF alignment. We are currently applying these methods
to examine CMT and wall patterns in Arabidopsis roots challenged by
drugs or mutations that target cell wall or CMT-specific functions. By
using this approach we hope to improve the understanding of wall
construction in plant cells.
 |
MATERIALS AND METHODS |
Plant Material and Growth Conditions
Arabidopsis ecotype Columbia was used throughout this study.
Seeds were surface sterilized in a mixture of 3% (v/v) hydrogen peroxide and 50% (v/v) ethanol for 2 min. After rinsing in
sterilized water, seeds were planted on nutrient-solidified agar (Bacto
Agar, DIFCO Laboratories, Detroit) plates [2 mM
KNO3, 5 mM Ca(NO3)2, 2 mM MgSO4, 1 mM
KH2PO4, 90 µM iron-EDTA complex,
46 µM H3BO3, 9.2 µM
MnCl2, 0.77 µM ZnSO4, 0.32 µM CuSO4, 0.11 µM
MoO3, 1% (w/v) Glc, 1.2% (w/v) agar, 530 µM
myo-inositol, and 50 µM thiamine hydrochloride]. Plates
were sealed with laboratory film and held vertically in a growth
cabinet under constant light (80 µmol m 2
s 1) and temperature (21°C).
Measurement of Root Growth
To measure the rate of root elongation, the position of the root
apex was scored with a razor blade on the outside of plastic Petri
plates every 12 h. At the end of experiments each plate was
photocopied and the distance between each mark was measured using the
digital image analysis program, Image I (Universal Imaging, West
Chester, PA). Root diameter was measured approximately 0.5 mm
from the root apex using a dissection microscope (SMZ-2T, Nikon, Tokyo)
equipped with an ocular micrometer. Average elongation rates and root
diameters with SD were calculated from data on at least 30 roots grown in three different plates.
Growth Kinetics
To examine the spatial distribution of elongation along roots,
carbon grains were applied to the upper surface of roots with an
eyelash and their positions photographed at 1-h intervals (Fig. 1).
Separation of carbon grains at specific distances from root tips was
measured on photographic prints using Image I software and REGRs were
calculated as follows:
where xf is the final distance
between two carbon grains, xi is the initial
distance between two carbon grains, and t is the time interval.
Cell Wall Preparation
Whole seedlings were fixed in 4% (v/v) formaldehyde made
up PEM buffer (25 mM PIPES
[1,4-piperazinediethanesulfonic acid], 0.5 mM
MgSO4, 2.5 mM EGTA, pH 7.2), rinsed three times
in PEM buffer, and cryoprotected in 25% and 50% (v/v) DMSO (in
PME buffer, 10 min each step). Root tips were excised with a razor
blade, placed on a nail head, and immediately frozen in liquid
nitrogen. After slicing off the surface of a root with a glass knife on a cryo-ultra-microtome (Ultracut E ultra-microtome with FC 4 cryo-microtomy attachment, Reichert-Jung, Vienna), while
maintaining samples and knives at 120°C, the remaining portion of
the root was thawed in 50% (v/v) DMSO, and then transferred
into PEM buffer. Several different treatments were tested to extract
membranes and other cytoplasmic materials: (a)
chloroform/methanol (1:1) twice for 1 h at 40°C, followed
by methanol and acetone treatment, each for 30 min; (b) 1%
(v/v) saponin for 48 h at 30°C; (c) 1% (v/v) Triton X-100 for 48 h at 30°C; (d) 0.1% (v/v) sodium
hypochlorite for 10 min; (e) boiling in acetic acid and nitric acid and
distilled water (8:1:2) for 1 h. After thoroughly rinsing in
distilled water, osmication in cold 0.5% (v/v) OsO4
for 15 min, rinsing in distilled water, and dehydration through a
graded ethanol series (30%, 50%, 70%, 95%, and 100% three times,
15 min for each step), specimens were critical point dried using
CO2.
FESEM
All specimens were mounted on stubs with double-sided sticky
carbon tape, cut surface facing upward, and coated with platinum at 2.4 mA for 195 s. Cell wall fine structure was examined on a Hitachi
S4500 FESEM (Hitachi, Tokyo), fitted with a solid state backscattered
upper electron detector at 5 kV. The condenser lens was set between 8 and 9 with a working distance between 5 and 8. Scanned images were
recorded with Image slave 2.11 (Meeco, Melbourne).
Cell Wall Digestion by Enzymes
To identify the chemical basis of cell wall structures observed
by FESEM, root specimens were treated with several
polysaccharide-degrading enzymes and proteases before
treatment with 0.1% (v/v) sodium hypochlorite. Tested
enzymes were (a) 0.1% (w/v) Cellulysin (Calbiochem-Novabiochem, San Diego) in PEM buffer for 1 h at root temperature, (b) 0.1% (w/v) Pectolyase Y-23 (Kikkoman, Tokyo) in PME buffer for
30 min at room temperature, (c) 0.125 units mL 1
endo-1,4- -glucanase (Megazyme International, Bray, County Wicklow, Ireland) in 50 mM sodium acetate (pH 4.7) for 5 h at
37°C, (d) 0.1% (w/v) Pronase (Boehringer Mannheim,
Mannheim, Germany) in Tris-buffered saline (pH 7.4) for
1 h at room temperature.
Immunofluorescence
Whole seedlings were fixed in 1.5% (v/v) formaldehyde
and 0.5% (v/v) glutaraldehyde made up in PEMT buffer (50 mM PIPES, 2 mM EGTA, 2 mM
MgSO4, 0.05% [v/v] Triton X-100, pH 7.2) for 40 min and rinsed in PEMT buffer three times for 10 min. They were subsequently digested with 0.05% (w/v) Pectolyase Y-23
(Kikkoman) in PEM buffer (50 mM PIPES, 2 mM
EGTA, 2 mM MgSO4) with 0.4 M mannitol for 20 min, rinsed in PEM buffer three times, treated with
20°C methanol for 10 min, and rehydrated in phosphate-buffered saline (PBS) for 10 min. Autofluorescence caused by free aldehydes from
glutaraldehyde fixation was reduced with 1 mg mL 1
NaBH4 in PBS for 20 min, followed by the treatment with 50 mM Gly in PBS (incubation buffer) for 30 min. Seedlings
were incubated with primary antibodies against tubulin at room
temperature overnight, rinsed in incubation buffer three times for 10 min, and secondary antibodies applied for 3 h at 37°C. After
rinsing in PBS three times, root tips were cut off from the rest of
seedlings, and mounted in 0.1% (w/v) para-phenylene diamine in 1:1
PBS-glycerol, pH 9. Cut cover glasses were used to space slide and
cover glasses so as to avoid crushing delicate root tips.
Antibodies
For this study, anti- tubulin (product N357, Amersham,
Buckinghamshire, England) was used at a dilution of 1:100. Fluorescein isothiocyanate-conjugated anti-mouse IgG (Silenus/Amrad Biotech, Boronia, Victoria, Australia), diluted 1:100, was used as a secondary antibody.
Confocal Laser Scanning Microscopy
Immunofluorescence images were collected with an MRC- Bio-Rad
600 (Microscience Division, Hemel Hempstead, UK) confocal
laser-scanning microscope, coupled to a Zeiss Axiovert IM-10 inverted
microscope. Excitation at 488 nm with an argon ion laser was used for
fluorescein isothiocyanate fluorochromes. Images were collected using a
Plan Neofluar 100-X objective lens, N.A. 1.30, following Kalman
averaging of six full scans. Three-dimensional images from the root
apex to the differentiation zone were obtained by collecting series of
approximately 60 optical sections, each section approximately 0.6 µm
thick, in the Z axis. Images were processed with image processing software programs including COMOS 7.0 (Microscience Division, Hemel Hempstead, UK), Confocal assistant 4.02 (written by
Todd Clark Brelje) and Adobe Photoshop 4.0 (Adobe Systems, Mountain
View, CA). All confocal images were flipped about the vertical axis to
adjust for the inverted stage of our microscope.
Measurement and Analysis of Microtubule and Cellulose
Alignment
The alignments of CMTs and CMFs were measured from printed
images obtained by confocal microscopy and FESEM, respectively. Angular
deviation from the longitudinal axis of roots was measured with Image I
software. Transverse orientation was defined as 90° to the long axis
plotted along the right vertical axis of the image, as viewed from the
outside of the cell (as happened in immunofluorescence) so that FESEM
images (taken looking from inside the cell) were flipped. Elements
aligned at angles between 0° and 90° (counterclockwise from top)
formed a left-handed helix while those elements oriented between 90°
and 180° formed a right-handed helix. For CMTs, each data point
represented a total of 60 CMTs from six different roots. CMF
measurements were taken from 2.4 µm × 1.5 µm wall patches,
which were sampled from several adjacent cells at each distance from
the apex. Regions around pit fields were avoided. Each data point
represented a total of 300 CMFs, combined from 30 wall patches from
three different roots. SD was calculated from CMT and CMF
angular measurements. Means between data points were compared by the
independent Student's t test for samples with unequal
variance (Microsoft EXCEL), at a significance level of 0.001. In
general, microfibrils were straight and uniformly aligned across the
observed window, except occasionally they appeared slightly wavy. Very
little variation between sample windows or between cells at a similar
position was observed. Fixation and all other processes involved for
the FESEM preparation did not cause any major artifact so that the
great majority of cells could be analyzed.
 |
ACKNOWLEDGMENTS |
We thank Frank Brink, Sally Stowe, and David Vowles of the ANU
Electron Microscopy Unit for their assistance and tuition with the
FESEM technique, and Tobias Baskin, Brian Gunning Nori Hasenbein, and
Owen Schwartz for helpful advice.
 |
FOOTNOTES |
Received August 9, 2000; modified September 14, 2000; accepted September 27, 2000.
1
Present address: Department of Cell Biology,
John Innes Centre, Colney, Norwich, NR4 7UH, UK.
*
Corresponding author; e-mail geoffw{at}rsbs.anu.edu.au; fax
61-0-2-6125-4331.
 |
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