Plant Physiol. (1998) 117: 501-513
Auxin Deprivation Induces Synchronous Golgi Differentiation in
Suspension-Cultured Tobacco BY-2 Cells1
Zev M. Winicur2, *,
Guo Feng Zhang3, and
L. Andrew Staehelin
Department of Molecular, Cellular, and Developmental Biology,
University of Colorado, Boulder, Colorado 80309-0347
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ABSTRACT |
To
date, the lack of a method for inducing plant cells and their Golgi
stacks to differentiate in a synchronous manner has made it difficult
to characterize the nature and extent of Golgi retailoring in
biochemical terms. Here we report that auxin deprivation can be used to
induce a uniform population of suspension-cultured tobacco
(Nicotiana tabacum cv BY-2) cells to differentiate
synchronously during a 4-d period. Upon removal of auxin, the cells
stop dividing, undergo elongation, and differentiate in a manner that
mimics the formation of slime-secreting epidermal and peripheral
root-cap cells. The morphological changes to the Golgi apparatus
include a proportional increase in the number of
trans-Golgi cisternae, a switch to larger-sized
secretory vesicles that bud from the trans-Golgi
cisternae, and an increase in osmium staining of the secretory
products. Biochemical alterations include an increase in large,
fucosylated, mucin-type glycoproteins, changes in the types of secreted
arabinogalactan proteins, and an increase in the amounts and types of
molecules containing the peripheral root-cap-cell-specific epitope JIM
13. Taken together, these findings support the hypothesis that
auxin deprivation can be used to induce tobacco BY-2 cells to
differentiate synchronously into mucilage-secreting cells.
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INTRODUCTION |
As already recognized by plant anatomists in the latter half of
the 19th century and recently rediscovered with the introduction of
novel antibody probes, plant tissues can be distinguished not only by
their size and shape, but also by the chemical properties of the cell
walls of each tissue (Knox and Roberts, 1989
; Pennell et al., 1989;
Knox et al., 1991; Roberts, 1994). Because the Golgi apparatus is the
site where the bulk of the cell wall matrix molecules are synthesized
(e.g. complex polysaccharides, AGPs, and Hyp-rich glycoproteins), it is
obvious that during tissue development the secretory apparatus of each
cell must adapt to the changing synthetic demands associated with the
production of different types of cell wall molecules (Driouich et al.,
1994).
Direct evidence for the tissue-specific retailoring of plant Golgi
stacks has come mainly from two sources: electron microscopic observations of cryofixed cells and immunocytochemical studies of cells
labeled with antibodies against defined epitopes of cell wall
molecules. For example, in tobacco (Nicotiana tabacum) and Arabidopsis root tips, characteristic changes in both the architecture and the cisternal-staining patterns of Golgi stacks have been shown to
accompany the developmental differentiation of meristematic cells,
first into gravity-sensing columella cells and then into slime-secreting peripheral cells (Staehelin et al., 1990). Similarly, activation of the secretory apparatus of barley aleurone cells by the
hormone GA3 leads not only to an increase in the
number and size of Golgi-associated vesicles, but also to changes in the size distribution of freeze-fractured particles in the Golgi membranes (Fernandez and Staehelin, 1985).
In clover root tips certain pectic polysaccharide epitopes are
typically associated with the medial and cis-Golgi cisternae in cortical cells, but in peripheral root-cap cells they are associated with the trans-Golgi cisternae and the TGN (Lynch and
Staehelin, 1992, 1995). Similar variations in Golgi-enzyme localization
have been found in different animal cell types (Roth, 1991).
Tissue-specific retailoring of plant Golgi stacks has also been found
to include the development of structures known as intercisternal filaments, which are located between trans cisternae and
sometimes between medial-trans cisternal pairs of Golgi
stacks in mucilage-secreting cells, but not in meristematic cells
(Mollenhauer, 1965; Turner and Whaley, 1965; Mollenhauer and
Morré, 1975; Staehelin et al., 1990). Although their function has
yet to be determined, the fact that they seem to align with protein
complexes in the cisternal membranes suggests that they may anchor the
Golgi enzymes involved in mucilage synthesis, thereby preventing them
from being dragged into the large secretory vesicles during the
packaging of these very large molecules (Staehelin et al.,
1990).
The characterization in plants of the nature and extent of Golgi
retailoring in biochemical terms requires the isolation and purification of Golgi stacks from specific cell types in quantities sufficient for biochemical analysis. Because cost-effective methods for
producing such Golgi fractions from plant tissues have yet to be
formulated, we have sought to develop an alternative approach by
inducing a uniform population of suspension-cultured cells to
differentiate in a synchronous manner.
Plant growth and development is controlled to a significant extent by
seven types of hormones: auxins, cytokinins, GAs, ethylene, ABA,
brassinosteroids, and jasmonates. In this study we have focused on the
role of auxins in the differentiation of slime-secreting cells. Auxins
are a group of natural and synthetic plant hormones that affect cell
growth and division (Taiz and Zeiger, 1991). For example, the
application of the natural auxin IAA to shoots stimulates cell
elongation, whereas its application to roots inhibits elongation and
promotes adventitious root formation. At the cellular level, one of the
earliest responses in pea stem epidermal cells to IAA treatment is a
transient increase in the percentage volume fraction of Golgi stacks in
the cytoplasm, but this increase lasts for less than 90 min
(Cunninghame and Hall, 1985). A more sustained increase in the amount
of Golgi material, in parallel with increased rates of cell elongation,
has been noted in IAA-treated oat coleoptiles (Quaite et al., 1983).
Auxin also affects a number of developmental processes, such as
gravitropism, leaf abscission, and fruit development. Removal of auxin
from the growth medium of suspension-cultured carrot cells has been
shown to cause their arrest in G1 (Nishi et al., 1977) and to induce
rapid cell elongation (Lloyd et al., 1980). Furthermore, auxin
deprivation can be used to induce anthocyanin production (Ozeki and
Komamine, 1981), alterations in cell wall polysaccharides (Masuda et
al., 1984), and glycosidase activities (Masuda et al., 1985).
Based on these findings, we hypothesized that by manipulating auxin
levels, we may also be able to manipulate the secretory activity and
functional organization of Golgi stacks in tobacco BY-2 cells in a
reproducible manner. The BY-2 cell line (Nagata et al., 1992) is well
suited for experimental studies; it grows quickly and its cell cycle
can be synchronized to about 70% with a combination of aphidicolin and
propyzamide (Nagata et al., 1992; Samuels et al., 1995). More
importantly for our studies, BY-2 cells can be grown with only the
synthetic auxin analog 2,4-D as a hormone supplement. Therefore, one
would expect the cells to respond to the removal of this hormone from
the growth medium.
Here we report that auxin deprivation can be used to induce tobacco
BY-2 cells to differentiate synchronously into a mucilage-secreting type of cell during a 4-d period. The cells stop dividing and undergo a
process of synchronized elongation and differentiation, which includes
morphological changes in Golgi stack membranes and biochemical changes
in glycoprotein and proteoglycan secretion. We discuss the potential
usefulness of this system in studying the molecular basis of
tissue-specific Golgi stack retailoring in plants.
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MATERIALS AND METHODS |
Plant Materials and Culture Conditions
Suspension-cultured tobacco (Nicotiana tabacum cv BY-2)
cells were kindly provided by Dr. R. Cyr (Department of Biology,
Pennsylvania State University, University Park) and cultured in a
modified Murashige-Skoog basal salt medium (no. 5524, Sigma) in which
KH2PO4, thiamine-HCl, and
inositol were increased to 370, 1, and 100 mg/L, respectively. The
cells grown in this medium were supplemented with 3% Suc and 0.2 mg/L
2,4-D at pH 5.0 on a shaker (125 rotations/min) at 25°C in the dark.
Cells were subcultured by regularly transferring 2 mL of a 7-d-old
culture into 100 mL of fresh medium in a 500-mL flask.
Auxin-deprivation experiments for microscopy were performed by briefly
centrifuging (30 s at 200g) 2 mL of a 7-d culture, washing
the cells twice with modified Murashige-Skoog medium prepared without
2,4-D, and resuspending the cells in 100 mL of the auxin medium in a
500-mL flask as before.
Light Microscopy and DAPI Staining
Light micrographs of control and auxin-deprived tobacco BY-2 cells
were taken (Axioskop MC100 microscope, Zeiss), and cell lengths and
widths were measured from negatives. In cells found in files, cell
length was defined as the dimension perpendicular to the plane of cell
division. In isolated cells, cell length was defined as the longest
dimension.
BY-2 cell nuclei were stained with DAPI. Cells were fixed with
ethanol:acetic acid, 3:1 (v/v) for 1 h, briefly spun in a
microcentrifuge at 3000g, resuspended in culture medium, and
brought to 1 µg/mL DAPI. The mitotic index (the percentage of cells
in mitosis) was determined using a fluorescence microscope (Zeiss) with
an excitation filter of 365 nm and a barrier filter of 420 nm.
The rat monoclonal antibody JIM 13 was a generous gift of Drs. Paul
Knox and Keith Roberts and is described by Knox et al. (1991).
High-Pressure Freezing and Freeze Substitution
Five- to 7-d-old cultures were harvested by centrifugation
(1000g) and mixed in a 1:9 (v/v) ratio with 20% (w/v)
aqueous dextran (Mr 38,800) in fresh
culture medium to act as an extracellular cryoprotectant. This cell
suspension was then poured onto a 30-µm nylon mesh and further
concentrated by wicking off the liquid over the lip of an Erlenmeyer
flask. Specimens were frozen in a high-pressure freezing apparatus
(model HPM010, Balzers, Hudson, NH), as described by Craig and
Staehelin (1988), and were stored in liquid nitrogen before freeze
substitution. Samples were freeze substituted by the method of Zhang
and Staehelin (1992), infiltrated at room temperature for 5 to 6 d
in increasing concentrations of resin diluted in acetone, and embedded
either in Embed 812 or London Resin White (Polysciences, Warrington,
PA) at 55°C overnight.
Electron Microscopy
Thin sections (about 0.1 µm) were cut (Ultracut E, Reichert,
Vienna, Austria) and picked up on Formvar-carbon-coated copper or
nickel grids (200 mesh, Polysciences). The Embed 812-embedded sections
were mounted on copper grids and counterstained with 2% uranyl acetate
in water for 5 min and Reynold's lead citrate for 1 min, and examined
at 80 kV in an electron microscope (model CM10, Philips, Eindhoven, The
Netherlands). Nickel grids were used exclusively for antibody-labeling
experiments.
The number of trans and total cisternae were counted from
electron micrographs of Golgi at 4 d for control cells and at 1.5 and 4.5 d for auxin-deprived cells (n > 45).
Comparisons were made between the groups using an ANOVA test.
The London Resin White-embedded sections were immunolabeled with JIM 13 (at full strength) and a 10-nm colloidal gold-conjugated goat anti-rat
secondary antibody using the method of Zhang and Staehelin (1992). The
average number of gold particles per Golgi and per equivalent area of
cytoplasm (0.14 µm2) were counted for controls
and for cells subjected to 1.5 and 4 d of auxin deprivation.
Comparisons were made between the groups using an ANOVA test.
Secreted Material
Secreted material was collected from BY-2 cells grown with or
without auxin at 3 d after subculture. Ten grams of cells was incubated in 200 mL of culture for 3 h at 24°C. The medium was collected and the insoluble material was spun out at 27,000g
for 40 min at 4°C. Secreted proteins were collected by dialyzing the samples against distilled water with a 12,000 to 14,000 Mr cutoff membrane (no. 25225-260, VWR
Scientific, West Chester, PA). Secreted proteins were analyzed by
standard SDS-PAGE on 12 to 15% gradient gels. Samples were normalized
for protein concentration before loading, and the gels were silver
stained (Blum et al., 1987).
Gels were blotted to nitrocellulose, and the expression of the JIM 13 epitope was analyzed by the standard western-blot procedure. The
expression of glycoproteins was analyzed by a modified western-blot procedure using horseradish peroxidase-conjugated concanavalin A (no.
L8146, Sigma) for the detection of high-Man N-linked glycans and UEA-1 (no. L6397, Sigma) for the detection of complex N-
and O-linked glycans.
AGPs were analyzed quantitatively and qualitatively by rocket and
crossed-gel electrophoresis, respectively. The method of van Holst and
Clarke (1986) was used, except that 15 (instead of 30) µg/mL of
-glucosyl Yariv's artificial antigen (no. 100-4, Biosupplies
Australia, Parkville) was used in the gel. A linear relationship
between AGP concentration and rocket height was confirmed with gum
arabic (Biosupplies Australia) as a standard.
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RESULTS |
Effects of Auxin Deprivation on Cell Division and Cell
Elongation
Tobacco BY-2 cells were subcultured into fresh Murashige-Skoog
medium containing either 0.2 mg/L 2,4-D (control) or no 2,4-D (auxin
deprived). Cells deprived of auxin remained physiologically active for
about 1 week, as judged by their cytoplasmic streaming and by their
ability to be stained with fluorescein diacetate, a vital dye (data not
shown).
Cells were fixed with ethanol:acetic acid, 3:1 (v/v), stained with
DAPI, and observed under a fluorescence microscope to determine the
effects of auxin deprivation on the mitotic index. Mitotic nuclei were
counted and the mitotic index was determined at different time points
after subculture. We found that in cells of a 7-d culture, after about
12 h of auxin deprivation, the mitotic index dropped from 1.5 to
0%, in contrast to that of control cells, which increased from 1.5 to
6% 1 d after subculture (Fig. 1).
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| Figure 1.
Inhibition of cell division is induced by
auxin-deprived growth conditions. The average mitotic index of control
cells ( ) increased from 1.5 to about 6% after 1 d. The average
mitotic index of auxin-deprived cells ( ) decreased from 1.5 to 0%
after 12 h. SE is indicated for each data point by
error bars.
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BY-2 cells were deprived of auxin for variable amounts of time and
transferred back into control growth medium to test whether the cells
cultured without auxin lost their ability to reenter the cell cycle.
Their mitotic indices were then monitored. These experiments
demonstrated that cells deprived of auxin for up to 5 d were able
to reenter the cell cycle, but the longer they went without auxin, the
longer they took to recover. For example, after 3 d of auxin
deprivation the mitotic index of the cells transferred to the
auxin-containing medium took about 7 d to return to control levels
(about 6%).
Auxin deprivation also caused a change in general cell size and shape
(Fig. 2). Within 24 h most of the
auxin-deprived cells began to form clumps, in contrast to the mostly
chain-like structures of control cultures. The lengths and widths of
more than 100 cells were measured after 3 d in both control and
auxin-deprived media (Fig. 3). The
scatterplots show that auxin deprivation not only caused an increase in
average length and width, but also in the variance of sizes. The
outliers in the scatterplot after auxin deprivation were both long,
thin cells and large, round cells.
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| Figure 2.
Light micrographs of BY-2 tobacco cells
3 d after subculture with (a) and without (b) auxin. Control cells
were found both in long chains (small arrow) and in clusters (large
arrow). Auxin-deprived cells were elongated and did not form chains.
Bars = 0.2 µm.
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| Figure 3.
Scatterplot showing length and width of both
control (top) and auxin-deprived (bottom) cells. Although auxin
deprivation caused an increase in both average cell length and width,
the most striking change was in cell length. The variance of the values
also increased with auxin deprivation.
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Cell length was measured at different times and the median length was
calculated for each to further analyze the effects of auxin deprivation
on cell size. An average median length was determined based on three
separate experiments (Fig. 4). This
dimension was chosen because the greatest overall change in cell size
was in length rather than in width (Fig. 3). Whereas the length of
control cells varied between 40 and 60 µm during the 6-d culturing
period (the slight decrease in length after 1 d corresponding to
the increase in mitotic index), auxin-deprived cells essentially
doubled their average length, from 54 to 110 µm during the first
3 d of culturing, and then maintained that length. An increase in
the size of the vacuoles appeared to drive most of this growth.
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| Figure 4.
Auxin-deprived growth conditions induced
elongation of BY-2 cells. The median cell length of the control cells
() varied between 40 and 60 µm and decreased after 1 d. This
decrease corresponded to the increase in mitotic index (Fig. 1). The
median cell length of the auxin-deprived cells () increased to 110 µm after 3 d and then tapered off. SE is indicated
for each data point by error bars.
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Morphological Changes in the Golgi Apparatus
Morphological changes in the Golgi apparatus of auxin-deprived
BY-2 cells were analyzed in cells preserved by
high-pressure-freezing/freeze-substitution methods and examined by
electron microscopy. In control cells the Golgi stacks were typically
dispersed singly or in small groups throughout the cytoplasm and did
not change their morphological characteristics during a 6-d culturing
period (Fig. 5a). In contrast, auxin
deprivation caused major clustering of the Golgi stacks in the vicinity
of the nucleus (Fig. 5b) and systematic changes in Golgi stack
architecture (Figs. 6 and
7). The clustering was already pronounced
within 24 h of treatment, whereas the morphological changes
progressed more gradually.
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| Figure 5.
Electron micrographs of Golgi stacks from control
(a) and auxin-deprived (b) BY-2 cells. In the control cells the Golgi
stacks had a uniform distribution throughout the cytoplasm. In the
auxin-deprived cells the Golgi stacks clustered together near the
nucleus. G, Golgi stacks; M, mitochondria; N, nucleus; and V, vacuole.
Bars = 0.5 µm.
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| Figure 6.
Electron micrograph of a Golgi stack from a
control BY-2 cell. The cis, medial, and
trans cisternae are labeled. Intercisternal elements are
indicated with arrows. Bar = 0.2 µm.
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| Figure 7.
Electron micrographs of Golgi stacks from
auxin-deprived BY-2 cells. Cells were fixed at 14 h (a), 1.5 d (b), and 4 d (c) after auxin deprivation. The
cis, medial, and trans cisternae are
labeled in b and c. Intercisternal filaments are indicated by
arrowheads. After 14 h one could already see hypertrophied cisternal margins and a clustering of the stacks. After 1.5 d an
increase in the size of the cisternal margins and an increase in the
staining of intercisternal elements could be seen. After 3 to 4 d
densely staining material appeared in the medial cisternae, and
misshapen or thickened regions appeared in the interior of the Golgi
stack. Bars = 0.2 µm.
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The auxin deprivation-induced alterations in Golgi stack architecture
involved both the structure of the different types of cisternae and
their marginal buds and the staining of the lumenal contents. A typical
control Golgi stack is illustrated in Figure 6. Based on the
morphological criteria defined by Staehelin et al. (1991), this Golgi
stack possessed one cis, four medial, and two
trans cisternae. Typical features of cis
cisternae are their end location, smaller-than-average diameter, wider
lumen, and relatively light staining of membrane and lumenal contents.
The medial cisternae occupy the central section of the stack. They have
a well-defined, flat central domain and margins that are both
fenestrated and bulbous. The contents of the relatively wide medial
cisternae exhibited a mottled staining pattern that increased with
density toward the trans cisternae. The trans
cisternae were recognized by their osmotically collapsed lumen, where
the membranes appeared tightly pressed together, and their darkly
staining products. A TGN was seen associated with some but not all
Golgi stacks.
Because the TGN was not infrequently displaced to one side of the Golgi
stacks in suspension-cultured cells (Fig. 6; see also Zhang and
Staehelin, 1992), the lack of a TGN in any given micrograph of a Golgi
stack may simply reflect the plane of the section. However, in some
instances the trans-most trans-Golgi cisterna assumes certain structural characteristics of a TGN, such as branched marginal domains, suggesting that it may assume TGN functions.
In response to auxin deprivation, the Golgi stacks underwent distinct
morphological changes in a synchronized manner (Fig. 7). By 14 h
after the cells were transferred to the auxin-free growth medium, the
first clustering of Golgi stacks was seen, producing large,
ribosome-excluding domains, and some of the trans cisternae
exhibited slightly hypertrophied cisternal margins (Fig. 7a). One to
1.5 d after auxin deprivation, large, dense vesicles accumulated
around the stacks, and the bulbous margins of all trans
cisternae became hypertrophied (Fig. 7b). In addition, the TGN
disappeared from the trans side of the Golgi stacks and was replaced by an accumulation of densely staining vesicles. After 3 to
4 d, the bulbous margins of the trans- Golgi cisternae
doubled in diameter from 45 to more than 90 nm, which corresponds to an 8-fold increase in volume, and their contents became more heavily and
uniformly stained (Fig. 7c). Densely staining contents also accumulated
in the medial Golgi cisternae, and the medial and trans
cisternae more frequently developed misshapen or expanded regions in
the interior of the Golgi stacks.
Although the total number of Golgi cisternae per stack did not change
significantly during auxin deprivation, the average number of
trans-type cisternae per stack increased from 1.9 for control cells, to 2.4 for 1.5-d auxin-deprived cells, to 3.3 for 4-d
auxin-deprived cells (Fig. 8). ANOVA of
the number of trans-type cisternae in the control, 1.5-d,
and 4-d groups showed P < 0.0001 (F value = 40.3), a
significant difference. There was no significant difference in total
number of cisternae between the three groups (P = 0.464, F = 0.772). The total number of Golgi stacks counted was 74 for the control
group, 54 for the 1.5-d group, and 47 for the 4-d group.
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| Figure 8.
Histogram illustrating the
auxin-deprivation-induced increase in the number and proportion of
trans-type cisternae (gray bars) in relation to the
total number of cisternae (stippled bars). The total number of
cisternae did not significantly change over time, but the average
number of trans-type cisternae increased after 1.5 and
4 d of auxin deprivation.
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Golgi stacks of control BY-2 cells also contain small sets of parallel,
3- to 5-nm-diameter, intercisternal fibers sandwiched between their
trans cisternae (and sometimes also between the trans-most medial cisternae [Fig. 6]; see also Staehelin
et al., 1990). In cells grown in auxin-deprived conditions, both the
number of intercisternal fibers and the general staining of these
fibrous domains increased significantly (Fig. 7c).
Changes in the Composition of Secreted Proteins
To characterize biochemically the effects of auxin deprivation on
the secretory pathway of tobacco BY-2 cells, we analyzed changes in the
secreted proteins by SDS-PAGE and by peroxidase-conjugated lectin
western blotting. BY-2 cells deprived of auxin for 3 d secreted
more than twice the amount of protein as control cells (4.2 ± 0.32 µg/mL total protein in control cells versus 8.9 ± 1.8 µg/mL in auxin-deprived cells). Analysis of the secreted proteins by
SDS-PAGE (Fig. 9a) demonstrated further
that auxin deprivation also produced qualitative changes in the profile
of secreted proteins. When samples were normalized for protein
concentration, the SDS gels of the secreted proteins of the control
cultures appeared to have more and darker-staining bands. Bands that
appeared in the control samples that were absent or at reduced levels
in the auxin-deprived samples were found at approximately 7, 18.5, 35 (a doublet in the control sample became a singlet in the auxin-deprived sample), 60, 92, and 134.5 kD. On the other hand, a small number of
bands in the auxin-deprived sample were found at higher levels of
staining than in the control sample, including proteins at 4, 27.2, 52, and >200 kD. Because this last band was very diffuse and barely
entered the gel, it possibly represented a very large glycoprotein, a
covalently cross-linked protein aggregate, or a very basic protein-like
extensin that does not migrate normally on SDS gels.
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| Figure 9.
SDS-PAGE (a) and western blots (b-d) of secreted
material from control (+) and auxin-deprived ( ) BY-2 cells. Molecular
mass in kilodaltons is indicated to the left of the gels. All lanes were loaded with equal amounts of proteins. Western blots were probed
with the lectins concanavalin A (Con A) and UEA-1 and with the
monoclonal antibody JIM 13. b, Concanavalin A, a Man-specific lectin, showed minor changes in a set of N-linked
glycoproteins between 45 and 97.5 kD. c, UEA-1, a Fuc-specific lectin,
showed a major increase in a single, diffuse band between 100 and 200 kD. d, JIM 13, a marker of root-cap differentiation, showed an increase
in staining between 40 and 180 kD. All gel images were digitized.
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Lectin western blots were used to characterize changes in secreted
glycoproteins. Concanavalin A is a Man-specific lectin, making it a
useful marker of N-linked glycoproteins. UEA-1 is a
Fuc-specific lectin that has been used as a marker of N- and O-linked mucin-like glycoproteins (Mitsui et al., 1990).
Concanavalin A faintly stained a number of bands between 45 and 97.5 kD
in both control and auxin-deprived samples (Fig. 9b). Although, on the
whole, the proteins in the control lane were of slightly higher apparent molecular mass than those in the auxin-deprived lane, there
were only a few differences in the banding patterns. The control
samples showed a greater staining of protein bands at 51 and 89 kD,
whereas the auxin-deprived samples showed a greater staining of a band
at 73 kD. UEA-1 stained a single large diffuse band between 100 and 200 kD in both control and auxin-deprived auxin lanes (Fig. 9c). The
auxin-deprived band was broader (extended farther down the gel) than
the control band, suggesting that there was an increase in the
production of a high-molecular-mass, mucin-like glycoprotein.
This band, although very large, still did not coincide with the
high-molecular-mass material at the top of the auxin-deprived polyacrylamide gel.
Analysis of AGPs
To assess total AGP secretion in 3-d-old control and
auxin-deprived cells, rocket-gel electrophoresis with Yariv's
artificial antigen (Yariv et al., 1962) was used. Auxin-deprived cells
secreted twice as much AGP as control cells, as judged by the height of the rockets (Fig. 10). Crossed-gel
electrophoresis of the control AGP samples showed a larger and broader
peak of material that migrated more slowly in the horizontal first
dimension (size separation), and a second, smaller peak that migrated
more quickly (Fig. 11, top). In
contrast, only one broad AGP peak with a slight shoulder on its
faster-migrating side is seen in the auxin-deprived cell sample (Fig.
11, bottom). Whereas this shoulder material seems to align with the
small peak material of the control cell samples, the major peak of the
auxin-deprived AGPs occupies a distinctly different position on the
gels between the two peaks of the control.
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| Figure 10.
Quantitative differences in AGPs from control (+)
and auxin-deprived () cells. Rocket-gel electrophoresis with
-glucosyl Yariv's artificial antigen was used to quantitatively
compare control and auxin-deprived secreted material from BY-2 cells. The auxin-deprived cells secreted twice as much AGP as did the control,
as shown by the height of the rockets.
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| Figure 11.
Qualitative differences in AGPs from control
(top) and auxin-deprived (bottom) cells. Crossed-gel electrophoresis
with -glucosyl Yariv's artificial antigen was used to qualitatively
compare control and auxin-deprived secreted material from BY-2 cells.
The wells in which the samples were loaded are indicated. The samples
were first run in the horizontal direction and then in the vertical direction. The AGPs from the control cells formed two distinct peaks,
whereas those from the auxin-deprived cells formed only one peak. The
peak from the auxin-deprived cells traveled farther in the first
dimension than did the major peak from the control cells.
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Immunoelectron Microscopy with the JIM 13 Antibody
Based on the changes in Golgi morphological features, the altered
staining properties of the contents of the enlarged budding vesicles on
the trans side of the stacks (Figs. 7 and 8), and the
changes in composition of the secretory products (Figs. 9-11), we
postulated that the auxin-deprived growth conditions induced a
retailoring of BY-2 cell Golgi stacks in a manner similar to the
retailoring of the Golgi that occurs during the differentiation of
slime-secreting root-tip cells (Staehelin et al., 1990; Lynch and
Staehelin, 1992). To further substantiate this hypothesis, we examined
the effects of auxin-containing and auxin-deprived growth conditions on
the production of molecules containing the JIM 13 epitope, a marker for
slime-secreting root-cap cells. Figure 12 demonstrates that JIM 13 is a marker
of root-cap differentiation in tobacco cells, since the monoclonal
antibody labeled the peripheral slime-secreting cells of the tobacco
root cap as it does the carrot root cap (Knox et al., 1991). If tobacco
BY-2 cells are related to root-cap cells, then auxin deprivation should
either induce the de novo appearance of molecules containing the JIM 13 epitope or increase the amount of JIM 13 epitope-containing molecules produced.
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| Figure 12.
Immunofluorescence micrograph of a tobacco root
tip stained with the monoclonal antibody JIM 13. Staining was
specifically localized to the peripheral root-cap cells. M, Meristem;
C, columella; and P, peripheral root-cap cells. Bar = 50 µm.
|
|
We probed western blots of secreted material from the control and
auxin-deprived cells with JIM 13 (Fig. 9d). Because the epitope is
normally localized to the outer plasma membrane and cell wall, we
hypothesized that it would also be released into the extracellular
medium. As shown in Figure 9d, both the control and the auxin-deprived
cells produced and secreted JIM 13-epitope-containing molecules, but
the auxin-deprived cells secreted more than the control cells. Although
this increase mimicked the increase in the UEA-1-stained fucosylated
glycoproteins (Fig. 9c), the staining patterns shown in Figure 9, c and
d, indicate that the JIM 13 epitopes are associated with a different
set of proteins than the UEA-1 epitopes. In both the control and
auxin-deprived lanes of Figure 9d, the JIM 13-stained material extended
from 40 to 180 kD. The most darkly staining zone in both the control
and auxin-deprived smears appeared at around 116 kD, and in the
auxin-deprived lane this broad band was both more heavily stained and
extended farther down the gel than the material in the control lane.
We also immunolabeled high-pressure-frozen/freeze-substituted BY-2 cell
sections with JIM 13. A comparison of the immunolabeling patterns of
Golgi stacks of control, 1.5-d auxin-deprived cells, and 4.5-d
auxin-deprived cells showed a progressive increase in JIM 13 labeling
of the Golgi cisternae with the increased time in auxin-deprived growth
conditions (Fig. 13). In the 4.5-d
cells, the number of Golgi-associated molecules that labeled with JIM 13 was 4-fold higher than in the control cells. ANOVA showed a significant difference between the control, 1.5-d, and 4.5-d groups (P = 0.0006, F = 8.682).
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| Figure 13.
Immunogold labeling of BY-2-cell Golgi stacks
with JIM 13. Immunoelectron micrographs of control (a) and
auxin-deprived (b) BY-2 cells showed an increase in the number of gold
particles over the Golgi stacks and surrounding cytoplasm in the
auxin-deprived cells. c, Histogram illustrating that gold-particle
labeling over the Golgi stacks (gray bars) increased from an average of
1.4 particles in control cells, to 3.7 particles in 1.5-d
auxin-deprived cells, to 6.0 particles in 4-d auxin-deprived cells.
Gold-particle labeling over the surrounding cytoplasm (stippled bars)
increased from 0.22 particle in control cells, to 0.40 particle in
1.5-d auxin-deprived cells, to 0.54 particle in 4-d auxin-deprived
cells. Bars = 0.2 µm.
|
|
A count of gold particles over an area of cytoplasm equivalent to the
surface area of an average Golgi stack cross-section (0.14 µm2) also showed a 2-fold increase. Because the
small secretory vesicles cannot always be adequately resolved
structurally in the samples processed for immunolabeling, they are most
likely responsible for this increase in cytosolic immunogold staining.
As with the Golgi labeling, there was a significant difference between
the control and auxin-deprived values (P < 0.01). The cell walls
of both control and auxin-deprived cells exhibited quite heavy gold labeling, but the labeling density was not statistically different between the two types of cells (data not shown). Most likely, the cell
walls have a limited capacity to bind JIM 13-epitope-containing molecules, which permits the excess molecules to leach into the growth
medium.
| |
DISCUSSION |
Auxin Deprivation Causes Tobacco BY-2 Cells to Become
Physiologically Synchronized
The goal of this study was to develop a method for inducing plant
suspension-cultured cells to differentiate synchronously in a manner
that involved a structural and functional retailoring of the Golgi
apparatus, analogous to what occurs in plant tissues during normal
development. To this end we have demonstrated that tobacco BY-2 cells
can be induced to differentiate synchronously by transferring them to
an auxin-free growth medium. Auxin deprivation blocked cell division,
as seen by a drop in mitotic index, and induced cell elongation, as
shown by a doubling of median cell length (Figs. 1-4). Similar
developmental responses, a cessation of cell division in G1 (Nishi et
al., 1977), and an increase in cell length (Lloyd et al., 1980;
Komamine and Kawahara, 1991), were found previously in auxin-deprived
suspension-cultured carrot cells.
The principal evidence in support of the hypothesis that auxin
deprivation induces tobacco BY-2 cells to undergo a tissue-specific type of differentiation came from changes in the AGPs produced by these
cells. AGPs are general markers of tissue differentiation (van Holst
and Clarke, 1986). In pea flowers, for example, specific AGPs are
excluded from the reproductive cells (Pennell and Roberts, 1990), and
in carrot roots, specific AGPs mark the development of such areas as
the stele and the root cap (Knox and Roberts, 1989). In maize
seedlings, AGPs demonstrate different temporal and spatial
expression patterns in cells designated for apoptosis (Schindler et
al., 1995). In this study auxin-deprived tobacco BY-2 cells not only
secreted greater quantities of AGP, but also produced a qualitatively
different type of AGP, as shown by rocket- and crossed-gel
electrophoresis (Figs. 11 and 12).
We have yet to determine whether these biochemical changes are
caused by an alteration in the protein or polysaccharide portion of the
AGPs. However, this qualitative change in AGP secretion is indicative
of a differentiation into a new tissue type (van Holst and Clarke,
1986), and suggests that the auxin-deprived cells do undergo a
developmental switch. Therefore, auxin deprivation of BY-2 cells
appears to be a suitable method for producing large quantities of
physiologically synchronized cells. Auxin-deprived cells fulfill the
general criteria of tissue differentiation (Taiz and Zeiger, 1991):
they are metabolically, structurally, and functionally distinct from
control cells. Auxin-deprived cells may therefore be exploited for
studying the retailoring of the Golgi apparatus during differentiation.
Auxin Deprivation Induces Changes in Golgi Stack Distribution and
Architecture
One of the earliest morphological changes in the Golgi apparatus
of auxin-deprived cells is a clustering of the stacks in the vicinity
of the nucleus (Fig. 5b). Golgi stack clustering also occurs in plant
cells exposed to the secretion-blocking drug brefeldin A
(Satiat-Jeunemaitre and Hawes, 1992; Driouich et al., 1993) and the
microfilament-depolymerizing drug cytochalasin D (Satiat-Jeunemaitre et
al., 1996). Most likely, this clustering is caused by the loss of
cytoplasmic streaming and by changes in the Golgi matrix, the
ribosome-excluding filamentous matrix that surrounds each Golgi stack
and its TGN (Staehelin and Moore, 1995). Because brefeldin A and
cytochalasin D induce clustering of the Golgi stacks by different
mechanisms (Satiat-Jeunemaitre et al., 1996), it has yet to be
determined if auxin deprivation alters Golgi stack distribution by
affecting the cytoskeleton, by affecting secretion, or by a combination
of both. Antibody staining against the microfilament cytoskeleton in
control and auxin-deprived cells is needed to further investigate the
effects of auxin deprivation on Golgi stacks.
Auxin deprivation also induced a number of morphological changes in the
BY-2 Golgi stacks, some of which are diagrammed in Figure
14. Of these, the most important would
appear to be the increase in the number and proportion of
trans-Golgi cisternae, a doubling of the diameter (which
corresponds to an 8-fold increase in volume) of the darkly staining
vesicles that bud from the margins of the trans cisternae,
and the loss of a distinct TGN system. Although these types of Golgi
changes do not resemble those found in a number of differentiated
tissues such as tapetal (Steer, 1977), pollen (Hess, 1993), or papillar
cells (Kishi-Nishizawa et al., 1990), they do resemble those that
accompany the retailoring of Golgi stacks in polysaccharide
slime-secreting cells. Golgi stacks with hypertrophied margins and
large, densely staining vesicles have been observed in the
slime-secreting alga Micrasterias denticulata (Meindl et
al., 1992), in pollen tubes of Lilium longiflorum (Lancelle
and Hepler, 1992), and in the slime-secreting epidermal and peripheral
root-cap cells of maize, tobacco, and Arabidopsis (Whaley et al., 1959;
Mollenhauer et al., 1961; Staehelin et al., 1990). In the latter three,
Golgi stacks have also been reported to contain increased numbers of
trans cisternae and TGN (Staehelin et al., 1990), and to
occasionally be clustered together near the nucleus (Whaley et al.,
1959). Taken together, the observed changes in Golgi stack architecture
of auxin-deprived BY-2 cells closely parallel the changes in Golgi
stack morphological features associated with the development of
slime-secreting cells.
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| Figure 14.
Model of the morphological changes in the Golgi
apparatus during auxin deprivation. Schematic diagrams of control and
auxin-deprived Golgi are shown with the cis, medial, and
trans cisternae, and the TGN. Auxin deprivation induces
a doubling in diameter of the budding vesicles around the cisternal
margins (A), an increase in densely staining vesicles beyond the
trans cisternae (B), an increase in the width and
staining of intercisternal filaments (C), an increase in the number and
proportion of trans cisternae (D), an increase in the
staining of the medial and trans cisternae (E), and a
decrease in staining of the cis cisternae.
|
|
Auxin-Deprived Tobacco BY-2 Cells Resemble Slime-Secreting
Root-Cap Cells
The BY-2 cell line was developed from a callus grown on a cultured
tobacco seed embryo (Nagata et al., 1992), so the precise tissue of
origin of the BY-2 cells is not known. Based on our findings to date,
however, it is possible that the BY-2 cells originated from
meristematic root cells. As discussed in the preceding sections, auxin
deprivation causes BY-2 cells to undergo morphological changes that
resemble those seen during the differentiation of polysaccharide
slime-secreting root-cap cells.
Root-cap cells are derived from the distal meristem in the root tip. As
the meristematic cells divide, cells are displaced outward in files and
differentiate first into the gravity-sensing columella cells and then
into the slime-secreting cells (Whaley et al., 1959; Staehelin et al.,
1990). The polysaccharide slime, or mucigel, produced by the
differentiated peripheral cells has been studied most extensively in
root tips from maize seedlings. Mucigel consists of hydrophilic
polysaccharides and swells in contact with water (Samtsevich, 1965).
The average molecular mass of the slime polymers produced by maize cv
SX-17 was found to be 2 × 106 kD by
exclusion chromatography, ultracentrifugal analysis, and the relative
viscosity of ethanol-precipitated material (Paul et al., 1975, as cited
by Rougier, 1981).
Mucigel molecules are synthesized by Golgi-associated enzymes of the
peripheral root-cap cells (for review, see Rougier, 1981). They contain
Glc, Gal, Xyl, Ara, and uronic acid (Jones and Morré, 1973; Moody
et al., 1988), and studies with radioactive precursors and fluorescent
lectins have also shown a high Fuc content (Paull and Jones, 1975;
Wright, 1975; Piché et al., 1985). Some of the glycoproteins in
the mucigel fraction possess polysaccharide side chains that are
attached to Thr residues via Xyl (Green and Northcote, 1978; Moody et
al., 1988).
Juniper and Roberts (1966) suggested that the appearance of
hypertrophied dictyosomes may be the result of a sudden increase in
carbohydrate supply from the breakdown of starch granules. This seems
likely because the mucigel is mainly composed of polysaccharides and
has a pectin-like composition (Wright and Northcote, 1974, 1976;
Wright, 1975). The mucigel molecule has PGA interspersed with regions
of Glc material and has a central -(1,4)-glucan core. It is possible
that this central core makes the molecule fibrillar, whereas an outer
matrix makes it hydrophilic.
If auxin deprivation induces a slime-secreting-cell-like
differentiation, as is suggested by the similarities in Golgi
morphological characteristics, one would expect to see an increase in
the secretion of fucosylated products and high-molecular-mass molecules
in auxin-deprived cells. As described below, both of these were found.
Auxin Deprivation Affects Protein Secretion
The proliferation of large, densely staining Golgi buds and
secretory vesicles in auxin-deprived BY-2 cells appears to reflect a
biochemical change in cell metabolism and secretion. Auxin deprivation induces an increase in total secreted protein and, as shown by SDS-PAGE
in Figure 9a, induces differences in the protein profile of secreted
material from control and auxin-deprived BY-2 cells. Although equal
concentrations of protein were loaded on all of the lanes of the
polyacrylamide gels (shown in Fig. 9), the intensity of the
silver-stained bands (Fig. 9a) appears more consistent with the control
lane having more protein. Furthermore, more bands appeared in the
control lane than in the auxin-deprived lane. This discrepancy is most
likely caused by a lack of reactivity of certain glycoproteins to the
silver stain, as shown by the fact that the UEA-1-stained,
Fuc-containing, high-molecular-mass bands (Fig. 9c) have no
counterparts in the silver-stained gels (Fig. 9a)
A major, slightly smeared band is evident at the very top of the
auxin-deprived lane in the silver-stained gel (Fig. 9a). The smeared
nature of this band suggests that it might also be a
high-molecular-mass glycoprotein. Alternatively, this material could be
a highly basic glycoprotein, such as extensin, that does not migrate
normally in conventional SDS gels (Memelink et al., 1993). If, as
hypothesized, auxin deprivation mimics root-cap development, this
material may be related to the mucigel secreted by the root-cap cells.
Anti-Fuc lectin western-blot analysis showed a major increase in a
large, Fuc-containing protein (100-200 kD) in auxin-deprived cells
(Fig. 9c). What was striking about this blot was that this very broad
band was the only band to be recognized by UEA-1. Because Fuc is a
large component of root-cap slime, this finding provides more evidence
for a relationship between auxin-deprived cells and
root-cap-slime-secreting cells. The Fuc band migrated considerably farther into the gel than the band at the top of the gel described in
the preceding paragraph, so it is unlikely to be part of the same
molecule. These two bands likely represent separate components of the
slime.
Auxin Deprivation Induces an Increase in Secretion of Molecules
Containing the JIM 13 Epitope
The JIM 13 monoclonal antibody was raised against AGP2 from
suspension-cultured carrot root cells by Knox et al. (1991), and predominantly labels the outer surface of the plasma membrane. In
carrot roots the antibody labels the epidermis and the future xylem in
the upper root and the peripheral slime-secreting cells in the root
cap. In tobacco root tips we found low staining in the future xylem but
very high staining in the peripheral root-cap and epidermal cells (Fig.
12). Therefore, we decided to use the antibody as a marker of
slime-secreting-cell development.
Western blots against secreted protein from control and auxin-deprived
BY-2 cells showed both a quantitative and a qualitative change in the
JIM 13 AGP epitope (Fig. 9d). When samples were normalized for total
protein concentration, an increase in JIM 13 staining was seen in the
auxin-deprived lane. Most of the staining in both control and
auxin-deprived cells was found in a diffuse band around 116 kD,
slightly higher (slower migrating) than for carrot
suspension-cultured-cell preparations (Knox et al., 1991). The major
JIM 13 band was slightly lower in the auxin-deprived lane than in the
control lane, suggesting a qualitative shift in the epitope. Because
JIM 13 recognizes a polysaccharide group of an AGP that contains up to
95% carbohydrate (Pennell et al., 1989; Knox et al., 1991), this shift
most likely represents an alteration in the polysaccharide content
rather than in the protein backbone. This shift in size is therefore
most likely a Golgi-mediated posttranslational event rather than a
translational event.
Because the JIM 13 band in both lanes migrated significantly faster
than the glycoprotein(s) detected by UEA-1 staining, it is unlikely
that the anti-Fuc lectin labeled the JIM 13-epitope-containing molecules. The Fuc-containing molecules in the polysaccharide slime are
most likely a separate component from the JIM 13 AGP.
It is not known how much of the total secreted AGP is represented by
the JIM 13 epitope, but a second, more lightly staining smear in the
auxin-containing lane appeared farther down the gel, at around 70 kD.
Because the crossed-gel electrophoresis also showed two peaks of AGP
for the auxin-containing cells and one major peak (with a shoulder) for
the auxin-deprived cells, the JIM 13 epitope may constitute a large
percentage of the total AGP.
McCann et al. (1993) found that both control and auxin-deprived carrot
suspension-cultured cells had large concentrations of JIM 13 epitope in
the cell wall. We were also unable to detect significant differences in
JIM 13 immunogold particle accumulation between control and
auxin-deprived cells (data not shown). Because this appears counter to
the increase in JIM 13 epitope we found in the secreted material from
auxin-deprived BY-2 cells, we needed to determine whether the increase
in secreted JIM 13 represented an actual increase in the production of
the epitope or simply a change in the release of the epitope to the
extracellular medium.
Immunoelectron microscopy with the JIM 13 antibody indicates that the
epitope is synthesized in the Golgi apparatus (Fig. 13) and that the
increase in JIM 13 staining on the western blots of the secreted
proteins (Fig. 9d) is attributable to an increased production of
molecules with JIM 13 epitopes in the Golgi apparatus. Because the JIM
13-epitope-containing molecules have to pass through the cell wall to
reach the extracellular medium, the lack of change of JIM 13 staining
in the cell walls of the control and auxin-deprived BY-2 cells suggests
that the cell wall-binding capacity for JIM 13-epitope-containing
molecules is already saturated in the control cells. Therefore, no
significant difference in gold-particle concentration is seen in the
walls, because as epitope production increases in the Golgi apparatus,
epitope secretion increases into the extracellular medium.
In tobacco root caps the meristematic and young columella cells produce
no detectable JIM 13 epitope (Fig. 12). The fact that the control BY-2
cells already synthesize some JIM 13-epitope-containing molecules
indicates that the control cells are not equivalent to root tip
meristematic cells or young columella cells, but rather are akin to
late-stage columella cells. Auxin deprivation therefore appears to
induce in tobacco BY-2 cells a series of morphological changes in the
Golgi apparatus and concomitant changes in the nature of secretory
products that mimic the final developmental changes in the formation of
slime-secreting cells from late-stage columella cells.
Future Studies
Auxin deprivation induces suspension-cultured BY-2 cells to
differentiate in a manner that resembles the development of
root-cap-slime-secreting cells. This has been demonstrated through
morphological changes in the Golgi stacks, qualitative and quantitative
changes in the secretion of general tissue differentiation markers, and
an increase in Golgi staining of the specific root-cap-slime-cell
marker JIM 13.
Further studies of auxin-deprived BY-2 cells may also lead to the
discovery of new markers for root-cap differentiation. For example, we
plan to investigate changes in secreted pectic polysaccharides, since
the ratio of methylesterified to unesterified polygalacturonic acid
residues has been shown to be related to peripheral root-cap-cell development (Hawes and Lin, 1990; Lynch and Staehelin, 1992; Stephenson and Hawes, 1994).
Finally, auxin deprivation opens up new doors for biochemical studies
of Golgi differentiation. As discussed in the introduction, it has been
nearly impossible in the past to purify Golgi stacks of a single tissue
type. By using auxin-deprived suspension-cultured cells, we can isolate
much larger quantities of Golgi membranes and study changes in the
integral membrane proteins involved in the morphological and functional
differentiation of the plant Golgi apparatus.
| |
FOOTNOTES |
1
This work was supported by a National Institutes
of Health grant (no. 18639 to L.A.S.).
2
Present address: Plant Biology Department,
University of Minnesota, 220 Biological Sciences Center, 1445 Gortner
Avenue, St. Paul, MN 55108.
3
Present address: Department of Biochemistry and
Biophysics, University of California, San Francisco, CA 94143-0554.
*
Corresponding author; e-mail zwinicur{at}biosci.cbs.umn.edu; fax
1-612-625-1738.
Received January 9, 1998;
accepted February 23, 1998.
| |
ABBREVIATIONS |
Abbreviations:
AGP, arabinogalactan protein.
ANOVA, analysis of
variance.
DAPI, 4
,6-diamidino-phenylindole.
TGN, trans-Golgi network.
| |
ACKNOWLEDGMENTS |
We gratefully acknowledge Drs. Paul Knox and Keith Roberts (John
Innes Centre, Norwich, UK) for their generous gift of the JIM 13 monoclonal antibody. Thanks go to Dr. Malcolm Bennett at the University
of Warwick (UK) and to Janet Meehl at the University of Colorado for
their comments and suggestions on the manuscript. Thanks also go to Dr.
Tom Giddings for help with the electron microscopy, and to Diane Lorenz
for help with the preparation of figures.
| |
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Top
Abstract
Introduction