Download From signal to form: aspects of the cytoskeleton

Journal of Experimental Botany, Vol. 48, No. 316, pp. 1881-1896, November 1997
Journal of
Experimental
Botany
REVIEW ARTICLE
From signal to form: aspects of the cytoskeleton-plasma
membrane-cell wall continuum in root hair tips
Deborah D. Miller, Norbert C.A. de Ruijter and Anne Mie C. Emons1
Department of Plant Cytology and Morphology, Wageningen Agricultural University, Arboretumlaan 4,
NL-6703 BD Wageningen, The Netherlands
Received 13 June 1997; Accepted 14 August 1997
Abstract
Introduction
Root hairs are excellent cells for the study of the
exocytotic process that leads to growth in higher
plants, because the exocytotic event takes place locally and because the cells are directly accessible for
signals, drugs, fixatives, microinjection, and microscopic observation. Well-characterized lipochitooligosaccharides, signal molecules excreted by
Rhizobium bacteria, induce root hair growth which can
be recorded microscopically in a root hair deformation
assay developed for Vicia sativa L. Root hair deformation is a morphogenetic process involving swelling of
the hair tip and subsequent new hair outgrowth from
that swelling. This response to the signal occurs at a
specific developmental stage, namely when hairs are
terminating growth. Thus, since polar growth can be
triggered intentionally, the system allows the study of
growth phenomena in higher plants at the cellular
level. Furthermore, important advances are being
made with molecular genetics that will allow the unravelling of the signal transduction pathways in root hair
morphogenesis leading to growth. This paper first discusses cytological phenomena involved in the process
of polar growth, such as cytoplasmic polarity, cytoplasmic streaming and the organization of actin filaments, the location of a spectrin-like antigen, the
distribution of intracellular calcium, cortical microtubules and cell wall texture, endocytosis by means of
coated pits, and physical aspects of the incorporation
of exocytotic vesicles into the plasma membrane. In
the second part, changes are discussed that occur in
some of these phenomena when growth is influenced
by growth regulators and mutations.
Plant form is the summation of the events involved in cell
morphogenesis and cell patterning. After cell growth the
form of a plant cell is maintained by its cell walls, but
during cell growth these walls are flexible and the genesis
of a particular shape depends on the site of expansion
(Roberts, 1994). Under physiological conditions, cell
expansion is linked to the process of wall synthesis. Since
the maximal elastic stretching of the plant plasma membrane is about 2% of its surface area (Wolfe and Steponkus,
1981, 1983), the expansion of the plasma membrane
requires insertion of new membrane. Furthermore, since
expanding walls do not become thinner, cell growth
requires exocytosis of wall materials (Roberts, 1994). Thus,
in the case of cell expansion, the exocytotic vesicle is the
unit of plant cell growth. Though natural cell growth does
not occur without exocytosis, it should be noted that
exocytosis in plant cells may take place without cell
expansion. This phenomenon happens in cases where the
exocytosed material is excreted outside the cell wall, for
instance nectar or the slime of root cap cells, as well as
during secondary wall deposition when cell wall deposition
occurs without cell expansion. Recent reviews by Battey
and Blackbourn (1993) and Battey el al. (1996) provide
insights into exocytosis in plant cells.
The location of exocytosis determines where, and for
cells in a tissue, in which direction, growth will take place.
One can discriminate between intercalary-, isodiametric-,
tip growth, and combinations of these growth types,
depending on which cell facets expand. The most common
type is intercalary growth, occurring in elongating cells,
such as the cells of the elongation zone of a root. In these
cells exocytosis occurs in all longitudinal cell facets.
However, since cortex cells also become thicker during
growth in length, some exocytosis should occur in the
Key words: Cytoskeleton, exocytosis, plant growth regulators, Nodulation factors, root hair, tip growth.
' To whom correspondence should be addressed. Fax: +31 317 485005. E-mail: [email protected]
© Oxford University Press 1997
1882
Miller et al.
transverse cell facets as well. In isodiametric growth
exocytosis occurs in all directions, whereas in tip growing
plant cells exocytosis occurs at one side of the cell and is
the reason why these cells are called tip growing or polar
growing cells. The well-known examples of tip growing
cells of higher plants are pollen tubes (Schnepf, 1986)
and root hairs (Derksen and Emons, 1990).
Root hairs are involved in the uptake of nutrients and
water (Abeles, 1992; Peterson and Farquhar, 1996) and
are involved in the interaction between plants and nitrogen-fixing bacteria (Mylona et al., 1995; Geurts and
Franssen, 1996). The fact that these cell types may grow
fast, that exocytosis takes place locally and that deformation by Rhizobium signals is specific and occurs only in a
predictable developmental stage makes legume root hairs
an attractive system for the study of exocytosis, plant
morphogenesis, and signal transduction at the cellular
level. Furthermore, root hairs are easily accessible for
treatments and cell biological analyses, and growing and
full grown hairs of the same root may be compared. Since
these cells are part of the plant body, data on fundamental
processes gained from them will, in principle, be relevant
for other higher plant cells.
Characteristics of cells with tip growth
Cell polarity, vesicle-rich region
Polarity is the quality or condition inherent in a body
that exhibits opposite properties in opposite parts or
directions; in biological terminology it is the observed
axial differentiation of an organism or tissue or cell into
parts with distinctive properties or form. A root hair with
the mechanism for cell elongation completely localized at
one site on the cell periphery is a typical polar cell. The
appearance of the polarized cyto-architecture of a root
hair can be seen in the light microscope with Nomarski
optics (Fig. 1). An emerging hair, which Dolan et al.
(1994) call a bulge, does not exhibit an apparent vesiclerich region, the region at the extreme tip of a polar
growing cell where only vesicles are located, in the light
microscope (Fig. la), and so this root hair initiation
phenomenon has to be studied in more detail with the
electron microscope. It is not known whether the initial
stage of pollen tube growth, a comparable system, is
lacking a vesicle-rich region. Growing root hairs (Fig.lb)
and growing pollen tubes have a vesicle-rich region at the
tip. It has been known for a long time that this vesiclerich region consists of Golgi vesicles both in pollen tubes
(Sassen, 1964; Rosen et al., 1964; confirmed with the
freeze substitution technique by Lancelle and Hepler,
1992) and in root hairs (Bonnett and Newcomb, 1966;
confirmed with the freeze substitution technique by
Emons, 1987; Ridge, 1988; Sherrier and Van den Bosch,
1994; Galway et al., 1997). The central vacuole in growing
Fig. 1. Differential Interference Contrast (DIC) images of developmental stages of Vicia saliva root hairs. (A) Emerging hair bulging
from a root epidermal cell; (B) growing hair with a vesicle-rich region
at the tip and an area with dense cytoplasm behind the tip; (C) hair
that is terminating growth with small vacuoles close to the tip; (D) fullgrown hair with central vacuole up to the tip and peripheral cytoplasm.
Bar= 1
root hairs is always located in the basal part of the cell
and the nucleus lies alongside the apical part of the large
vacuole (Galway et al., 1997) and migrates at the same
pace as the growing tip (Ridge, 1992). When root hairs
stop growing, the vesicle-rich region initially becomes
smaller and the main vacuole moves closer to the tip
(Fig. lc). The organelle zonation in root hairs has not
been studied in detail as in pollen tubes (Derksen et al.,
1995a).
Cytoplasmic streaming and arrangement ofactin filaments
Many tip growing cells exhibit a specific type of cytoplasmic streaming called reverse-fountain streaming, a
term introduced by Iwanami (1956) for pollen tubes. This
type of streaming allows transports of Golgi vesicles from
their subapical point of origin to the vesicle-rich region
at the hair tip (root hairs: Emons, 1987; Shimmen et al.,
1995; pollen tubes: Derksen et al., 1995a; Derksen, 1996).
The most characteristic feature of this type of streaming
is that the flow of cytoplasm does not reach the cell apex,
but reverses direction when it reaches the vesicle-rich
region. This type of streaming is particularly evident in
Roof hair tip growth 1883
broad root hairs such as those of Hydrocharis (Shimmen
et al., 1995) in which the flow of cytoplasm is upwards
at the cell borders and downwards in the cell centre. In
relatively thin hairs such as Vicia saliva L. (De Ruijter
et al., unpublished observations) the downward flow, as
well as the upward flow, may be close to the plasma
membrane. Nevertheless, it is catagorized as reversefountain streaming and not circular streaming as seen in
cells with cytoplasmic strands because the flow never
reaches the plasma membrane at the utmost tip of the hair.
In full-grown hairs there is a completely different type
of cytoplasmic streaming compared to growing hairs. The
flow does not reverse before it reaches the hair tip as in
growing cells, but flows through the apical hemisphere in
a thin layer of cytoplasm between the large central vacuole
and the plasma membrane. This type of cytoplasmic
streaming has been called rotation streaming (Iwanami,
1956). Cytoplasmic polarity is lost (Fig. Id) in these hairs.
It is known that cytoplasmic movement in plant cells
is caused by the actin-myosin complex (for a review see
Mascarenhas, 1993). Since myosins travel along actin
filaments from the pointed (minus) end to the barbed
(plus) end, Shimmen et al. (1995) surmise that the actin
filaments in the cytoplasmic strands in the cell centre of
Hydrocharis root hairs should be arranged with the
pointed ends toward the cell tip since the flow in these
strands is toward the base of the hair. The filaments in
the peripheral strands would have the opposite polarity
since the flow in these strands is towards the tip. Yet,
these statements have not been confirmed directly.
Following the same line of thought, it can be deduced
that the orientation of actin filaments in the cytoplasmic
strands in growing and full-grown hairs is different. This
orientation has to be proven by, for instance, heavy
meromyosin decoration of individual actin filaments in
the bundles.
The rate of streaming in root hairs within one species
depends on hair maturity (Emons, 1987; Sattelmacher
et al., 1993) and growth rate (Soran and Lazar-Keul,
1978). Video microscopy was utilized in root hairs to
track particles in the stream of cytoplasm indicating that
particles move along the long axis of the hair in the
subapical and basal portions of the hair, yet significantly
less movement (Seagull and Heath, 1980) or no streaming
was observed at the extreme tip (Emons, 1987). When
Hydrocharis root hairs were treated with cytochalasin B
or mycalolide B, cytoplasmic streaming ceased and
strands were destroyed (Shimmen et al., 1995). This study
shows that actin filaments act as tracks along which
cytoplasmic streaming occurs and act as a cytoskeleton
in maintaining the transvacuolar strands.
In tip growing cells the Golgi bodies lie subapically
(Fig. 2), but their vesicles are inserted into the plasma
membrane at the cell apex where they deliver their contents. Therefore, the vesicles have to be transported from
the Golgi bodies to the base of the vesicle-rich region and
then onwards to the extreme tip. Transport to the base
of the vesicle-rich region most likely occurs by cytoplasmic
streaming, but how vesicles move within the vesicle-rich
region is unclear. When root hairs of Equisetum hyemale
and Limnobium stoloniferum were rapidly frozen by plunging in liquid propane and freeze-substituted for electron
microscopy, microfilaments were not detected in the vesicle-rich region between the Golgi vesicles, and staining
with rhodamine phalloidin showed no actin filaments in
the hair tip (Emons, 1987). Also, rapid-freeze, freezesubstitution of Vicia hirsuta root hairs by Ridge (1988,
1990) indicated an absence of actin filaments in the apical
dome. Seagull and Heath (1979) have identified two
populations of actin filaments in radish root hairs by
electron microscopy: filament bundles throughout the
cytoplasm except at the extreme tip, and single cortical
filaments associated with microtubules again with the
exception of the tip. With electron microscopy some fine
filaments were visible adjacent to microtubules in the
cortical cytoplasm of the hair dome in root hairs of
Equisetum hyemale (Emons, 1987), but the nature of
these filaments has not been studied with immunogold
labelling. Since no cytoplasmic streaming was observed
with video microscopy in the vesicle-rich region, it was
suggested that the fine filaments alongside the cortical
microtubules contribute to directed targeting of vesicles
toward the site of insertion (Emons, 1987). High resolution video microscopy of Lilium pollen tubes shows the
tip region filled with vigorously vibrating units, presumably vesicles, that display a chaotic agitation with a high
frequency, but a low amplitude, suggesting that migration
of vesicles to the plasma membrane is not a smooth flow
of particles but a very turbulent process (Pierson et al.,
1990). At present the force responsible for this process is
unknown. A mobile vesicle supply centre such as the
Spitzenkorper in fungi (reviewed by Harold, 1997) has
never been reported for root hairs.
A more physical, passive phenomenon could also lead
to transport of vesicles to the tip. If Golgi vesicles are
attached to actin filaments, they will be brought from the
Golgi bodies to the vesicle-rich region by the tipward
cytoplasmic streaming. If they uncouple from the filaments at the vesicle-rich region, and if vesicles are constantly incorporated into the plasma membrane at the
other side of the vesicle-rich region, there may be no need
for an active transport mechanism within this region.
An absence of actin filaments within the apical dome
of tip growing cells has been observed in several types of
tip growing cells. Jackson and Heath (1993) found a
region lacking actin filaments at the very apex of growing
Saprolegnia hyphae. Actin filaments in living polarized
plant cells have been studied only in pollen tubes of
Lilium by Miller et al. (1996). Thick actin bundles are
longitudinally oriented in the cytoplasm, but they taper
1884 Miller etal
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Fig. 2. Electron micrograph of a growing root hair of Equisetum hvemale showing Golgi vesicles in the hemisphere of the tip and Golgi bodies
(GB) subapically. B a r = l /xm.
towards the edge of the pollen tube when nearing the
vesicle-rich region and are absent from the vesicle-rich
region proper (Miller et al., 1996). Treatment with
caffeine, a substance that greatly diminishes the calcium
influx and abolishes the tip-focused calcium gradient
(Pierson et al., 1996), causes the actin filaments to redistribute in such a way that they are closer to the tip,
thereby shortening the vesicle-rich region. When caffeine
is removed, the vesicle-rich region re-establishes its normal
size simultaneously with the removal of actin filaments
from this area (Miller et al., 1996). For further discussion
of pollen tube actin, see Miller et al. (1996). Since pollen
tubes and root hairs are both polar growing cells with
similar morphology, organelle distribution and cytoplasmic streaming patterns, it would seem logical that the
actin distribution within growing root hairs is similar to
that of a pollen tube. Future experiments should address
this issue.
Spectrin-like antigen
Animal spectrins are multifunctional molecules that
belong to a family of proteins including actin binding
protein 1, a-actinin, dystrophin, and fimbrin. These proteins have many binding sites with at least two actin
filament binding sites, and binding sites for calcium and
calmodulin (Hartwig, 1994). Spectrin and its associated
proteins in red blood cells are now thought to provide
organizational stability to a cell by controlling integral
membrane protein distribution (Bennett and Gilligan,
1993; Devarajan and Morrow, 1996). Based on data from
adrenal chromaffin cell secretion Hays et al. (1994) suggest that an increase of cytosolic calcium concentration
([Ca 2+ ] c ) dissociates actin from spectrin thereby allowing
vesicle fusion.
Immunoblot analysis of young tomato leaves detected
a 220 kDa protein labelled with human erythrocytespectrin (Michaud et al., 1991), and Faraday and
Spanswick (1993) detected a 230 kDa protein with antispectrin in the purified plasma membrane fraction of rice
roots. In addition, De Ruijter and Emons (1993) detected
spectrin-like epitopes in several tissues of maize and
carrot, predominantly at the plasma membrane of cells
growing along their whole length. This spectrin-like epitope is also localized at the tip of growing root hairs
(Fig. 3), which is comparable to its localization at the
tips of tobacco pollen tubes (Derksen et al., 19956) and
the tips of growing hyphae of Saprolegnia ferax
(Kaminskyj and Heath, 1995). Even though the nature
of the plant epitope found in root hairs is still not known,
its location suggests a role in exocytosis.
Calcium gradient
External Ca 2 + ions at the appropriate concentration and
a Ca 2 + influx are required to elicit exocytosis in
Arabidopsis root hairs (Schiefelbein et al., 1992). These
authors observed a net influx of Ca 2 + at the tips of
growing root hairs with a calcium selective vibrating
probe, whereas no influx was found at the sides of the
hair nor into non-growing hairs. Selective vibrating probe
analysis indicated Ca 2 + currents only in the tips of
growing Sinapis root hairs (Herrmann and Felle, 1995;
Felle and Hepler, 1997), which responded to changes in
external concentrations of calcium by transient growth
differences rather than an altered steady-state growth rate
(Herrmann and Felle, 1995). The [C&1+\. increased in the
apical area in the presence of increased external calcium
and decreased when the external calcium concentration
was lowered (Felle and Hepler, 1997). In the latter study
non-growing hairs responded to changes in external calcium as well, but the [Ca 2+ ] c increase was uniform
Roof hair tip growth 1885
Fig. 4. Confocal ratio image of the median-optical plane of a growing
Vicia sativa root hair loaded with the Ca 2 + indicator Indo-1 AM
showing high [Ca2*],. at the tip (red-orange). The vacuole appears green
due to sequestering of Indo-1 AM. Calibrated [ C a 2 ^ represented in
pseudocolour, ranges from 150 nM at the base (blue-green) of the hair
to 1.8 mM (red-orange) at the growing tip. Bar= 15 ^m.
Fig. 3. (A) DIC image of growing Vicia sativa root hairs after chemical
fixation at the tip; (B) the same cells labelled with anti-spectnn
antibody, showing the spectrin epitope in the root hair tip. Bar= 15 ^m.
throughout the hair. Lew (1991) observed a K + influx at
the tips of living Arabidopsis root hairs. Jones et al.
(1995) used selective vibrating microelectrodes to measure
the magnitude and spatial localization of Ca 2 + , K + and
H + fluxes in growing and non-growing root hairs of
Limnobium stoloniferum. The Ca 2 + and H + inward fluxes
were restricted to tips of growing hairs, while K + influx
was uniform along the length of the whole hair.
Circumstantial evidence for the existence of calcium
channels in plant plasma membranes is based on the
inhibitory effects of calcium channel blockers such as
nifedipine (Reiss and Herth, 1985; Schiefelbein et al.,
1992) and verapamil (Bednarska, 1989). Nifedipine eliminated the Ca 2 + influx and at the same time caused
Arabidopsis root hairs to stop growing (Schiefelbein et al.,
1992). The observed Ca 2 + influx in Sinapis root hairs
could be inhibited by nifedipine (Herrmann and Felle,
1995) and La 3 + , another calcium channel blocker
(Herrmann and Felle, 1995; Felle and Hepler, 1997).
Jones et al. (1995) inhibited Ca 2 + influx by applying
Mg 2 + at various concentrations. Inhibition of 90% of the
Ca 2 + influx by the competing Mg 2 + did not significantly
change the growth rate suggesting that the magnitude of
the calcium flux does not directly determine the growth
rate in Limnobium stoloniferum. Since no outward Ca 2 +
flux was detected in Arabidopsis (Schiefelbein et al., 1992)
or in Sinapis (Herrmann and Felle, 1995), Lommel and
Felle (1997) believe that the large, expanding vacuole is
most likely acting as a Ca 2 + sink.
Using Fura-2 Clarkson et al. (1988) have found an
internal [Ca 2+ ] c tip-to-base gradient in tomato root hairs,
but this result was not consistently seen. In root hairs of
Vicia sativa the [Ca 2+ ] c at the tips of growing and fullgrown hairs were statistically different (De Ruijter et al.,
unpublished results). The tip of a full-grown hair has a
[Ca 2+ ] c similar to the level at the base of the root hair of
100-150 nM. Meanwhile, in a growing tip, a tip-focused
[Ca 2+ ] c was found that can reach up to 2.2 ^.M (Fig. 4).
In root hairs of Sinapis Herrmann and Felle (1995) and
Felle and Hepler (1997) found that the tip-focused [Ca 2+ ] c
levels are approximately three times higher in the tip of
the hair than in the basal part, whereas a pH gradient
was not found within the Ca 2 + gradient. The addition of
La 3+ eliminated the inward flow of Ca 2 + into the tip
of the root hair, while the addition of the plasma membrane H + -ATPase inhibitor, dicyclohexylcarbodiimide
(DCCD), stopped root hair growth but did not abolish
the [Ca2+]<. gradient or the basal calcium level (Felle and
1886 Miller etai.
Hepler, 1997). The authors suggest that an elevated apical
[Ca2+]c may be important for tip growth, while DCCD
inhibition for long time periods may slowly cause the
decline of the gradient and eventually cause the breakdown of the plasma membrane transport processes.
In plant cells, as in animal cells (Clapham, 1995),
exocytosis appears to be dependent on, and triggered by,
a rise in the [Ca2+]c. Ca2+ channels are integral membrane
proteins, which are most likely delivered to the plasma
membrane locally by the fusion of secretory vesicles with
the plasma membrane. Insertion and activation of these
channels at the tip of growing root hairs would lead to
localized influx of Ca2+ and hence polarized growth.
These Ca 2+ channels must be subsequently deactivated
or endocytosed allowing for continued insertion and use
of new Ca 2+ channels. For further discussion of this
matter in pollen tubes, see Steer and Steer (1989) and
Miller et al. (1992).
Cortical microtubules and cell wall microfibrils
In root hairs, microtubules lie mainly in the cortical
cytoplasm next to the plasma membrane in a longitudinal
or helical orientation (Newcomb and Bonnett, 1965;
Seagull and Heath, 1980; Lloyd and Wells, 1985; Ridge,
1988; Emons, 1989; reviewed in Ridge, 1996) and appear
to be continuous with the endoplasmic microtubules
located between the nucleus and tip (Lloyd et al., 1987).
Microtubules have been clearly shown to be in the tips
of root hairs (Lloyd and Wells, 1985; Trass et al., 1985)
and adjacent to the plasma membrane in a random
orientation (Emons, 1989).
Adjacent to primary cell walls that expand, cortical
microtubules are parallel to the last cellulose microfibrils
(Wymer and Lloyd, 1996). Root hair tips are in this
respect no different from other cells. Their tips expand
more or less isodiametrically and in the tip hemisphere
the cortical microtubules as well as the cellulose microfibrils, which are being deposited, lie in all directions
(Emons, 1989). Adjacent to many secondary cell walls
cortical microtubules are not in parallel alignment with
the last cellulose microfibrils deposited in the cell wall
(Emons et al., 1992a). A logical conclusion drawn from
such, often made, observations is that cortical microtubules do not direct the cellulose microfibrils under deposition, but do relate to expansion. They may determine
which walls can expand by directing the Golgi vesicles to
these cell facets. Application of the microtubuledepolymerizing drug colchicine in a concentration range
between 1-10 mM caused widening of the tips of root
hairs of Equisetwn and Raphanus (Emons et al., 1990)
indicating that cortical microtubules may be involved in
determining the width of a tip growing cell.
The tonneau {ton) (Traas et al., 1996) and fass
(McClinton and Sung, 1997) Arabidopsis mutants pro-
duce morphologically normal root hairs. However, the
root cells themselves lack microtubules adjacent to the
plasma membrane in the cortical cytoplasm and indeed
these cells do not grow properly and yield a shorter,
thicker root (TON, Traas et al., 1996; FASS, McClinton
and Sung, 1997). However, Traas et al. (1996) mention
that the microtubule cytoskeleton in the ton root hairs is
normal, which suggests that different genes are involved
in cytoskeletal arrangement in intercalary- and tipgrowing cells.
Site and amount of endocytosis
Endocytosis is the process of internalizing membrane by
sorting out particular intramembrane proteins and their
ligands; it is a manner of making membrane domains
with dedicated function. Neither the composition of the
membrane nor the vesicle content is known in plant cells.
The vesicles are surrounded by a clathrin coat and bud
off as coated pits (Coleman et al., 1988). The subapical
plasma membrane of growing root hairs (Emons and
Traas, 1986; Ridge, 1988), as well as Nicotiana tabacum
L. pollen tubes (Derksen et al., 1995a), is covered with
coated pits. The density of coated pits in growing root
hairs of Equisetum is 2.5-4.5 ^.m"2 and in Raphanus
4.0-8,0 fxm~2 (Emons and Traas, 1986). In full-grown
root hairs the density is only 0.1-0.6 fim"2, showing that
more membrane turnover takes place in growing hairs
than in full-grown ones. Pits in all stages of assembly and
in all stages of pinching off as coated vesicles were
observed (Fig. 5). Derksen et al. (1995a) counted a coated
pit density of 15 fxm~2 at a distance of 6-15 /nm behind
the extreme apex in tobacco pollen tubes. Galway et al.
(1997) report coated pits among secretory vesicles at the
extreme tip of Arabidopsis root hairs. Restricted localization of a plant clathrin heavy chain antibody to the apex
of Lilium longiflorum pollen tubes was seen by Blackbourn
and Jackson (1996). These results suggest that exocytosis
and endocytosis occur near each other in tip growing
plant cells. In animal cells calcium is required for
exocytosis, but is generally not essential for endocytosis
while annexin proteins are implicated in steps involved in
both processes (Gruenberg and Emans, 1993).
Exocytosis
Exocytosis links the inside and the outside of the cell and
therefore it has to be a precisely regulated process. The
following cascade of interactions takes place: budding of
vesicles from the Golgi bodies; translocation and delivery
of vesicles to the plasma membrane; docking and attachment of vesicles at the plasma membrane; priming of
vesicles, i.e. waiting for Ca 2+ ; fusion of vesicles with the
plasma membrane, triggered by Ca 2+ ; pore widening and
discharge of vesicle content (Battey and Blackbourn,
Root hair tip growth
1887
Fig. 5. Electron micrograph of the subapical plasma membrane of a Raphanus sativus root hair with microtubules (MT), coated pits and coated
vesicles, encircled in left part of the micrograph. Bar = 300 nm.
1993). The putative role for synaptobrevin homologue in
plant cell exocytosis has been reviewed by Battey and
Blackbourn (1993) and the function of annexin-membrane interactions has been described by Battey and
Blackbourn (1993), Clark and Roux (1995) and Moss
(1997).
Physical aspects of vesicle insertion
In growing root hairs exocytosis takes place at the hair
tip. The physical aspects of vesicle insertion and content
delivery can be studied in freeze-fracture preparations,
which has been done in Equisetum hyemale root hairs
(Emons, 1985). However, hardly any fractures were
through the plasma membrane of the extreme tip. Vesicle
insertion into the membrane and delivery of their contents
has best been observed in non-embryogenic (Staehelin
and Chapman, 1987) and embryogenic suspension cultures (Emons et al., 19926) of Daucus carota.
From such data the following hypothesis for the course
of events has been extrapolated for plant cells under
turgor pressure (Staehelin and Chapman, 1987; Emons
et al., 19926): (i) an exocytotic vesicle fuses with the
plasma membrane, initially forming a pore (Fig. 6a),
(ii) the vesicle content is delivered into the cell wall, (iii)
the fusion between vesicle and plasma membrane changes
into a slit (Fig. 6b), (iv) the vesicle tips over (Fig. 7) and
(v) the vesicle membrane is incorporated into the plasma
membrane giving rise to bulges, depressions, and horseshoe-shaped configurations in freeze-fracture images
(Fig. 6c, d, e). The latter three are 160±20 nm in diameter
(Emons et al., 19926). Another fracture plane through
the inside of a fusing vesicle (Emons et al., 19926)
(Fig. 6f), confirms the model.
Thus, pores, slits and bulges/depressions/horseshoes
are regarded to be three successive stages. Bulges and
depressions are the same feature seen in the exoplasmic
(Fig. 6b) and protoplasmic (Fig. 6d) fracture faces,
respectively. Bulges/depressions (Fig. 6b, d) and horseshoes (Fig. 6c, e) are not two different developmental
stages but different fracture planes through the vesicle
during insertion into the plasma membrane (Fig. 7).
These data give important insights into the process of
exocytosis in plant cells, but such data have yet to be
collected in tip growing cells.
Calculation of the amount of vesicle insertion
The exocytotic vesicles of Equisetum hyemale root hairs
from freeze-substitution studies have a diameter of 300 nm
(Emons, 1987). Thus, one vesicle inserts 0.28 ^m 2 (4-rrr2)
membrane. The diameter of a growing Equisetum hyemale
root hair is approximately 20 ^m and under greenhouse
conditions its growth speed is 40 f i m h " ' (Emons and
Wolters-Arts, 1983). If this rate is taken into consideration then it means that in 1 h a membrane area with the
dimensions of a tube with a diameter of 20 ^m and a
length of 40 fim is inserted (2500 ^m 2 ). If the above value
of 0.28 fim2 is taken for one vesicle insertion, in 1 h 8900
vesicles will be inserted into the plasma membrane of the
tip. This value is accurate if no endocytosis takes place.
However, during tip growth in root hairs massive
1888 M/7/eretal.
Fig. 6. Exocytotic configurations in freeze-fracture preparations of embryogenic suspension cells of Daucus carola. (A) Exoplasmic Fracture Face
(EF-face) with an early stage of vesicle fusion; (B) EF-face with slit-like fusions (arrow) and bulges; (C) EF-face with bulges and horseshoe-shaped
configurations; (D) Protoplasmic Fracture Face (PF-face) with depressions; (E) PF-face with horseshoe-shaped configurations; (F) EF-face with
fractures through the inside of exocytotic vesicles. Bar = 500/tm.
cytoplasm
Fig. 7. Schematical representation of the formation of (A) bulges in the EF-face of the plasma membrane during vesicle-plasma membrane fusion,
and depressions in the PF-face, if the fracture plane goes through the lower side of the tipped-over vesicle, and (B) horseshoe-shaped configurations
in both membrane faces, if the fracture goes through the plasma membrane.
Roof hair tip growth
endocytosis takes place (Fig. 5; Emons and Traas, 1986).
On the basis of coated pit density and the assumption
that coated pit lifetime is as in other cells (Pearse and
Bretscher, 1981; Tanchak et al., 1984) plasma membrane
turnover in these growing hairs has been calculated to be
20-40 min (Emons and Traas, 1986). Without any growth
taking place, in approximately 30 min this 2500 ixm2 area
of plasma membrane is replaced, which is 5000 ^mh" 1 .
If one vesicle insertion consists of 0.28 ^m2 membrane, it
amounts to 17 800 vesicles h" 1 . In that case, endocytosis
and growth together need to have 26 700 vesicles h" 1 or
445 vesicles min ~1 inserted. Morre and van der Woude
(1974) estimated the rate of vesicle formation required to
sustain growth of Liliitm pollen tubes to be 1000
vesicles min"1. These authors based their estimation on
the assumption that a unit volume of vesicle contents
supplied to the cell surface leads to a unit volume increase
in the growing wall. Based on vesicle accumulation in
Tradescantia pollen tubes after cytochalasin treatment the
estimate was 3000-5000 vesicles min"1 (Picton and Steer,
1983). However, the growth speed of pollen tubes is
considerably higher than that of root hairs, for example,
~ 2 ^.m min" 1 in lily pollen tubes. Assuming that
exocytosis takes place in the whole hair dome, and that
the root hair tip is hemispherical, which is a simplification,
445 vesicles are inserted every min into a membrane area
of 628 fim2 (ATTV2). If all these vesicles would be visible as
exocytotic configurations at the same time, then 0.7
configurations fim"2 would be seen on the plasma membrane of the hair dome. This estimate falls well within
the counted number of exocytotic configurations in
embryogenic (0.57 + 0.13 ^m" 2 ) and fast growing nonembryogenic (1.24+ 1.34 /xm"2) suspension cells of carrot
(Emons et al., 19926). For other methods to measure
exocytosis see the recent review by Battey et al. (1996).
Effects of growth regulators on calcium, and on
the cytoskeleton, plasma membrane and cell wall
continuum
Exogenous application of growth regulators
Plant growth regulator action mediates morphological
change by stimulating changes in the cortical cytoskeletal
elements. Over the last 30 years many investigations have
shown that cortical microtubules lie transverse to the
direction of elongation of a plant cell (Cyr, 1994; Wymer
and Lloyd, 1996; Hush and Overall, 1996), including root
hair tips (Emons, 1989). Light (Nick et al., 1990; Nick
and Schafer, 1991, 1994; Zandomeni and Schopfer, 1993)
and plant growth regulators mediate their effect on the
direction and amount of cell growth by changing the
orientation and number of cortical microtubules (Lloyd
et al., 1996; reviews: Shibaoka, 1991, 1994; Cyr, 1994;
Hush and Overall, 1996; for ethylene see Dolan, 1997).
1889
Auxins (Nick and Schafer, 1994; Millner, 1995; Mayumi
and Shibaoka, 1996; Masucci and Schiefelbein, 1996),
and gibberellins (Shibaoka, 1993, 1994; Uoydetal., 1996)
generally promote a cortical microtubule orientation perpendicular to the cell axis, and promote growth in cell
length. Abscisic acid (ABA) (Sakiyama and Shibaoka,
1990; Ishida and Katsumi, 1992; Sakiyama-Sogo and
Shibaoka, 1993), cytokinins (Su and Howell, 1992, 1995;
Shibaoka, 1994), and ethylene (Steen and Chadwick,
1981; Lang et al., 1982; Roberts et al., 1985) promote a
cortical microtubule orientation in the direction of the
cell axis and consequently diminish growth in cell length
and may induce radial expansion. Mayumi and Shibaoka
(1996) have shown that in cells where microtubules cycle
between longitudinal and transverse orientation, auxin
promotes reorientation to the transverse position.
6-dimethylaminopurine (DMAP), a protein kinase inhibitor, favours microtubule reorientation from transverse
to longitudinal (Mizuno, 1994).
All growth regulators have influence on root hair
growth and appearance (Ridge, 1996), but studies on the
involvement of the cytoskeletal system and calcium in
this process have only just begun. Felle and Hepler (1997)
showed a transient 70% growth reduction in Sinapis root
hairs treated with 10~7 M indole acetic acid (IAA). This
reaction occurred concomitantly with a rapid depolarization of the plasma membrane and a drop in [Ca2 + ] c along
the entire length of the root hair but more in the tip than
the base. Kinetin and ABA cause root hair branching
and other aberrations such as swelling of the tip or base,
twisting, curling and bending (Ridge, 1996 and references
therein). For example, Arabidopsis roots treated with
ABA produce short bulbous root hairs (Schnall and
Quatrano, 1992).
Ethylene stimulates root hair production in pea, faba
bean and lupin, induces root hair formation in areas
normally lacking root hairs in radish and mustard, and
induces root hair formation in tulip even though this
species normally lack root hairs (Abeles et al., 1992). In
an environment of excess ethylene, root hairs have been
found to arise from the atrichoblasts, the root epidermal
cells that in control conditions do not form root hairs
(Dolan et al., 1994). Roots grown in the presence of the
ethylene precursor 1-aminocyclopropane-l-carboxylic
acid (ACC) produce root hairs on the atrichoblasts
(Tanimoto et al., 1995), whereas aminoethoxyvinylglycine
(AVG) a blocker of ethylene biosynthesis (Masucci and
Schiefelbein, 1994; Tanimoto et al., 1995; Heidstra et al.,
1997), or Ag + , a blocker of ethylene perception (Heidstra
et al., 1997), inhibit the formation and growth of root
hairs. When Vicia sativa roots growing between a
coverslip and a slide (Fahraeus, 1957) are treated with
AVG or Ag + , the growing root hairs obtain a cytoarchitecture typical for hairs that are terminating growth
(Heidstra et al., 1997), i.e. the vesicle-rich region almost
1890
Miller et al.
disappears, but the reverse fountain streaming remains.
Moreover, the presence of ACC renders the blockers
ineffective (Heidstra et al, 1997). In the same study,
ACC oxidase mRNA is shown to accumulate in the cell
layers opposite the phloem poles. It is likely that the
location of ACC oxidase mRNA accumulation is the site
of ethylene production. In roots such as Arabidopsis the
trichoblasts are located above the anticlinal walls of two
underlying cortical cells, while the atrichoblasts are located over a single periclinal wall of a cortex cell. The
trichoblasts appear more accessible than the atrichoblasts
to putative signals transported through the apoplast,
possibly explaining why additional ethylene can induce
atrichoblasts to form hairs.
Roof hair morphogenesis mutants
Several genes have been discovered that affect root hair
production, elongation and growth characteristics in
Arabidopsis (Wilson et al, 1990; Schiefelbein and
Somerville, 1990; Su and Howell, 1992; Kieber et al.,
1993; Chang et al., 1993; Schiefelbein et al, 1993; Reed
et al, 1993; Aeschbacher et al, 1994; Galway et al, 1994,
1997; Dolan et al., 1994; Masucci and Schiefelbein, 1994,
1996; Masucci et al, 1996; Di Cristina et al, 1996;
Chang, 1996; Leyser et al, 1996; Cernac et al, 1997;
Kieber, 1997; Wang et al, 1997). The genes known so
far to be associated with ethylene and/or auxin effects on
root hair development are Constitutive Triple Response]
(CTR1), Auxin Resistant! (AXR1), Auxin Resistant2
(AXR2), Auxin Resistant3 (AXR3), Ethylene Response!
(ETRJ), and Root Hair Defective6 (RHD6). Other genes
such as RHD2, RHD3, RHD4, and TIP1 are involved in
altered tip growth (Aeschbacher et al, 1994), and
Transparent Testa Glabra (TTG) and Glabra2 (GL2)
(Galway et al, 1994; Masucci and Schiefelbein, 1996;
Masucci et al, 1996; Di Cristina et al, 1996) are involved
in early regulation of epidermis cell fate. Mutation studies
are providing insights into the function of the cytoskeletal
system, calcium, and plant growth hormones in root hair
development and growth.
Kieber et al. (1993) found a Raf-like protein kinase
encoded by the CTR1 gene which is postulated to be a
negative regulator of ethylene signal transduction, while
Dolan et al (1994) found that the recessive Ctrl mutation
causes root hair formation on non-hair forming epidermal
cells. Thus, the CTRl gene encodes a negative regulator
of hair formation while ethylene acts as a positive regulator of hair formation. ETR1 has sequence similarity to
the two-component regulators in bacteria (Chang et al,
1993). Thus, in Arabidopsis ethylene signalling involves
two known regulators, one of which resembles the 'twocomponent' regulators found in bacteria (Chang, 1996).
The auxin resistant mutant ax! has a reduced number
of root hairs (Cernac et al, 1997). Even though these
mutants can initiate root hairs many cannot elongate
properly. Whereas a suppressor of auxin resistance SARI
has the opposite effect on root hair development; the root
hairs are thicker and a higher number of ectopically
placed hairs are found (Cernac et al, 1997). Cloning of
the axrl gene indicates it encodes a protein related to the
ubiquitin-activating enzyme (El) known to function in
the ubiquitin conjugation pathway (Leyser et al, 1993).
Roots of the auxin-resistant Arabidopsis axr2 dominant
mutant lack root hairs almost completely and are insensitive to high concentrations of auxin, ethylene and abscisic
acid (Wilson et al, 1990). Application of exogenous
auxin restores root hair formation suggesting that auxin
is essential for root hair development. Another auxinresistant mutant, Arabidopsis axr3, has pleiotropic defects,
many of which belong to auxin-related processes, and
these roots lack root hairs (Leyser et al, 1996). The
recessive rhd6 mutation causes a reduction in root hair
production and the site of root hair emergence differs
from wild type but once initiated the root hairs appear
normal (Masucci and Schiefelbein, 1994). Rescue of the
rhd6 mutant phenotype occurs when auxin or the ethylene
precursor ACC is added to the growth medium. In
addition, wild-type seedlings grown in the presence of the
ethylene biosynthesis inhibitor AVG look like rhd6 mutants. These results suggest that the RHD6 gene is involved
in root hair initiation and processes involving auxin and
ethylene. Su and Howell (1992) found that a cytokininresistant mutant (ckl) produces hairs shorter than usual
while the phytochrome B mutant hy3 has longer hairs
than usual when grown in the presence of light (Reed
et al, 1993). The genes TTG and GL2, which promote
hairless cells, seem to act early in the developmental
pathway by negatively regulating the ethylene and auxin
pathways (Masucci and Schiefelbein, 1996; Masucci et al,
1996). For instance, losing the function of TTG (Galway
et al, 1994; Masucci et al, 1996) and GL2 (Masucci
et al, 1996; Di Cristina et al, 1996) allows the development of root hairs in all files, suggesting that these genes
are positive regulators of non-hair fate. Therefore, it is
believed that the main role for TTG and GL2 is to inhibit
non-hair cells from responding to the hormonal signals.
For a model speculating on how these genes function in
root hair initiation and development see Masucci and
Schiefelbein (1996).
Proper initiation of root hairs appears to require the
RHD1 gene product, while hair elongation requires
RHD2, RHD3 and RHD4 gene products (Schiefelbein
and Somerville, 1990). Schiefelbein et al. (1993) found
the tip! mutant has shorter branched root hairs. Analysis
of the tip! mutant indicates that pollen tube growth as
well as root hair growth is affected, suggesting that the
TIP! gene is involved in a fundamental aspect of tip
growth in plant cells (Schiefelbein et al, 1993). The rhcB
mutant in Arabidopsis produces short and wavy root hairs
Root hair tip growth
1891
that have approximately one-third the volume of a wildtype hair. Galway et al. (1997) determined that vacuole
enlargement and thereby normal cell expansion in these
hairs is hindered. Asymmetric growth is the probable
cause of the wavy-hair phenotype observed in these hairs.
The RHD3 gene has been cloned and encodes a novel
protein with putative GTP-binding motifs (Wang et al.,
1997).
The precise mechanism whereby ethylene and auxin
influence root hair initiation and outgrowth is not understood. Since root hair initiation involves a change in the
direction of cell expansion from mainly longitudinal to
the root axis to one of tip growth, signals that influence
root hair initiation and growth must have an influence
on the cytoskeleton. Discovery and analysis of new mutants will help clarify the role of these gene products in
root hair formation.
Perception and transduction of signals from rhizobia in
legume root hairs
Lipochito-oligosaccharides, also called nodulation factors
(Nod factors), are well-characterized signal molecules
excreted by Rhizobium bacteria (Lerouge et al., 1990);
Spaink; 1992; Fisher and Long, 1992; Denarie and
Cullimore, 1993; Carlson et al., 1995) that induce the
formation of nitrogen-fixing nodules in legumes (Mylona
et al., 1995). The first morphogenetic modification
observed after Nod factor application is root hair
deformation. In the assay used for Vicia sativa, the root
hairs terminating growth are susceptible to Nod factors
(Heidstra et al., 1994), and root hair deformation is the
reinitiation of growth in these hairs (De Ruijter et al.,
unpublished results). This process starts with a swelling
of the root hair tip (Fig. 8), which is an active process
requiring protein synthesis (Vijn et al., 1995). After
application of lipochito-oligosaccharides, [Ca 2+ ] c (Fig. 9)
and the spectrin-like antigen (Fig. 10a, b) are mobilized
to the area of cytoplasm bordering the plasma membrane
of the swelling, showing that root hair deformation in
this assay is a two-step process: first unpolarized isodiametric cell expansion followed by reinitiation of tip
growth.
It is important to decipher which signal transduction
pathways are exploited by the bacteria. Van Workum
et al. (1995) have hypothesized that Nod factors elicit
hormonal changes including the production of ethylene.
This hypothesis is based on the observation that an
exopolysaccharide-deficient mutant of Rhizobium leguminosarum biovar viciae, which has impaired root nodule
formation on its host plant, can nodulate that host if the
level of ethylene production in the host is minimized by
AVG or by growing the root in the dark. Further evidence
for a role for ethylene in Nod factor-induced signalling
comes from experiments in which roots of Vicia sativa
Fig. 8. DIC image of Vicia sativa root hair after application of 10 10 M
Nod factor (A) at 1 h 10 min, swelling; (B) at 1 h 25 min, forming a
vesicle-rich region; (C) at 1 h 45 min, outgrowth. Bar= 15 ^m.
ssp. nigra, grown under aphysiological conditions, i.e. in
the light, were inoculated with Rhizobium leguminosarum
bv. viciae, or with Nod factors derived from these bacteria.
The roots remained short, became twice as thick, and
produced an increased number of hairs. This phenotype
could be mimicked by addition of the ethylene releasing
compound ethephon and inhibited by adding AVG (Van
Spronsen et al., 1995), suggesting that the phenotype is
caused by excessive ethylene production. These results
taken together with the fact that root hair formation in
Arabidopsis as well as legumes involves ethylene, suggests
that ethylene could be part of Nod factor-activated signal
transduction. However, ethylene does not appear to be
an intermediate compound of the Nod factor-induced
signal transduction pathway leading to root hair deformation in the Vicia sativa root hair deformation assay
(Heidstra et al., 1997).
Ardourel et al. (1994) hypothesize that two types of
receptors are involved in the plant response, one that
prepares the plant for symbiotic infection by triggering
root hair tip growth and cell division in the cortical cells
and a second receptor with more stringent structural
requirements needed for cell invasion. Felle et al. (1995)
tested whether Nod factors with modified backbones,
saturated acyl chains or without sulphate could initiate
the depolarization response. The plasma membrane
1892
M/7/eretal.
Fig. 9. Confocal ratio image of the median-optical plane of a View
saliva root hair 1 h 15min after Nod factor application and loaded
with the Ca2+ indicator Indo-1 AM showing high [Ca2*],. in the
peripheral cytoplasm of the swollen tip (red-orange). The vacuole
appears green due to sequestering of Indo-1 AM. Calibrated [Ca2+l-,
represented in pseudocolour, ranges from 150 nM at the base (bluegreen) of the hair to 1.8 mM (red-orange) at the plasma membrane of
the swelling. Bar= 15 fim.
Fig. 10. (A) DIC image of a Vicia saliva root hair 1 h 20min after
Nod factor application, showing a new vesicle-rich region at one site in
the swelling; (B) the same cell labelled with and spectrin antibody,
showing the spectrin epitope in the new hair tip. Bar = 30^m.
responded differently to the various structural motifs of
the signal molecule, suggesting that the different parts
may confer different functions to the signal. Kurkdjian
(1995) found that membrane depolarization in Medicago
sativa depended also on the structural form of the Nod
factor added. Concomitant with the plasma membrane
depolarization, Felle et al. (1996) found rapid alkalinization in alfalfa root hairs after Nod factor application.
The most sensitive response was to the sulphated tetrameric Nod factor that normally causes deformation and
nodulation in alfalfa. Non-sulphated Nod factors could
cause a pH change at high concentrations, but did not
cause plasma membrane depolarization. The different
responses of the sulphated and non-sulphated Nod factors
led the authors to suggest that there are two Nod signal
perception systems, one that is host-specific and one that
recognizes a generic Nod factor structure.
Even though there is no direct evidence for the presence
of Nod factor receptors yet, the depolarization response
suggests that plasma membrane-localized receptors exist.
An indication of the type of signal transduction cascade
that Nod factors might induce comes from the work of
Ehrhardt et al. (1996), who observed spiking, i.e. periodic
[Ca2 *\. oscillations in alfalfa root hairs following hostspecific Nod factor addition. The spiking began after an
average lag period of 9 min with a 1 min interval and
exhibited host specificity (see also Long, 1996). An alfalfa
mutant lacking root hair curling and cell division also
lacked [Ca2 +\. spiking. Plasma membrane depolarization
occurs within 1 min (Ehrhardt et al., 1992), long before
the beginning of the [Ca 2+ ] c spiking, indicating that the
spiking is probably not causal to or mechanistically
related to membrane depolarization.
[Ca 2+ ] c spiking in animal cells is typically associated
with the inositol phosphate signal transduction pathway.
Whether [Ca 2+ ] c spiking in plant cells relates to the same
mechanism is unknown. Some indirect evidence that
plants may utilize this pathway comes from pollen tube
work in which a Ca 2+ -dependent phospholipase C (PLC),
capable of hydrolysing
phosphatidylinositol-(4,5)bisphosphate (PIP 2 ), can lead to the production of
inositol-(l,4,5)-triphosphate (IP 3 ) (Franklin-Tong et al.,
1996). In another experiment, the release of caged IP 3
was correlated with increased [Ca2"1^ and inhibition of
pollen tube growth (Franklin-Tong et al., 1996).
It could be hypothesized that PIP 2 is instrumental in
the reorganization of the actin cytoskeleton via actin
binding proteins, such as profilin, in a manner such that
Golgi vesicles accumulate at the hair tip, allowing insertion of Ca 2 + channels in the plasma membrane there,
leading to more Ca 2 + influx and sustained tip growth.
Alternatively, the elevated [Ca 2+ ] c at the plasma membrane of the root hair tip after Nod factor application
(Fig. 10) may cause a reorganization of the actin cytoskeleton without interference of the inositol signal
Woof hair tip growth
transduction pathway. Whether the formation of this
gradient is caused directly by a local depolarization of
the plasma membrane or by a signal transduction cascade
in which [Ca 2+ ] c oscillations act as second messenger, still
has to be studied.
Conclusions
This review has focused on structural elements of the tip
growth process since knowledge about these structural
elements is needed to understand plant cell morphogenesis. Progress in the identification of signalling pathways
leading to morphogenesis is just beginning, but it is clear
that the cytoskeleton is involved. In pollen tubes of pea,
Lin et al. (1996) found a Rho GTPase called Rop
GTPase. In other organisms molecules such as these
regulate the organization of the actin cytoskeleton. This
Rop GTPase is concentrated in the cortical region of the
pollen tube apex, and on the periphery of the generative
cell. These results suggest that Rop GTPase may be
involved in the signalling mechanism that controls the
actin-dependent tip growth of pollen tubes. Other signalling systems are likely to be involved. Smith et al. (1994)
applied two inhibitors of serine/threonine protein phosphatases and found inhibition of both the initiation and
elongation of root hairs in Arabidopsis.
A cell can only exert its specific functions after it has
developed a specific morphology, i.e. function follows
form (De Loof et al., 1996). The cytoskeleton plays an
important role in establishing this form. Exocytosis, one
of the important phenomena of plant morphogenesis, is
by its very nature a cellular event. The phenomenon, and
the cascades of cellular reactions leading to it, can only
be understood by using a multidisciplinary approach.
Concepts must ultimately be tested in whole living cells.
This type of research will be a very active field in the next
few years and root hairs possess the characteristics necessary to serve as an experimental system for studying
interactions at the plant cell surface, triggered from the
inside or from the outside of the cell leading to cell
morphogenesis. The use of genetic analysis and signal
molecules such as Nod factors will give insights into the
signal transduction pathways involved in root hair growth
and plant morphogenesis.
Acknowledgements
We wish to thank Drs Ton Bisseling, Claire Grierson, Peter
Hepler, Nicholas Battey, and John Schiefelbein, for helpful
discussions, Jan Vos for kindly providing the data on carrot
suspension cells, and Allex Haasdijk for the artwork. This work
was supported by the National Science Foundation through a
Postdoctoral Fellowship in Biosciences Related to the
Environment granted to DDM. Grant number DBI 9509441.
1893
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