Download Membrane trafficking and polar growth in root hairs and pollen tubes

Journal of Experimental Botany, Vol. 58, No. 1, pp. 65–74, 2007
Intracellular Compartmentation: Biogenesis and Function Special Issue
doi:10.1093/jxb/erl059 Advance Access publication 26 July, 2006
SPECIAL ISSUE PAPER
Membrane trafficking and polar growth in root hairs and
pollen tubes
Prisca Campanoni* and Michael R. Blatt
Laboratory of Plant Physiology and Biophysics, IBLS Plant Sciences, Bower Building, University of Glasgow,
Glasgow G12 8QQ, UK
Received 20 February 2006; Accepted 23 May 2006
Abstract
Root hairs and pollen tubes extend by rapid elongation
that occurs exclusively at the tip. Fundamental for
such local, tip-focused growth (so-called ‘tip growth’)
is the polarization of the cytoplasm that directs secretory events to the tip, and the presence of internal
gradients and transmembrane flux of ions, notably
Ca21, H1, K1, and Cl2. Electrophysiological and imaging studies using fluorescent markers have sought to
link ion gradients with growth and membrane trafficking. Current models recognize membrane trafficking as
fundamental to tip growth, notably its role in supplying
lipid and protein to the new plasma membrane and cell
wall that extend the apex of the cell, and a complementary role for endocytosis in retrieving excess membrane and in recycling various protein fractions. The
current state of knowledge is reviewed here in order
to highlight the major gaps in the present understanding of trafficking as it contributes to polar growth in
these cells and recent results, that suggest a role for
membrane trafficking in the active regulation of ion
channel turnover and activity during polar tip growth,
are discussed.
Key words: Cytosolic free calcium, cytosolic pH, endocytosis,
exocytosis, ion channel, K1 channel, root hair, SNAREs, tip
growth, vesicle transport.
Introduction
Root hairs develop at the base of competent root epidermal
cells (trichoblasts), and they serve to increase the total root
surface area for water and nutrient acquisition, to anchor
the plant to the soil, and to support symbiotic interactions
with micro-organisms living in the soil (Gilroy and Jones,
2000). Pollen grains develop pollen tubes on the floral
stigma and grow through the female tissues of the pistil to
deliver their genetic material to the egg (Taylor and Hepler,
1997; Lord, 2000). To accomplish these tasks, both root
hairs and pollen tubes display a highly specialized growth
form in which the cell elongates unidirectionally and only
at the very tip. This extreme form of polarized growth, or
tip-growth, sustains prodigious rates of elongation: Root
hairs grow at rates of 10–40 nm sÿ1 (Galway et al., 1997;
Wymer et al., 1997), comparable to that of animal neuron
growth cones (Gomez and Spitzer, 1999), and lily pollen
tubes (Lilium longiflorum L.) can achieve the astonishing
growth rate of 250 nm sÿ1 (Pierson et al., 1996; Messerli
et al., 2000). It is no surprise, therefore, that to support these
growth rates root hairs and pollen tubes must possess an
appropriately amplified turnover of cytoskeleton, cytoplasmic structures and organelles, and an accelerated traffic of
membrane vesicles that deliver membrane and cell wall
material to the growing tip.
In root hairs and in pollen tubes growth is restricted to
the very apex of the tip: in root hairs of Medicago
truncatula, for example, maximal extension occurs in an
annulus just behind the apex (Shaw et al., 2000) and, as
a result, the tip is not perfectly hemispherical, but flattened
at the very apex and tapered along the edges toward the
cylindrical portion of the cell. Presumably these variations
in extension rate across the apex dome reflect underlying
variations in the cellular machinery for growth, notably in
the delivery of secretory vesicles and release of cell wall
constituents. Internally the cytoplasm exhibits a high degree of polarized zonation (Fig. 1): root hairs typically
show a clear region of vesicle accumulation, the hyaline
‘cap’ in the extreme apical 2–3 lm of the tip and, behind
this zone, a region of extremely densely packed cytoplasm
containing endoplasmic reticulum (ER), Golgi apparatus
* To whom correspondence should be addressed. E-mail: [email protected]
ª The Author [2006]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
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66 Campanoni and Blatt
Fig. 2. Polarization of endomembrane secretory system at the side of
root hair tip growth. Arabidopsis seedlings transformed with a YFPHDEL construct as a marker for the ER/Golgi interface. Pictures were
taken using a confocal laser scanning microscope and stacks were
overlapped for three-dimensional reconstruction of the image. (A)
Bulging root hair at the stage of establishment of polar tip growth. (B)
Growing root hair. Arrows indicate the region of ER accumulation at
the growing apex. Scale bars, 10 lm.
Fig. 1. Cytoplasmic organization of a growing root hair. Schematic
diagram of the internal zonation of a growing root hair. The tip is
characterized by a high cytoplasmic calcium concentration (indicated
by shadowing), and is filled by densely packed secretory vesicles, which
fuse to the membrane. Organelles including mitochondria, ER, and Golgi
(endomembrane system) are excluded from the apex. The nucleus (N)
follows the growth at a fixed distance behind the tip. The vacuole (VAC)
occupies the bulk of the root hair. Cortical microtubules and cytoplasmic
actin cables run parallel to the direction of growth. Shorter microfilaments
guide the delivery of the vesicles to the clear zone.
and mitochondria (Fig. 1; endomembrane system). The
nucleus follows the tip at a fixed distance (Galway et al.,
1997; Geitmann and Emons, 2000). Pollen tubes similarly
display a clear zone at the extreme apex that contains vesicles, while larger organelles and other structures are excluded from this zone and locate several microns behind
the apex (Geitmann and Emons, 2000; Lennon and Lord,
2000). Further back from the tip, the bulk of both root
hairs and pollen tubes is dominated by a large vacuole.
The cytoplasm is restricted to a thin peripheral layer, with
the exception of accumulation in the extreme apex.
Membrane trafficking in tip growing cells
Tip growth correlates strictly with the polarization of
cytoplasm and secretion in root hairs. Once the pattern of
trichoblasts is specified along the root epidermis, each cell
establishes a characteristic apical–basal polarity, with the
nucleus migrating to the centre of the trichoblast and then
to its base, where the dome of the future hair first forms.
It is at this stage that the polar tip growth is established:
the cytoplasm beneath the dome polarizes to initiate a hyaline zone, and vesicle fusion at the plasma membrane
must accelerate locally. Polarization of other organelles accompanies these changes, including a recruitment of the
endoplasmic reticulum and Golgi behind the new growing
tip. Figure 2A shows the accumulation of the ER at the
base of the cell and beneath the dome, visualized here by
expressing an ER-retained YFP–HDEL construct after transient transformation of root epidermal cells (P Campanoni,
C Davis, J-U Sutter, MR Blatt, unpublished data). This
organization of the cytoplasm is retained and must sustain
tip growth during the development of the root hair (Fig.
2B). On maturation, the root hair stops growing and the
endomembrane system redistributes along the hair,
now largely vacuolarized (Dolan, 2001; Carol and Dolan,
2002). Remarkably, although membrane traffic is clearly
central to these processes, very little is known of its
mechanism, much less of its co-ordination. The following
discussion serves to outline our current understanding of
the membrane trafficking events involved in plant polar
tip growth.
Exocytosis
In general, the endomembrane system involved in vesicle
transport in plants is very similar to that described for
other eukaryotic cells and, indeed, many regulatory and structural proteins involved in membrane traffic are wellconserved between plants and animals (reviewed in Battey
et al., 1999; Sanderfoot and Raikhel, 1999; Vitali and
Denecke, 1999; Pratelli et al., 2004; Sutter et al., 2006a).
Knowing the function and localization of these proteins in
tip growing cells will certainly add to an understanding of
the relationship between vesicle transport and tip growth.
To date, several tip growth-impaired phenotypes have
been correlated with mutations in different ras-like GTPases,
such as the Rop-GTPases that contribute to cellular polarity
by regulating the actin cytoskeleton, ARFs (ADP-ribosylation factors) that are essential for vesicle formation at the
donor membrane, and Rab-GTPases that are responsible
for vesicle docking to the target membrane (for review on
small GTPase proteins in plants, see Molendijk et al., 2004).
Membrane trafficking in tip growth 67
Among these, Rop2 and RabA4b have been shown to
localize at the tip during root hair elongation and appear
to function as positive regulators of root hair growth
(Molendijk et al., 2001; Jones et al., 2002; Preuss et al.,
2004, 2006). Expression of constitutively active AtRop4
and AtRop6 GTPases abolished polarized growth
(Molendijk et al., 2001).
By contrast, only two mutations have been reported so
far that affect other elements of the machinery involved in
mediating vesicle fusions to the plasma membrane and
yield tip growth-defective phenotypes. In Zea mays the
mutation sec3/rth1 (roothairless1) correlated with defects
in root hair elongation (Wen et al., 2005), and in Arabidopsis thaliana sec8 mutants display defects in pollen
tube initiation and growth (Cole et al., 2005). Rth1/Sec3
and AtSec8 are orthologues of two subunits of the exocyst
complex that, in yeast and animal cells, guides polarized
exocytosis. Intriguingly, in these cells Sec3 and Sec8 are
part of the same multi-protein complex, composed of eight
different subunits (Exo70, Exo84, Sec3, Sec5, Sec6, Sec8,
Sec10, and Sec15; for review see Li and Chin, 2003).
Although it is known that Arabidopsis and rice genomes
contain genes orthologous to all the exocyst subunits (Elias
et al., 2003), only recently have any of these subunits been
shown to act as a complex (Zarsky et al., 2004). The fact
that neither the sec3 nor the sec8 mutants were impaired
concurrently both in root hair and in pollen tube growth,
suggests that the two proteins may function within different pathways in each case. Alternatively, the contrasting
phenotypes might be due to a redundancy in the genome,
with specific isoforms differentially expressed into the two
cell types.
Astonishingly, no clear connection has yet been made
between tip growth and SNARE proteins. SNARE complexes mediate the final stages of vesicle fusion throughout
the endomembrane system and at the plasma membrane
(reviewed by Sanderfoot and Raikhel, 1999; Pratelli et al.,
2004; Sutter et al., 2006a). Therefore it would be reasonable to expect that alterations in SNARE function might
lead to tip growth defects. At present, however, the only
Arabidopsis SNARE mutants reported to display defects in
pollen development are members of the Syp2 and Syp4
SNARE subfamilies of syntaxin-like proteins. These two
subfamilies are thought to contribute to vesicle traffic
between the vacuole and trans-Golgi network (Sanderfoot
et al., 2001). No examples of mutations in SNARE proteins
have yet been reported which impair root hair tip growth.
Endocytosis
Along with exocytosis, localized endocytosis is welldocumented as a phenomenon associated with budding in
yeast. For example, the endocytic protein Sla2p/End4p
which links endocytosis with the actin cytoskeleton, is
crucial for establishing zones of polarized growth in yeast
(Castagnetti et al., 2005). In plants, recognition is fairly
recent of endocytosis as a common cellular process, as for
many years it was argued that endocytic events could
not occur against the opposing force of cellular turgor
pressure. Experiments demonstrating an internalization of
impermeant fluorescent dyes provided a first direct confirmation of endocytosis in plants (O’Driscoll et al., 1993;
Carroll et al., 1998). Subsequent studies showed that
KAT1, an inward-rectifier K+ channel, was constitutively
retrieved together with styryl dyes as putative membrane
markers, and that these events proceeded from the plasma
membrane against high turgor pressure, both in Vicia faba
protoplasts and in the intact guard cells (Hurst et al., 2004;
Meckel et al., 2004). For recent reviews on endocytosis,
see Geldner (2004), Šamaj et al. (2005), Murphy et al.
(2005), and also Sutter et al. (2006a, b).
Plant secretory vesicles are commonly 100–150 nm in
diameter; they bud from the Golgi at a rate of 2–4 minÿ1 per
Golgi stack and, in hypersecretory cells such as growing
pollen tubes, they arrive at the plasma membrane at rates of
several thousands per minute within the region of the tip
(Steer and Steer, 1989; Picton and Steer, 1983). Calculations of secretion rate in the growing tip of pollen tubes
(Picton and Steer, 1983; Steer, 1988; Derksen et al., 1995)
and root hairs (Morre and Van der Woude, 1974; Emons
and Traas, 1986) have indicated that the total membrane
surface area added to the plasma membrane exceeds its
expansion by as much as 5-fold. Clearly, on the basis of
these calculations, the excess membrane must be retrieved
by endocytosis, but many details are still lacking. Whereas
the vast bulk of exocytotic events occurs in the apical dome,
because lateral displacements of membrane material are
likely over the surface of the tip and behind, it is less
obvious where endocytosis might occur or even whether
such events are localized. In fast-growing tobacco pollen
tubes, clathrin-coated endocytic vesicles are known to form
in the subapical region 6–15 lm below the tip (Derksen
et al., 1995) but, by contrast, endocytosis was found to take
place at the apex in Arabidopsis pollen tubes even though
their growth rates are generally much slower (Blackbourn
and Jackson, 1996). Tip-localized internalization of the
styryl dye FM4-64, a putative membrane marker, has also
been reported in lily pollen (Parton et al., 2001).
In root hairs, these differences are probably related to
maturation state. For example, Emons and Traas (1986),
Galway et al. (1997), and Voigt et al. (2005) reported that
the distribution of coated vesicles along the tip varied with
both age and growth rate. Coated vesicles were observed
to emerge from the subapical plasma membrane in the
clear zone during active growth, but the frequency of these
events dropped off once root hairs reached maturity. At
least some of these endocytotic vesicles were recycled for
secretion in the tip or fused to larger bodies and proceeded
to the base of the hair to fuse with the vacuole through
a microfilament-driven ER-independent pathway (Ovečka
et al., 2005; Voigt et al., 2005).
68 Campanoni and Blatt
An important, but often overlooked point is that the
exo- and endocytosis are almost certainly coupled. Obviously, it is essential for tip growth that a balance between
the rates of secretion and endocytosis is preserved to ensure
an appropriate flux of material for growth. It is also
important that membrane components are recycled in
a temporally and spatially consistent manner to support
growth. Thus, it is no surprise that perturbing one process
can result in changes in the other, or that inhibitors that
affect membrane trafficking, such as brefeldin A (BFA),
also rapidly suppressed cell growth (Schindler et al., 1994;
Baskin and Bivens, 1995; Cho and Hong, 1995; Morris
and Robinson, 1998). Remarkably, in pollen tubes treated
with BFA, inhibition of exocytosis has been reported to
accompany an increase in the rate of endocytosis (Wang
et al., 2005; but also see Parton et al., 2001), suggesting
that BFA-dependent growth inhibition is a direct consequence of both exocytosis inhibition and endocytosis
stimulation.
Ion gradients
Both exocytosis and endocytosis are often sensitive to
free cytosolic calcium levels (Taylor and Hepler, 1997;
Bibikova et al., 1997; Sutter et al., 2000; Camacho and
Malhó, 2003), and such observations correlate well with
cytoplasmic tip-focused Ca2+ gradients that have been
measured in tip-growing cells. The phenomenology of
Ca2+ gradients is also closely coupled to internal gradients
and transmembrane flux of several ions. In addition to Ca2+,
a substantial body of electrophysiological and imaging data
have associated K+, H+, and Clÿ with tip growth.
Calcium
Several studies have shown an intracellular calcium
gradient focused on the apex of both growing pollen tubes
(Taylor and Hepler, 1997; Gilroy and Jones, 2000) and root
hairs (Bibikova et al., 1997; Felle and Hepler, 1997;
Wymer et al., 1997). This gradient is tightly coupled to
polar tip growth and it corresponds closely with that of
vesicle fusion and retrieval, as shown in Fig. 1 (Bibikova
et al., 1997; Felle and Hepler, 1997; Taylor and Hepler,
1997; Wymer et al., 1997; Gilroy and Jones, 2000). In both
root hairs and pollen tubes, the concentration of calcium
decreases sharply, from very high levels near the inner
surface of the plasma membrane at the tip, to basal levels
some 20 lm behind the tip. Calcium gradients measured in
the medium surrounding root hairs (Gilroy and Jones,
2000) and pollen tubes (Holdaway-Clarke et al., 1997;
Messerli et al., 1999) also indicate a strong flux of calcium
directed inward at the apex. Tip growth normally cannot be
uncoupled either from these calcium gradients or from
transmembrane flux of the divalent, although such a statement belies a complexity of detail behind their relationship,
especially in relation to oscillations in growth and in free
Ca2+ concentration (below).
Protons
Studies of lily pollen tubes with fluorescent pH-indicators
first uncovered the presence of a constitutive alkaline band
at the base of the clear zone and an acidic domain at the
extreme apex, where active growth takes place (Feijó et al.,
1999). In agreement with the intracellular proton gradient,
measurements of the extracellular currents showed that
growing pollen tubes maintained a circuit of protons with
a net influx at the extreme apex and an efflux in the region corresponding to the alkaline zone (alkaline cytosolic
pH). It has been postulated that this circuit of protons
could be driven by plasma membrane H+-ATPases that are
polarly localized along the tube (Feijó et al., 2004), but the
balance of H+ flux implies that current return, presumably
via H+-coupled transporters, is likely to be just as, if not
more important for establishing any pH gradients.
Although the gradient of pH is generally recognized as
fundamental for pollen tube tip growth, the presence of
proton gradients in growing root hairs is still an issue of
contention. A localized rise in cytosolic proton concentration in root hairs is detectable just at the very early
stages of root hair development, at a time when the local
cell wall acidification and loosening led to a bulging of the
new hair from the trichoblast (Bibikova et al., 1997;
Takahashi et al., 2003). This proton gradient is closely
associated with the production of reactive oxygen species
(Foreman et al., 2003) and related to the state of nutrition
(Shin et al., 2005). Once the polarity is established, however, a proton gradient along the growing hair was no
longer observed (Herrmann and Felle, 1995; Bibikova
et al., 1997; Parton et al., 1997).
What difficulties lie behind measurements of H+? Because
+
H is highly mobile by comparison with other ions in
solution, local proton gradients are easily perturbed during
measurement and lost. Other problems relate to the buffering
capacity of pH indicators. Thus, an excess in the concentration
of pH indicator dyes used during the experiments can have
a disproportionate effect in shuttling protons, buffering and
dissipation of local gradients. In short, questions of pH
gradients in root hairs and their physiological correlates still
need addressing. In fact, strong currents of protons have been
measured in the extracellular vicinity of root hairs, with an
influx towards the apex and a marked efflux toward the base
of the hair (Gilroy and Jones, 2000), suggesting the presence
of an internal proton gradient. It is worth noting, too, that
an alkalization of the cytosol was observed in conjunction
with changes in growth direction of root hairs in response to
Nod factors (Cárdenas et al., 2000).
Potassium
If proton gradients have a minor role in driving tip growth
in root hairs compared to pollen tubes, potassium ions are
Membrane trafficking in tip growth 69
of fundamental importance to root hair tip growth as the
predominant inorganic solute (Hepler, 2001). Recent
studies of mutants with aberrant root hair phenotypes have
served to underline this point, as well as raising many more
questions, especially in relation to the potassium transporter
TRH1. This transporter belongs to the AtKT/AtKUP/HAK
gene family and was isolated from a mutant screen for
altered root hair morphology and elongation. The Arabidopsis trh1 (tiny root hair 1) mutant plants failed to
establish the cytoplasmic zonation characteristic of tip
growth in root hairs (Rigas et al., 2001; Desbrosses et al.,
2003). A link between TRH1 and auxin transport is also
a feature of these mutants: subsequent studies of VicenteAguillo et al. (2004) indicated that trh1 plants show ectopic
expression of an auxin-induced reporter gene in the root
stele as a result of an impaired transport of auxin (VicenteAgullo et al., 2004). The effect of trh1 loss-of-function on
root hair elongation, then, might be explaned as a secondary effect due to suboptimal concentrations of auxin in
the root epidermis. In short, it remains to be established
whether the primary action of the TRH1 gene product is in
auxin rather than in potassium transport.
The major channels responsible for K+ transport across
the plasma membrane of root hairs themselves are the
outward-rectifier K+ channel GORK and the inward-rectifier
K+ channels AKT1 and AKT2/AKT3 as well as the ‘silent’
K+ channel AtKC1, identified from expression profile and
electrical property studies (Ivashikina et al., 2001; Reintanz
et al., 2002; Pilot et al., 2003). Among the inward-rectifier
K+ channels, AKT1 underlies the predominant inward K+
current in root hairs and is probably modulated by AtKC1.
Significantly, disrupting the AKT1 gene resulted in complete
loss of the inward K+ current in these cells.
In root hairs AKT1 also seems to have a role in polar tip
growth. Desbrosses et al. (2003) found that Arabidopsis
plants homozygous for the akt1 loss-of-function mutation
displayed altered root hair tip growth. Curiously, mutant
root hairs were longer than in the wild type when the plants
were grown in the absence of potassium, but shorter if
grown in potassium concentrations higher than 10 mM.
Thus, at low external concentrations of potassium AKT1
seemed negatively to regulate hair tip growth, exerting
a function opposite to that of TRH1. Moreover, the root
hair phenotype of the double akt1-trh1 mutant did not
exhibit a clear epistasis, reinforcing the idea that the two
channels affect root hair growth via very different mechanisms (Desbrosses et al., 2003).
If much is already known about potassium transport in
root hairs, little is known about it in pollen tubes.
Nevetheless, potassium ions also seem to play a role in
tip growth in pollen tubes. Mutations in the pollen-specific
Arabidopsis inward-rectifying K+ channel SPIK, also
belonging to the Kv channel family, impared the growth
of the mutant pollen tubes with a consequent decrease in the
competitive ability for fertilization (Mouline et al., 2002).
Chloride
Chloride ions have also been associated with tip growth.
Intriguingly and in contrast to potassium, chloride is lost
from the apex and it is recovered by an influx behind the tip
(Zonia et al., 2001). Inhibition of the chloride flux via
treatments with inhibitors of Clÿ channel activity led to
a complete block of tip growth and to an increase in pollen
tube volume (Zonia et al., 2002). These observations are
consistent with a role for chloride ions in supporting growth
and the associated increase in cell volume of pollen
tubes. Interestingly, the Clÿ flux appeared to be regulated
by inositol 3,4,5,6-tetrakisphosphate (Zonia et al., 2002).
Phosphoinositides and phospholipids are also known to
contribute to the regulation of vesicle trafficking and polar
tip growth (Malhó, 1998; Zonia and Munnik, 2004;
Monteiro et al., 2005; Vincent et al., 2005; for review see
Martin, 1998).
Coupling tip growth with ion gradients and
membrane trafficking
Understanding the temporal relationship of membrane
traffic in relation to tip growth presents a number of
challenges. Of these, one of the most intriguing puzzles
relates to oscillations in Ca2+, associated ion flux and their
coupling to growth and, presumably, to secretions that are
characteristic of these cells, especially pollen (Pierson
et al., 1996; Wymer et al., 1997). Much attention has
focused on an examination of phase relationships associated with these oscillations on the assumptions that the
calcium gradient at the apex of tip-growing cells participates in secretory processes (Battey et al., 1999) and that
it should be possible to extract hierarchical data giving
information regarding the identity of the primary regulator
of tip growth (Feijó et al., 2001). Indeed, the Ca2+ gradient
oscillates in magnitude in close relation to oscillations in
growth rate (Holdaway-Clarke et al., 1997; Messerli and
Robinson, 1997; Feijó et al., 2001) and styryl dye internalization (Parton et al., 2001). Nevertheless a direct
correspondence between Ca2+ oscillations, vesicle delivery
and turnover, and growth in pollen has proven more
difficult to assess, since the peak in calcium oscillations
lags the peak of growth rate by 4 s (Messerli et al., 2000).
Still more confusing, the influx of calcium from extracellular medium also oscillates, but with a lag of 11–15 s
on growth rate, and with a significant delay in relation to
the fluctuations in cytoplasmic-free Ca2+ concentration
(Holdaway-Clarke et al., 1997; Messerli et al., 1999).
Proton flux, pH gradient, and chloride flux also oscillate in
relation to growth, but none perfectly in phase with the
growth rate. H+ influx shows a lag relative to growth rate of
about 8 s (Feijó et al., 1999); chloride oscillations exhibit
a lag of 3 s (Zonia et al., 2001). The counter-intuitive nature
of these delays led Messerli et al. (2000) to argue that Ca2+
gradient oscillations and secretion are in fact uncoupled.
70 Campanoni and Blatt
The problem is all the more complex in fact, since
intracellular calcium stores and calcium chelation by cell
wall proteins introduce noise in experimental measurements and thereby confound further analysis. Moreover,
exocytosis of new membrane material at the tip and its
retrieval by endocytosis can be expected to affect channel
and other transporter populations and their patterns of
localization. In short, analysing the cause/effect relationships among ion flux, ion gradients, and vesicle transport at
this point will benefit from direct experimental monitoring and manipulation of membrane traffic and secretion.
For example, studies of pollen tubes treated with Yariv
reagent, which precipitates arabinogalactans proteins and
interferes with the normal deposition of the cell wall,
showed that pollen tube extension, at least, may be uncoupled from secretion in some circumstances (Roy et al.,
1999). Intriguingly, these experiments also showed that
the cells retained tip-focused calcium influx, locally high
cytosolic calcium concentrations, and vesicle transport,
despite the inhibition of growth.
It is worth considering two prevalent models for the
integration of traffic with ion transport that centre (i) on
the turnover of transporters in the membrane and (ii) on
direct protein–protein interactions for the control of transport, although these alternatives are not mutually exclusive per se. In many animal cells, exo- and endocytosis of
membrane transporters and receptors are regulated in
response to external stimuli and lead to longer-term
changes in their activities. Traffic of the EGF (Epidermal
Growth Factor) receptor and the glucose transporter
GLUT4 are two of the best characterized examples. The
EGF receptor, a protein kinase, is negatively regulated
by EGF in animal epithelia by evoked endocytosis of the
EGF-receptor complex which is delivered to the lysosomes
for degradation (for review see Katzmann et al., 2002).
Traffic of the GLUT4 glucose transporter is tightly
regulated in adipose and muscle cells in response to
insulin (Zeigerer et al., 2002). In this case, the hormone
triggers the exocytosis of GLUT4, and can generate a
4–5-fold rise in its population at the plasma membrane of
cells in these tissues over a period of 10–20 min. As
blood glucose is stabilized, GLUT4 recovers from the
membrane dynamically by endocytosis and is recycled
to endosomal compartments (Zeigerer et al., 2002;
Karylowski et al., 2004). Thus, membrane traffic in these
instances is strongly biased to transport control through
what might be considered a ‘transport-passive’ mechanism.
By contrast, the second model is characterized by
interactions between vesicle trafficking proteins and ion
channels, and thus might be considered to be ‘transportactive’ in regulating ion flux. Such direct control is an
important part of the mechanisms regulating neurotransmitter release. Notably, Ca2+ channel interactions with QSNAREs, including mammalian syntaxin 1A, affect the
gating, and hence efficacy of Ca2+ channels that facilitate
Ca2+ influx for neurotransmitter release as well as channel
interactions with other second messengers (Stanley and
Mirotznik, 1997; Bezprozvanny et al., 2000; Jarvis et al.,
2002; Arien et al., 2003; Swayne et al., 2005; Yokoyama
et al., 2005). Direct interactions of SNAREs with neuromuscular and epithelial K+ and Clÿ channels also impact
on the activity of these channels (Cormet-Boyaka et al.,
2002; Tsku et al., 2005).
At present there is little concrete evidence for either
mechanism in plants, but there is no doubt that ion transport
and membrane transport are closely linked. Transportpassive mechanisms of regulation have been proposed to
account for the basipetal distribution of the PIN family of
auxin transporters (Geldner et al., 2001, 2003; Friml et al.,
2002; Blakeslee et al., 2004), for auxin-evoked changes
in the activity of the H+-ATPase (Hager et al., 1991), and
for osmotically-driven changes in KAT1 K+ channel
activity in guard cells (Hurst et al., 2004; Meckel et al.,
2004). To date, none of these studies has offered substantial data that could demonstrate the molecular mechanics
of the trafficking events, but they do set precedents for
traffic-related changes in the population and distribution
of transporters at the plasma membrane.
Support for transport-active mechanisms of regulation
remains thinner still, but there is at least one hint of such
control of ion channels mediated by a SNARE. The
syntaxin NtSyp121 (=NtSyr1) was isolated by screening
a tobacco cDNA library for sensitivity to abscisic acid
(ABA) using Xenopus oocytes (Leyman et al., 1999). The
basis of the screen in oocytes was an evoked, endogenous
Clÿ conductance that is normally sensitive to Ca2+-induced
activation; however, a key line of evidence for function in
the plant was that adding a truncated peptide, corresponding to the cytosolic domain of NtSyp121, suppressed ABAdependent regulation of the K+ and Clÿ channel currents in
Nicotiana guard cells. The simplest interpretation of these
results was that the truncated peptide competed with the
native, full-length protein for interacting partners, thereby
suppressing ABA action on the ion channels. Subsequent
studies (Geelen et al., 2002) indicated that the same peptide
also suppressed vesicle traffic to the plasma membrane.
It might be argued in this case that the influence of the
truncated peptide in suppressing ABA-mediated channel
control reflected its action on ion channel populations,
a transport-passive mechanism, rather than a more direct
mechanism. The difficulty with this argument is that ion
channel response to ABA is normally extremely rapid,
typically complete within some tens of seconds for two of
the three dominant channel currents, and its block was
similarly rapid upon direct injection of the truncated
peptide (Leyman et al., 1999). These effects are more
than an order of magnitude faster even than insulin-evoked
exocytosis of GLUT4 (above) and, hence, are difficult
to explain in the context of vesicle traffic alone. In fact,
recent work from this laboratory (Sutter et al., 2006b) has
Membrane trafficking in tip growth 71
demonstrated that KAT1, the Arabidopsis homologue to
one of these ion channels, is anchored within discrete
microdomains in the plasma membrane when expressed in
tobacco leaves. Intriguingly, this microdomain architecture,
and channel anchoring, is severely disturbed when the
KAT1 protein is co-expressed with the same truncated
fragment of the SNARE that affects ABA control of the
channels. These results indicate a role for the SNARE in
stabilizing and localizing the channel at the plasma
membrane and they raise some interesting questions about
roles for the vesicle trafficking protein in regulation of the
channel through its situation in the membrane. Indeed, it
is very likely that spatial position and associations with
other proteins are essential for correct transmission of
evoked signals. It will be important now to identify the
nature of the proteins that associate locally with these ion
channels and the SNAREs.
A complementary model could be proposed for instance
to explain the pattern of H+ flux around the tip of the
growing pollen tube. According to this model, the polar
distribution of the H+ flux maps to the distribution of the
H+-ATPase that is thought to be intercalated within the
plasma membrane by exocytosis at sites distal from the tip
and then to migrate within the plasma membrane and to be
actively retrieved outside the alkaline zone. Alternatives
could entail targeted exocytosis of the H+-ATPase and/or
localized endocytosis and depletion of H+-ATPase without
significant lateral movements. These same ideas extend to
the ion channels and other transporters that contribute to
ion flux and gradients during tip growth, as shown in Fig. 3.
For example, anchoring mechanisms involving cytoskeleton, binding proteins, and phospholipids could be important
for local ‘trapping’ of selected, laterally-mobile transporters
either at the apex or over more distal regions of the plasma
membrane. Alternatively, these proteins could be spatially
targeted with exo- and/or endocytosis ensuring their exclusion from the adjacent regions (Fig. 3B). Some hint for
targeted exocytosis and endocytotic depletion may be
found in recent studies of trafficking in root hairs and pollen
using FM dyes (Ovečka et al., 2005; Wang et al., 2005).
It remains to be seen whether these observations extend to
the pattern of distribution of H+-ATPase and ion channels.
Fig. 3. Schematic model for ion channel secretion at the apical plasma
membrane of a tip growing cell. (A) Diffuse exocytosis (white arrows) of
the channels (black dots) to plasma membrane. Lateral diffusion within
the membrane (shadowed arrows) brings the channels at the apex, where
they anchor enriching the membrane locally. (B) Site-directed exocytosis
(white arrow) of ion channels (black dots) at the apex. Localized
endocytosis (grey arrows) depletes the channels from behind the tip.
hairs and pollen tubes over the past two decades. By
contrast, our understanding of membrane traffic in these
cells and the molecular players involved remains sparse. It
is expected that recent advances in molecular tagging
strategies and approaches to single-cell measurements, as
well as mutant analyses, should aid significantly in resolving the links between ion gradients, ion flux, and
membrane trafficking in polar growth.
Acknowledgement
Conclusions
Although evidence is mounting for a close interplay
between membrane vesicle traffic and the regulation of
ion transport in plants, there are still large gaps in our
current knowledge. Key elements of these processes relate
to targeting and localized activities in many cell types, but
none more so than in cells that exhibit tip growth. Detailed
information on the characteristics of ion transport and
intracellular ion gradients has been drawn from electrophysiological and fluorescence imaging studies of root
This review was prepared with support from the Biotechnology and
Biological Sciences Research Council.
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