Download Organelle motility in the pollen tube: a tale of 20 years

Journal of Experimental Botany, Vol. 60, No. 2, pp. 495–508, 2009
doi:10.1093/jxb/ern321 Advance Access publication 26 December, 2008
REVIEW PAPER
Organelle motility in the pollen tube: a tale of 20 years
Giampiero Cai* and Mauro Cresti
Dipartimento Scienze Ambientali, Università di Siena, via Mattioli 4, I-53100 Siena, Italy
Received 10 September 2008; Revised 18 November 2008; Accepted 18 November 2008
Abstract
Organelle movement is an evident feature of pollen tubes and is essential for the process of tube growth because it
enables the proper distribution of organelles and the accumulation of secretory vesicles in the tube apex. Organelles
move along the actin filaments through dynamic interactions with myosin but other proteins are probably responsible
for control of this activity. The role of microtubules and microtubule-based motors is less clear and somewhat
enigmatic. Nevertheless, the pollen tube is an excellent cell model in which to study and analyse the molecular
mechanisms that drive and control organelle motility in relation to plant cell expansion. Current knowledge and the
main scientific discoveries in this field of research over the last 20 years are summarized here. Future prospects in the
study of the molecular mechanisms that mediate organelle transport and vesicle accumulation during pollen tube
elongation are also discussed.
Key words: Actin filaments, cell growth, cytoplasmic movement, kinesin, microtubules, myosin, organelles, pollen tube.
Introduction: the importance of actin filaments
Sexual reproduction in seed plants (gymnosperms and
angiosperms) shows a critical evolutionary step consisting
of the use of a specialized male gametophyte (pollen) and its
developmental product (the pollen tube). The advent of the
pollen grain enabled male gametes of seed plants to be
transported over long distances and to resist adverse
environmental conditions. Development of pollen and
pollen tubes, coupled with the evolution of specialized
organs (flowers) in angiosperms, allowed plants progressively to do without water for fertilization and to invade dry
land (Cresti et al., 1992). The pollen tube is a tubular cell
that allows the male gametes to navigate from male to
female flower organs and to do without specialized motile
devices, like flagella, that are required for sperm movement
in lower plants (Tiezzi et al., 1991).
The growth of pollen tubes is a specialized type of cell
expansion (apical growth) characterized by the fusion of
Golgi-derived secretory vesicles in a restricted cellular
domain (the tube tip). Three main domains of pollen tubes
have been proposed: the apical region is the growth domain,
the subapical region is a transition domain, while the base
domain is more or less similar to other plant cells and
contains the typical repertoire of plant cell organelles. As in
other plant cells, organelles and vesicles move rapidly back
and forth along the main axis of the pollen tube: this
activity (‘cytoplasmic streaming’) is required to maintain the
distribution of membraneous structures and to focus Golgi
vesicles in the growth domain of the pollen tube. Movement
of pollen tube organelles is persistent but single organelles
behave independently of each other and move at different
speeds (Fig. 1A). Trafficking of membrane structures occurs
toward the tip, mainly along the tube flanks; in the subapex,
more or less in alignment with a specific actin array (the
actin fringe), organelles reverse their direction of movement
and turn back along the central actin cables. Golgi-derived
secretory vesicles accumulate in the tip domain by moving
along peripheral actin cables; vesicles are probably finally
delivered to the tip region by the actin fringe while recycling
and endocytotic vesicles are then driven back along the
central actin filaments (Fig. 1B). Cytoplasmic streaming is
a key feature of plant cells, whose large size obliged them to
set up an intense circulation of organelles in order to
facilitate intracellular transport and the distribution of
resources.
The pollen tube is, for many reasons, an excellent cell
model in which to study the molecular events underlying
* To whom correspondence should be addressed: E-mail: [email protected]
ª The Author [2008]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.
For Permissions, please e-mail: [email protected]
496 | Cai and Cresti
Fig. 1. (A) Movement of organelles in tobacco pollen tubes as shown by DIC video-microscopy. A single pollen tube is shown after 3 h
of growth. Specific organelles are indicated as representative of different patterns of motility. The organelle indicated by an arrowhead
moves very fast (1.5 lm s 1), while the two organelles indicated by arrows move very slowly; in particular, the organelle marked by
a black arrow is apparently immobile. The organelle labelled with a white line moves slowly (0.5 lm s 1). Numbers at the bottom left are
time in seconds. Bar: 5 lm. (B) Schematic drawing of organelle and vesicle trafficking in a pollen tube. Both structures move toward the
apex along the flanks of the pollen tube; in the subapical region, organelles reverse direction and turn back in the centre of the pollen
tubes. Vesicles are directed toward the apex where they fuse with the plasma membrane; vesicles that do not penetrate the apex are
recycled together with endocytotic vesicles.
organelle transport. It is a single haploid cell, the ultrastructure of which is well known in different species, and can be
manipulated easily. Although most growth events occur in
a spatially-limited region (the tip), the cell shows many
features typical of plant cells, such as interactions with
other cells (the female stigma cells), polarized growth, and
time-dependent cell wall construction. Pollen tubes can be
analysed by different techniques, such as immunocytology,
biochemistry, genetic transformation, and live cell imaging.
This is strictly possible in all species, but a proper combination of different pollen species (Nicotiana tabacum, Lilium
longiflorum, Arabidopsis thaliana, etc) provides a valuable
plant cell model. The aim of this review is to summarize
scientific discoveries in the field of organelle movement and
pollen tube growth in the last 20 years. We start by
reviewing preliminary observations on the critical role of
actin filaments and end by describing the final fate of
secretory vesicles and the interplay between vesicles and
actin filaments in endo/exocytosis. The choice of this span
of time was intentional. In 1988, the 10th Symposium on
Sexual Reproduction in Higher Plants (co-organized by M
Cresti), was held in Siena (Italy). The opening lecture by
Professor Jack Heslop-Harrison (1920–1998) was focused
on organelle transport in pollen tubes (‘The pollen tube:
motility and the cytoskeleton’) and on the use of videoenhanced microscopy to correlate the movement of pollen
tube organelles with the organization of actin filaments and
microtubules (Heslop-Harrison and Heslop-Harrison,
1988). In their meticulous pioneering work, the HeslopHarrisons showed the motility of organelles in older segments of pollen tubes and a putative correlation between
organelle transport and actin cytoskeleton. In 1988, two
other key manuscripts on organelle movement in pollen
tubes were published. Heslop-Harrison et al. (1988) showed
Organelle movement in pollen tubes | 497
that treatment with colchicine did not affect pollen tube
growth and organelle transport but may influence shaping
of the generative cell during its course through the tube.
Work by Kohno and Shimmen (1988) revealed that isolated
pollen organelles moved along Characeae actin bundles
in an ATP-dependent manner and that motility was
influenced by Ca2+. The manuscripts indicated that actin
filaments were the main tracks along which pollen tube
organelles moved and indirectly suggested that myosinbased organelle movement was regulated by variations in
Ca2+ concentrations.
Organization of the cytoskeleton in angiosperm pollen
tubes was already partly known. Condeelis (1974) identified
F-actin in pollen tubes of Amaryllis belladonna, later related
to the movement of pollen nuclei (Cresti et al., 1976). Many
years later, the availability of specific probes enabled
researchers to show that actin filaments and microtubules
are arranged along the longitudinal axis of pollen tubes,
indicating the extent of overlapping as well as several
differences in the apical (growth) region (Pierson et al.,
1986). It was immediately clear that cytoskeletal organization in the tip domain of pollen tubes was hard to decipher
because of fixation artefacts due to the dynamics of the tip
cytoskeleton. Using different chemical procedures, Pierson
(1988) showed differential organization of actin filaments in
the tip domain of pollen tubes and presumed ‘actin foci’ in
the pollen tube flanks; since similar structures are involved
in cell–cell interaction in animal cells, the pollen ‘actin foci’
were postulated to have a role in the process of communication between pollen tube and style. Soon after, the actinbased motor, myosin was discovered. Using antibodies to
animal myosins, Tang et al. (1989a) identified a myosin
heavy-chain polypeptide of 175 kDa in Nicotiana pollen
tubes. Based on its localization, myosin was deduced to be
involved in the movement of vesicles, organelles, and the
male germ unit in pollen tubes. Similar results were
obtained with different types of myosin antibodies (HeslopHarrison and Heslop-Harrison, 1989b), sustaining the
model that actin and myosin were the main protein
mechanism guiding organelle transport.
In order to understand the function of myosins, more
information was also needed on the distribution of actin
filaments. Anti-actin antibodies revealed axially oriented
strands of actin filaments distributed alongside the pollen
tube except at the very tip. Conventional staining with
rhodamine–phalloidin provided similar results (Tang et al.,
1989b). Actin filaments apparently organized as bundles
along the pollen tube length but showed a distinct pattern in
the growing domain, suggesting that differential organization of actin was required for tube elongation. Similar
results were obtained using microwave-accelerated DMSOpermeabilisation and TRITC-phalloidin staining (HeslopHarrison and Heslop-Harrison, 1991).
How do the distribution of myosins and the organization
of actin filaments correlate with organelle transport and
zonation in pollen tubes? Zonation is a way of describing
the different domains of pollen tubes based on their
membrane content: the vesicular, organelle, nuclear, and
vacuole regions (Cresti et al., 1977). However, these areas
are not separated by defined boundaries and exchange of
materials among them is a prerequisite for growth. Thus,
zonation is the result of dynamic activities that, for
example, constrain vesicles in the tip region and prevent
large-sized organelles from entering. Correlations between
actin filaments and organelle distribution were analysed by
Heslop-Harrison and Heslop-Harrison (1989a), who found
that cytochalasins B and D affected pollen tube growth and
rapidly stopped organelle circulation and movement of
vegetative nuclei and generative cells. A striking characteristic of organelle circulation in pollen tubes is their tipward
movement along the tube flanks and backward transport in
the centre (the so-called ‘reverse fountain’). The reversal of
organelle flow (which occurs in the apical 5–15 lm of
growing pollen tubes) prevented large-sized organelles (like
lipid globuli and amyloplasts) from penetrating the apical
region but allowed them to arrest and reverse their
movement in the subapical domain (Heslop-Harrison and
Heslop-Harrison, 1990). These findings suggested that the
subapical region contained a sort of filter that discriminated
the different organelles. Based on cytological observations,
the main candidate for filtering activity was the actin
cytoskeleton, described as a network of cross-linked actin
filaments in the tip and ordered, orientated bundles in the
older parts. The inability of large-size organelles to enter the
apex was also presumed to depend on the dense mass of tiplocalized vesicles (Pierson et al., 1990). Vacuoles in the
central zone were observed to be very motile and the
cytoplasmic strands in the vacuolar region contained
elements of the endoplasmic reticulum along which organelles moved in a saltatory manner (Pierson et al., 1990).
These findings suggested that the endoplasmic reticulum
was associated with actin filaments, which, in turn,
supported the movement of organelles. This speculative
hypothesis would later be confirmed using GFP-tagged
Golgi bodies (Nebenführ and Staehelin, 2001). The relationships between endoplasmic reticulum and Golgi bodies were
also investigated. Pollen tubes treated with Brefeldin A
showed dense cytoplasm containing tubular reticulum in the
tip domain and rapidly dissociating and disappearing Golgi
bodies, which fused with the endoplasmic reticulum (Rutten
and Knuiman, 1993).
The indecipherable role of microtubules
Most of the evidence discussed above indicated that actin
filaments were the main pathways along which organelles
and vesicles moved in the pollen tube. The basic evidence
that colchicine did not affect pollen tube growth was
indirect proof that microtubules were not critical for
organelle transport and tube elongation. However, a number
of researchers were investigating the distribution and
putative function of microtubules in pollen. Aström (1992)
showed that most pollen tube microtubules contain tyrosinated a-tubulin whereas acetylated a-tubulin was detected
in the generative cell, kinetochore fibres, and cell plate of
the phragmoplast; acetylation of a-tubulin was thought to
498 | Cai and Cresti
be correlated with more stable microtubules. A breakthrough in the pollen microtubule world was the immunological identification of a 105 kDa kinesin-related protein in
Nicotiana tabacum (tobacco) pollen tubes (Tiezzi et al.,
1992). The kinesin-related homologue of pollen tubes
localized in the apical and subapical regions of pollen tubes,
probably in association with vesicular material. The protein
also co-localized with pollen tube microtubules (Cai et al.,
1993). Further confirmation of the presence of kinesinrelated proteins came from studies in Corylus avellana
pollen, which contained a 100 kDa kinesin-related protein
associated with Golgi vesicles (Liu et al., 1994). The
presence of kinesin-related proteins in pollen tubes cast
doubts on the role of microtubules in organelle transport,
however, detailed observations on the distribution of microtubules in tobacco pollen tubes did not clarify the presence
of microtubules in the tip domain and their involvement in
vesicle transport (Del Casino et al., 1993). In addition, the
absence of microtubule-based motility data weakened the
preliminary indications on the role of microtubules in
vesicle transport.
The microtubule-motor story became a little more complicated a couple of years later when dynein-related polypeptides were identified in tobacco pollen tubes (Moscatelli
et al., 1995) associated with pollen tube organelles (Moscatelli
et al., 1998), indicating that the microtubule motor repertoire of pollen tubes may comprise different motor classes.
Although the presence of dynein in plants is still debated,
dyneins seem to have been lost from higher plants
(Wickstead and Gull, 2007) and their role seems to have
been replaced by the large kinesin family. Although the
microtubule story was highly promising, the indisputable
lack of colchicine- and oryzalin-related effects on tube
elongation was a crucial obstacle for these preliminary
discoveries. To be more precise, oryzalin was reported to
affect the pulsatory growth of pollen tubes (Geitmann et al.,
1995) and generative cell movements (Aström et al., 1995),
but these findings were hardly related to organelle movement
and motors.
Multiple myosins?
Although actin–myosin interactions were considered to play
a major role in organelle transport in pollen tubes, the only
myosin identified was a 175 kDa polypeptide detected by
heterologous antibodies (Tang et al., 1989a). Like other cell
systems, it seemed possible that pollen tubes contained
different classes of myosins, putatively involved in different
functions. Using antibodies to different myosin classes,
Miller et al. (1995) observed that myosin-I seemed to move
the generative cell and vegetative nucleus along the pollen
tube, while myosin-V moved small organelles (vesicles) and
myosins-I and II moved large organelles in two directions.
A different antibody against heavy meromyosin detected
a 174 kDa polypeptide associated with cytoplasmic organelles of tobacco pollen tubes, most fluorescent spots being
concentrated in the tip region below the plasma membrane.
This finding was also confirmed by immunoelectron microscopy that revealed gold particles around vesicle-like structures (Tirlapur et al., 1995). To sum up, data obtained in
the middle 1990s indicated the presence of various myosins,
whose different staining patterns were probably due to the
use of heterologous antibodies and the cross-reactivity of
antibodies with different myosin classes. A unified model
was missing.
Immunological detection was not the only way to reveal
myosin-like proteins in pollen tubes. Kohno et al. (1990)
had already described the main biochemical properties of
one pollen myosin; the same research group was able
partially to purify one myosin polypeptide (Kohno et al.,
1992) and isolate the complete 170 kDa motor protein
(Yokota and Shimmen, 1994). The myosin of Lilium longiflorum (lily) pollen tubes showed properties typical of the
myosin family and was localized as spots in the tip domain
of Lilium longiflorum pollen tubes and throughout the
pollen tubes of tobacco (Yokota et al., 1995). These results
were the first evidence of a pollen-specific myosin and
suggested that the protein might be associated with, and
involved in trafficking of different organelle classes. It was
evident that angiosperm pollen tubes contained at least one
myosin of 170 kDa; proofs for other myosins essentially
depended on the antibody used for detection. The presence
of different myosins in pollen tubes was also suggested
again more recently (Lovy-Wheeler et al., 2007) after
evidence that different organelle classes of pollen tubes
move differentially. Since the genome of Arabidopsis
thaliana contains 17 myosins belonging to families VIII and
XI (Lee and Liu, 2004), pollen tubes presumably contain
different members of the myosin XI family, each involved in
the movement of different organelles and vesicles. Clearly,
interpretation of the function of myosin(s) in pollen tubes is
limited by the absence of genetic data. The presence of
genes coding for myosin VIII and XI in the plant genome
(Jiang and Ramachandran, 2004) suggests that the previous
localization of myosin-I, II, and V should be re-examined
in a new light. For example, the previously identified 175
kDa (Tang et al., 1989a) and 174 kDa myosins (Tirlapur
et al., 1995) could correspond to the 170/175 kDa myosins
identified later (Yokota et al., 1995), assuming that polyclonal antibodies to myosin-II cross-react with plant
myosin-XI. Since there is functional homology between
animal myosin-V and plant myosin-XI (Li and Nebenführ,
2007), antibodies to myosin-V may indirectly indicate that
plant myosin-XI moves organelles and vesicles in the pollen
tube (Miller et al., 1995). Myosin-VIII is yet to be
discovered and may be responsible for the endocytotic
process in the pollen tube apex (Golomb et al., 2008). The
binding of different myosins with specific organelles is
a further fascinating research field; myosins probably
associate with the membrane of plant organelles by means
of Rab proteins (Hashimoto et al., 2008), which regulate the
binding and, consequently, the activity of motor proteins.
In any case, to learn more about myosin functions, cDNAs
encoding myosins expressed in pollen tubes need to be
cloned and used for functional studies.
Organelle movement in pollen tubes | 499
Fast progress on actin dynamics through
gene cloning and GFP technology
In the second half of the 1990s, advances in molecular
biology techniques opened new research perspectives in the
field of organelle movement. Immunological techniques had
been extremely important in uncovering the distribution of
cytoskeletal proteins, however, alternative procedures were
needed to obtain new information. Techniques of genetic
engineering and DNA cloning allowed researchers to move
from basic cytoskeletal proteins to the discovery of many
different accessory proteins involved in actin dynamics.
Analysis of organelle movement made progress through the
observation of single organelles after labelling with specific
dyes, exploiting specific computer software. Researchers
inferred that movements of membraneous structures were
random in the pollen tube tip but were more directed in
older cell regions (de Win et al., 1998). These findings
indicated indirectly that the organization of the actin
cytoskeleton changed from the tip to the base of the pollen
tubes. The differential distribution of actin in living pollen
tubes was also revealed by genetic transformation with
GFP-labelled actin-binding proteins (reviewed in Wilsen
et al., 2006). However, a characteristic pattern of labelling
was obtained with certain probes. High expression levels of
probes also caused structural rearrangements of actin
filaments and inhibited growth. Although these markers are
important technological advances, their use does not
yet allow complete visualization of the actin cytoskeleton in
growing pollen tubes and may cause artefacts and prevent
pollen tube growth.
At the same time, interest in pollen tubes was captured
by the study of mechanisms regulating growth and transduction of external signals into internal processes. In this
context, analysis of ion influx and dynamics provides
insights into the relationships between ion concentrations
and actin filaments. Dissipation of the tip-focused Ca2+
gradient enhanced entry of large organelles into the tube
tip (Pierson et al., 1994), suggesting that Ca2+ controlled the
organization of actin filaments, as proposed by Kohno and
Shimmen (1987). Another paper described the presence of
an alkaline band delimiting the apical domain and defining
the acidic tip of pollen tubes (Feijo et al., 1999). It was
suggested that the apical acidic domain correlated with the
growth process, probably facilitating vesicle movements and
exocytosis. The alkaline band probably also represented the
area where organelles reversed their motion (‘reverse
fountain’) by regulating the assembly of actin filaments
through pH-sensitive actin-binding proteins. Reversal of
organelle motility in the subapical region presumably
required appropriate local organization of actin filaments.
In this context, Li et al. (2001) described the presence of
circular F-actin bundles in pollen tubes, missing in the
initial 10-20 lm of the tubes and well correlated with
cytoplasmic streaming of organelles. Although the circular
F-actin bundles were not further described, they could
speculatively represent the track for bidirectional cytoplasmic streaming in pollen tubes.
The so-called former ‘actin foci’ represented an attempt
to associate actin dynamics with extracellular signals as in
animal cells. To understand the dynamics of actin filaments
in relationships with external cues it was necessary to
discover proteins that mediate the response of actin to
extracellular signals. Rop (‘Rho-of-plants’) GTPases were
found in the growth sites of pollen tubes and were assumed
to control polar growth (Zheng and Yang, 2000). RopGTPases influence the dynamics of tip-localized actin in
pollen tubes by transforming short actin bundles of the
apical domain into a filamentous network; overexpression
of Rop-GTPases also induces the formation of transverse
actin bands behind the tip (Fu et al., 2001). Speculatively,
external signals are converted by tip-localized Rop-GTPases
into messengers that modulate the organization of actin
filaments and thus the motility of organelles and the growth
of pollen tubes.
The Rop-mediated equilibrium between G- and F-actin
requires the activity of accessory intermediate proteins.
Villin was identified in lily pollen tubes and localized in
association with actin bundles (Vidali et al., 1999). Villins
are Ca2+-dependent, severing, capping, and nucleating
proteins and may participate in F-actin fragmentation and
nucleation in the pollen tube apex following activation by
the Ca2+ gradient. Like villin, LdABP41 is a Ca2+dependent protein that cuts actin filaments into short filaments and may operate in combination with the tip-focused
Ca2+ gradient (Fan et al., 2004). Villin also stimulates the
formation of unipolar bundles of actin filaments. Pollen
tubes combine villin with gelsolin (Huang et al., 2004;
Xiang et al., 2007) and profilin (Vidali and Hepler, 1997) to
organize actin filaments into two different and partly
autonomous arrays (the actin cables in pollen tube shanks
and shorter filaments in the apical and subapical regions),
required for the processes of organelle transport and tube
growth. Although tip growth is more sensitive to actin
inhibitors than cytoplasmic streaming, the two activities are
related because tip growth requires a continuous supply of
secretory vesicles, indicating that pollen tube growth
requires the assembly of different subsets of actin filaments,
which have different sensitivity to actin drugs (Vidali et al.,
2001). Consequently, pollen tube growth is the result of
cytoplasmic streaming, accumulation of secretory vesicles in
the tip domain (which depends on longitudinal actin
filaments) and the rate of vesicle fusion (which may require
a different actin array).
The number of actin-modulating proteins of pollen tubes
discovered increased rapidly. Actin-depolymerizing factors
(ADF) were identified as proteins showing pH-sensitive
actin-binding and depolymerizing activity (Allwood et al.,
2002). The alkaline band in pollen tubes was therefore
postulated to enhance the activity of ADFs and shift
actin toward the monomeric form. Thus the tip-focused
Ca2+ gradient and the alkaline band are critical markers of
reverse streaming activity (Cardenas et al., 2005). The
organization of actin filaments was also found to depend on
the activity of formin, a group of actin-nucleating proteins
that enhanced the formation of actin cables in the pollen
500 | Cai and Cresti
tube (Cheung and Wu, 2004) and promoted reversal of
streaming in the apical domain. The entire process of pollen
tube growth (comprising organelle/vesicle transport and
endo/exocytosis) necessarily required equilibrium to be maintained between G- and F-actin. Recognizing the number and
role of the molecular actors, which ultimately regulate actin
dynamics, and their relationships with differential ion
concentrations is a crucial forthcoming challenge.
Dynamics of specific organelle classes
Analysis of actin dynamics was accompanied by the
visualization of individual membrane classes in living pollen
tubes using specific markers. For example, FM4-64,
a marker of vesicles in the tube apex, highlighted the
organization of secretory vesicles in the ‘apical clear zone’
and revealed distinct pathways of vesicle movement toward
and away from the apex (Parton et al., 2001). Vesicle
distribution was found to fluctuate in the apical domain of
pollen tubes, suggesting that accumulation of secretory
vesicles in the pollen tube apex is not continuous but shows
a periodicity that depends on the actin cytoskeleton but not
on the secretion rate (Parton et al., 2003). Again, these
findings suggested that organelle and vesicle motility is
a process unrelated to secretion and growth. Visualization
of GFP-Rab11b-tagged vesicles in the inverted conical
region of tobacco pollen tubes after Latrunculin treatment
again indicated that accumulation of vesicles in the tip is
closely linked to the actin cytoskeleton (de Graaf et al.,
2005). Other cell compartments (specifically, endoplasmic
reticulum, mitochondria, and vacuoles) were monitored
during pollen tube growth and their movement was shown
to depend on actin filaments but apparently not on microtubules. Movement of endoplasmic reticulum followed the
direction of the main cytoplasmic streaming in pollen tubes,
traveling apically along the pollen tube flanks and reversing
its motion in the ‘actin fringe’ domain. Mitochondria were
found predominantly in the subapical region and showed
a movement similar to that of the endoplasmic reticulum. In
contrast, the vacuole was formed by thin longitudinal
tubules undergoing active motion but never extending into
the apical domain (Lovy-Wheeler et al., 2007). Although
cytoplasmic streaming generated the circular motion of
membraneous structures throughout the pollen tube, the
above findings showed that organelle distribution was
neither uniform nor accidental but precisely controlled. A
key question now is whether different myosins or different
regulation of the same myosin may be responsible for
differential organelle distribution.
Organelles and vesicles also move along
microtubules
Apart from preliminary observations on putative kinesinrelated protein in pollen tubes, many findings suggested that
a role of microtubules during in vitro pollen tube growth was
unnecessary. Evidence that colchicine and oryzalin treatment
caused few or no effects on the elongation rate of pollen
tubes led to the consideration that actin filaments were the
only motor system supporting organelle transport. More
consistent evidence that microtubules may take part in
organelle movement came with the discovery of pollen tube
proteins showing microtubule-gliding ability. A 90 kDa
kinesin-like protein from tobacco pollen tubes was identified
after screening of proteins co-pelleting with microtubules and
reacting with a peptide antibody against a highly conserved
region in the motor domain of the kinesin superfamily (Cai
et al., 2000); the purified 90 kDa protein induced microtubules to glide in motility assays. This finding was followed
by evidence that organelles from tobacco pollen tubes were
capable of moving with different velocities along microtubules in vitro (Romagnoli et al., 2003). A 105 kDa
organelle-associated motor was identified as functionally,
biochemically, and immunologically related to kinesin. This
indicated that the movement of pollen tube organelles was
not exclusively actin-based but also depended on microtubule-based motors. Further support for this finding came
from evidence that isolated pollen tube mitochondria moved
along microtubules and actin filaments in vitro (Romagnoli
et al., 2007). Movement was slow and continuous along
microtubules but very fast and irregular along actin filaments. Results also indicated that pollen vesicles and
mitochondria bound to identical myosins but specific kinesins, because the 90 kDa kinesin was found in association
with mitochondria but not with Golgi vesicles. The sum of
these elements suggested that microtubules and kinesins
contributed to the disposition of mitochondria in pollen
tubes. Similar results were also found in cells of Arabidopsis
thaliana (Ni et al., 2005). The presence of an additional
kinesin-like protein associated with Golgi bodies of Nicotiana
pollen tubes (Wei et al., 2005) further sustained the model
that organelles also move along microtubules and that pollen
tubes may contain different microtubule-based motors.
Golgi-associated kinesins are not peculiar to pollen tubes,
also being found in Arabidopsis trichomes (Lu et al., 2005).
Although the contribution of microtubules and microtubulebased motors may be superfluous during pollen tube growth
in vitro, it is hypothesized that the role of microtubules in
organelle motility may be more essential in vivo, when pollen
tube growth is fastest (Joos et al., 1995).
Actin-microtubule and related proteins
The presence of an additional microtubule-dependent transport system suggested that actin filaments and microtubules
may co-operate in the motility of organelles and vesicles
during pollen tube growth. The information that the
velocity of membranes along the two filamentous systems
was drastically different indicated that the specific role of
actin- and microtubule-motors was different. The combined
use of myosin and kinesin to achieve the proper final
delivery of organelles (so-called ‘functional co-operation’)
Organelle movement in pollen tubes | 501
suggested (albeit not necessarily) that actin filaments and
microtubules communicated in some way. While the
organization of actin filaments has been described in great
detail and the cortical actin fringe is acknowledged as
a typical feature of growing pollen tubes (Lovy-Wheeler
et al., 2005), detailed information on the distribution of
microtubules in living pollen tubes is still lacking or under
progress (Cheung et al., 2008).
Microtubules and actin filaments are organized differently
in pollen tubes but also show some similarities. Both are
disposed in bundles along the main axis, with actin filaments
also being present in the cytoplasm while microtubules are
mostly found in the cell cortex. In the cortex, single actin
filaments are usually observed to be aligned with microtubules and elements of the endoplasmic reticulum; microtubules appear to be cross-linked to actin filaments and to
the plasma membrane (Lancelle et al., 1987). Close association of microtubules with actin filaments has also been
shown by immunogold labelling, indicating that microtubules may act as guide elements for actin filaments or vice
versa (Lancelle and Hepler, 1991). Co-localization of microtubules and actin filaments in the pollen tube cortex has also
been revealed by double labelling with fluorescent probes,
suggesting functional relationships with organelle movement
and the organization of the cell cortex (Pierson et al., 1989).
A schematic drawing of actin filaments and microtubules
(based on visualization with GFP-fluorescent probes) and
their hypothetical relationships in pollen tubes is shown in
Fig. 2A.
Although a kind of interplay between microtubules and
actin filaments probably exists, its implications are enigmatic
and controversial. Microtubules are probably not involved in
the capacity of pollen tubes to overcome mechanical
obstacles, whereas actin filaments seem very important.
Conversely, maintenance of the direction of growth in vitro
was susceptible to microtubule degradation whereas actin
filaments were less or non critical (Gossot and Geitmann,
2007). In addition, the organization of microtubules probably depended on actin filaments, while actin filaments seemed
to be independent of microtubules. These findings suggested
that microtubules and actin filaments were physically connected in some way. In this context, a microtubule-associated
protein, SB401, was identified in the pollen of Solanum
berthaultii and shown to bind to actin filaments (Huang
et al., 2007). The possible interplay between actin filaments
and microtubules and the concept of ‘functional co-operation’ possibly implied the need for proteins mediating interactions between actin filaments and microtubules and their
effects of their mutually dependent architecture on organelle
movement. Inadequate data on microtubule-associated proteins (MAPs) in pollen tubes makes any type of realistic
hypothesis even more difficult, although a search of the
literature and BLAST comparison of proteins brought forth
several MAP-homologous proteins expressed in Arabidopsis
thaliana and putatively involved in different functions
(Gardiner and Marc, 2003). MAPs have still to be demonstrated in pollen tubes and it is not clear whether they are
required for organelle movement. Results of in vitro motility
assays indicated that pollen tube organelles move constantly
along single microtubules, suggesting that microtubule bundles may not be necessary (Romagnoli et al., 2007). The same
ambiguity also concerns post-translational modification of
tubulin. In addition to microtubule stabilization, acetylated
and tyrosinated a-tubulins (Aström, 1992; Del Casino et al.,
1993) may possibly be correlated with the activity of motor
proteins and the regulation of microtubule-mediated trafficking of organelles, as proposed by Dunn et al., (2008). The
effects of MAP-mediated differential density of microtubules
on organelle movement is an open question.
Actin, vesicles and the regulation of
exocytosis/endocytosis
The final fate of Golgi-derived secretory vesicles in pollen
tubes is fusion with the apical plasma membrane. Relationships between endoplasmic reticulum, Golgi bodies, and
vesicles have been studied using Brefeldin A, which induced
subapical membrane accumulation derived from the backflow of apical vesicles and from Golgi-derived membranes.
The Brefeldin-induced membrane aggregation showed periodic movements, which were not related to periodic
fluctuations of Ca2+. This evidence suggests that the Golgi
system is a central element in the production and recycling
of vesicular materials; these activities depend on the actin
cytoskeleton (Parton et al., 2003). Thus, the growth of
pollen tubes is essentially based on focusing Golgi-derived
vesicles that accumulate through the motor activity of the
actin–myosin complex and fuse in specific sites of the pollen
tube apex (polarized exocytosis). From the early work of
Steer and Steer (1989), it emerged that the number of
secretory vesicles in the pollen tube apex largely exceeded
the number effectively required for growth, suggesting that
excess plasma membrane must be recovered by other
mechanisms (endocytosis).
The relationship between vesicle trafficking and endo/
exocytosis is a critical aspect of pollen tube growth. Vesicle
streaming and tube growth can be unplugged by Latrunculin B (Cardenas et al., 2005), suggesting that vesicle
movement and fusion are separated by an undefined
boundary. Thus, current discussion about vesicle motility
(which is based on cytoskeleton-membrane interactions)
also needs to consider the subsequent phase of vesicle fusion
(which includes regulation and the dynamics of this process). Recent research indicated that exocytosis is mediated
by small (400 nm) vesicles and occurs in a region 3–10 lm
from the tube apex (Zonia and Munnik, 2008). This model
of subapex-delimited vesicular secretion has also been
suggested by other authors using spatiotemporal correlation
spectroscopy and fluorescence recovery after photobleaching; secretory vesicles were shown to move towards the apex
in the pollen tube cortex and accumulate close to the tip,
returning through the tube centre (Bove et al., 2008). The
velocity of secretory vesicles matched the speed of Golgi
vesicles as measured in vitro along actin filaments and
microtubules (Romagnoli et al., 2007). These findings imply
502 | Cai and Cresti
Fig. 2. (A) Schematic distribution of actin filaments and microtubules in a pollen tube as shown by GFP-tagged proteins. Actin filaments
are found in the cortical and central regions of the tube. They presumably arise as very short filaments from formin-containing membraneassociated proteins (only shown at the apex/subapex border); the actin fringe may represent a transition domain and a growth regiondelimiting domain; actin cables in pollen tube shanks are probably formed by bundle-forming proteins (such as villin). Microtubules exist
prevalently in bundles (formed by uncharacterized proteins) in the cortical cytoplasm and their organization in the subapex is still unclear;
presence of a microtubule fringe has been proposed (Lovy-Wheeler et al., 2007), but its organization and spatial relationships with the
actin fringe are unknown. Microtubules probably associate with the plasma membrane through p161 (Cai et al., 2005) and, hypothetically,
with actin filaments through uncharacterized proteins. Sites of origin and polarity of microtubules are not known. Monomers of actin and
tubulin are probably present in the apical region. Objects are not drawn to scale. (B) Regulation of cytoplasmic streaming. This picture is
derived from (A) but is essentially focused on the relationships between the signal transduction pathway and the pollen tube cytoskeleton.
Ca2+ is considered to be a main effector in the regulation of organelle movement. It enters the pollen tube in the tip region (1) and
generates a local increase in concentration (2), which also depends on the concerted activity of phospholipase C (PLC), phosphoinositol
biphosphate (PIP2), inositol triphosphate (IP3) (3), and of Rop GTPases activity (4). The oscillatory increase in Ca2+ concentration activates
the villin/gelsolin/profilin group, shifting the equilibrium of actin toward the monomeric form (5). This protein group works in combination
with ADFs (not reported) and the alkaline band, the latter produced by active efflux of H+ (6). The increase in Ca2+ concentration affects the
Organelle movement in pollen tubes | 503
relationships between the zone of active secretion and the
actin cytoskeleton. Vesicles are known to move along the
actin filaments, but the precise role of actin in the secretion
mechanism is still unclear. Identification of the actin fringe
in pollen tubes (Lovy-Wheeler et al., 2005) suggested the
hypothesis of its role in vesicle trafficking and fusion. Since
the actin fringe starts about 1–5 lm behind the apex and
extends for 5–10 lm, the region of the actin fringe
approximately coincides with the region of vesicle secretion.
Speculatively, exocytic vesicles may traffic directly to the
site of exocytosis through the actin fringe or the actin fringe
may somehow label or delimit the site of exocytosis. A
putative model may therefore be that accumulation of
secretory vesicles in the tube apex depends on longitudinal
actin filaments (Cardenas et al., 2008) while the actin fringe
probably acts as a final track for conveying vesicles
specifically to the site of growth. The apex and subapex of
pollen tubes may also possibly be the region of actin
assembly. New actin filaments probably arise from membraneassociated protein complexes containing formin. Overexpression of formin in pollen tubes induces the formation of
actin filaments, arising from the plasma membrane, in the
cytoplasm. Formin-dependent actin polymerization is therefore important for polar growth of pollen tubes (Cheung
and Wu, 2004). Actin filaments almost certainly arise from
the plasma membrane as very short filaments and then
organize into the actin fringe, which is therefore a transition
domain between growing actin filaments in the apex/
subapex and longitudinal actin cables. The actin fringe may
also act as a functional domain to delimit the region of
growth. Little is known about the organization of microtubules in the pollen tube apex. Despite evidences that
microtubules are absent from the apical domain, GFPlabelled EB1 indicated that microtubules may lie closer to
the pollen tube tip than previously thought (Cheung et al.,
2008). The presence of microtubules in close proximity to
the tip may also depend on the bending features of pollen
tubes (Foissner et al., 2002) suggesting a role of microtubules in the guiding process. The presence of a microtubule fringe in the subapex of pollen tubes was suggested by
microtubule labelling with fluorescent taxol (Lovy-Wheeler
et al., 2007) but the implications for growth and relationships with the actin fringe are unclear.
The exocytotic event is closely related to the opposite
process of endocytosis, which most likely occurs in two
distinct regions of the pollen tube, along the apex, and in
the distal region (Zonia and Munnik, 2008; Moscatelli
et al., 2007). In this context, hydrodynamic flux was
assumed to regulate exocytosis and endocytosis because
hypotonic treatment (and thus cell swelling) stimulated
exocytosis and reduced endocytosis, while hypertonic treatment (and cell shrinking) stimulated endocytosis but decreased exocytosis. The growth rate of pollen tubes is,
consequently, the result of fine-tuning of the apical volume
of pollen tubes, which influences the activity of endocytosis
(at the apex and along the distal tube) and exocytosis (in the
subapical region) and thus defines the boundary of the
growth zone. Visualization of endocytosis by fluorescent
phospholipids (Lisboa et al., 2008) indicated that the
plasma membrane retrieved was re-integrated into the
internal membrane pool and redistributed at the base and
apex of the pollen tube. Depolymerization of actin filaments
abolished plasma membrane retrieval, whereas oryzalin had
no effect, suggesting that actin filaments but not microtubules may be responsible for the endocytotic process.
Exocytotic/endocytotic activity is closely related to the
process of pollen tube growth, which is, in turn, regulated by
several external cues and intracellular messengers. The latter
presumably mediate the effects of growth regulators by
affecting the rate of vesicle fusion and the delivery of vesicles
to the apical domain through modulation of the actin
cytoskeleton. Among the different molecules that establish
signal transduction, Ca2+ was suggested to play a key role in
vesicle-dependent secretion of cell wall components, while
Rop-GTPases may play a crucial role in the fusion of
secretory vesicles and endocytosis (Camacho and Malho,
2003). Other authors have reported that Rop-GTPase
activity influences F-actin dynamics in the pollen tube tip
(Fu et al., 2001). Such activity probably depends on two
counteracting downstream pathways involving Rop-interactive CRIB-containing proteins (RICs), which contain a binding motif to Cdc42/Rac (Wu et al., 2001). Specifically, it was
found that RIC4 (overexpression of which promoted actin
assembly) determined the accumulation of secretory vesicles
but inhibited exocytosis; on the other hand, disassembly of
actin by RIC3 was required for exocytosis (Lee et al., 2008).
This again showed that the process of vesicle transport in the
tube tip and exocytosis depend on distinct mechanisms and
that fine-tuning of the equilibrium between F- and G-actin is
essential for tube growth. The activity of Rop-GTPases is
probably mediated by protein kinase activity. The presence
of type 2A protein phosphatase inhibitors had dramatic
effects on the cytoskeleton (randomly oriented actin filaments and microtubules, alteration of actin bundle alignment, and disassembly of microtubules) (Foissner et al.,
2002), which were similar to those observed after overexpression of Rop1 (Fu et al., 2001). Phosphatidylinositol
4,5-bisphosphate, inositol 1,4,5-trisphosphate, and phosphatidic acid have also been suggested to participate in the
control of vesicle fusion/retrieval and to modulate the
motor activity of myosin (7) through the associated calmodulin light chain (not shown). Regulation of actin filaments and inactivation of
myosin prevent large-sized organelles from entering the apical domain (8). Secretory vesicles also move along actin filaments using myosin
activity (9) but it is not known whether this myosin is affected by variations in Ca2+ concentrations. Vesicles enter and accumulate in the
apical domain (10) and fuse with the plasma membrane (11) or turn back toward the grain (12). Nothing is known about the regulation of
microtubules, but they could be regulated by phospholipase D (not shown). The polymerization state of actin filaments may also regulate
microtubule dynamics (13). Objects are not drawn to scale.
504 | Cai and Cresti
organization of actin filaments (Monteiro et al., 2005a).
Expression of a phospholipase C (PLC) in pollen tubes of
Petunia inflata raised the apical Ca2+ gradient, which
disorganized the actin filaments (Dowd et al., 2006). This
finding suggested that PLC hydrolysed phosphatidylinositol
4,5-bisphosphate to the signalling molecules inositol 1,4,5trisphosphate and diacyl glycerol (DAG). Inositol 1,4,5trisphosphate is thought to regulate intracellular Ca2+
concentrations, which are known to affect actin organization.
The negative effects of expression of PLC were reversed by
treatment with low concentrations of the actin drug Latrunculin B, suggesting that PLC alters actin structure in growing
tube tips and restricts growth to the very tip of pollen tubes.
Overexpressed PLC also inhibited tube growth and accumulated laterally at the pollen tube tip plasma membrane
mimicking the distribution of phosphatidylinositol 4,5bisphosphate. Diacylglycerol (DAG) also showed a localization similar to that of phosphatidylinositol 4,5-bisphosphate
(Helling et al., 2006). Phospholipase D (PLD) and its
product phosphatidic acid (PA) also play a role in the
polarization of pollen tubes and their activity can be blocked
by primary alcohols, like 1-butanol. When inhibited by 1butanol, pollen tube growth can be restored by taxol
treatment, indicating that microtubules may be one target of
PLD activity. This hypothesis is supported by the finding
that PLD is linked to microtubule dynamics (Dhonukshe
et al., 2003).
The role of microtubules (if any) in vesicle targeting and
fusion is unclear. Although kinesin-related proteins have
been localized in apical vesicles (Tiezzi et al., 1992),
disorganization of microtubules does not influence pollen
tube growth. Nevertheless, the evidence that oryzalin
treatment affected pulsatory growth (Geitmann et al., 1995)
and directional growth of pollen tubes (Gossot and
Geitmann, 2007) indirectly suggests that microtubules may
interfere with the growth mechanism. In addition, the
finding that plant phospholipase D associates with membranes and microtubules (Gardiner et al., 2001), coupled
with the effects of protein phosphatase inhibitors on pollen
tube microtubules (Foissner et al., 2002), indicates interplay
between the signal transduction pathway and the microtubule cytoskeleton.
On the basis of the above information, it is possible to
postulate relationships between the actin cytoskeleton and
the endo/exocytotic process (Fig. 2B). Ca2+ is presumed to
enter pollen tubes at the very tip (1) increasing its local
concentrations (2) (Messerli and Robinson, 1997). Variation
in Ca2+ concentrations is also thought to increase after
stimulation of the phospholipase C (PLC), phosphoinositolbiphosphate (PIP2) and inositol-triphosphate (IP3) pathway
(3) (Monteiro et al., 2005b). Rop-GTPases are also believed
to regulate Ca2+ concentrations (4) (Hwang et al., 2005).
Local oscillatory increases in Ca2+ concentration are
considered to activate the protein group consisting of villin,
gelsolin, and profilin, shifting the equilibrium towards Gactin and preventing polymerization/elongation of actin
filaments (5) (Yokota et al., 2005). This protein group
probably works in combination with ADFs (not reported in
the figure) and the alkaline band, which is probably
produced by active discharge of H+ (6) (Chen et al., 2002).
An increase in Ca2+ concentrations also affects the motor
activity of myosin (7) through the associated calmodulin
light chain (not shown) (Yokota et al., 1999). Regulated
elongation of actin filaments along with the inactivation of
myosin probably prevents large-sized organelles from entering the apical domain (8) but allows them to turn back.
Secretory vesicles also move along actin filaments using the
motor activity of myosin (9). It is not known whether
vesicle-associated myosin is affected by increases in Ca2+
concentrations; in any case, vesicles penetrate and accumulate in the apical domain (10) where they fuse with the
plasma membrane (11). Vesicles that do not fuse may return
to the subapical region, together with endocytic vesicles
(12); this tip-to-subapex movement may rely on actin and
myosin. Nothing is known about the regulation of microtubule activities; speculatively, they may be regulated by
phospholipase D (not shown), or the polymerization state
of actin filaments may regulate microtubule dynamics (13)
(Gossot and Geitmann, 2007; Poulter et al., 2008).
Conclusions and future prospects
Advances in the last 20 years of research have offered
important new insights into the functions of actin filaments
and microtubules in organelle transport inside pollen tubes.
Since the early evidence of organelle movement along actin
filaments, current research is now focusing on the relationships between actin/tubulin dynamics, organelle/vesicle
transport, and pollen tube growth. A great amount of data
indicates that actin filaments are the main tracks of
organelle and vesicle transport; actin filaments support
cytoplasmic streaming in the basal domain and accumulation of vesicles in the tip domain. External signals appear to
be critical for redirecting tip growth when the pollen tube
has to follow signals from female tissues; external signals
presumably modulate the activity of motor proteins and
organization of actin filaments. The role of microtubules is
still unclear, although their function could be more
prominent for the growth of pollen tubes in vivo. Whether
actin and the microtubule cytoskeleton co-operate in motor
activity is still unclear and there are major challenges to be
solved in the near future. Some of these questions are listed
below.
(1) Do organelles of the same class move independently? The motility pattern of mitochondria, Golgi
bodies, endoplasmic reticulum, and vesicles needs to be
clarified. We do not know whether single organelles
move independently of the others or ignore whether or
not the trafficking of each organelle class depends on
that of other classes. Do specific motor receptors exist?
Does this imply involvement of multiple myosins?
(2) Is organelle transport regulated by functional cooperation? It is current opinion that organelle transport
in eukaryotic cells is achieved by regulating the activities
Organelle movement in pollen tubes | 505
Fig. 3. Hypothetical interactions of pollen tube organelles with both actin filaments and microtubules. Speculatively, organelles move
rapidly along actin filaments using myosin as motor protein; organelles may be stopped in specific sites of the pollen tube for finer
positioning through dynamic interaction of kinesin with microtubules. Microtubule-actin interacting proteins are hypothetical; they may
help organize cytoskeletal architecture but their role in organelle transport is hard to decipher. The different sized arrows indicate the
approximate intensity of motility along actin and microtubules. Objects are not drawn to scale.
of motor proteins of different families (Goode et al.,
2000). This model has been the subject of relatively little
study in plant cells, although there is evidences that
myosin and kinesin may ultimately be required for
organelle trafficking (Wei et al., 2005). Since organelles
interact dynamically with actin filaments and microtubules, the pollen tube may be an excellent model for
studying this process; unfortunately, we do not know
how many myosins and kinesins are present in pollen
tubes, and how motor proteins co-operate in organelle
and vesicle movement. A speculative basic model is
reported in Fig. 3.
(3) What is the relationship between external signals,
motor-mediated organelle/vesicle delivery, and pollen tube
growth? Delivery of vesicles to the apical region supports
pollen tube growth, which is under the control of external
signals from female tissues (Herrero, 2001). Although
indications suggest that myosin activity in pollen tubes is
controlled by Ca2+ concentrations (Yokota et al., 1999),
no direct evidence as yet links the signal transduction
pathway to motor activity. Does control of motor activity
simply rely on the regulation of cytoskeletal filaments?
(4) What is the exact role of microtubule motors in
organelle/vesicle transport? There is evidences that pollen
tube organelles and vesicles move along microtubules
(Romagnoli et al., 2007) and that microtubules may
affect the pulsed directional growth of pollen tubes
(Gossot and Geitmann, 2007; Geitmann et al., 1995);
however, more information is needed on the role of
microtubule motors in these processes. Gene sequences
of kinesins and inhibition of expression by iRNA may
provide some answers.
(5) What is the effect of localized vesicle delivery on cell
wall construction in pollen tubes? In order to appreciate
the contribution of the cytoskeleton to the pollen tube
shaping process, it is essential to determine the secretion
pattern of cell wall-synthesizing enzymes, their activation
scheme, and dependence on exocytosis and actin filaments or microtubules. Is proper organization of the
cytoskeleton required for pollen tube cell wall construction? How do these activities depend on molecular
motors?
(6) What is the relationship between the actin fringe, the
alkaline band, and secretion of vesicles? This feature
merits considerable attention because the actin fringe
(presumably under the control of the alkaline band) is
a key element in vesicle secretion. Does the actin fringe
define the secretion zone or does it drive secretory vesicles
to the secretion site? What is the role (if any) of the actin
fringe during endocytosis? Where does the responsibility
of myosins end in the transport of secretory vesicles?
Acknowledgements
We sincerely thank Professor Rui Malhó (Department de
Biologia Vegetal, FCL, Campo Grande, 1749-016 Lisboa,
Portugal) for his kind revision of the manuscript and for
helpful suggestions.
References
Allwood EG, Anthony RG, Smertenko AP, Reichelt S,
Drobak BK, Doonan JH, Weeds AG, Hussey PJ. 2002. Regulation
of the pollen-specific actin-depolymerizing factor LlADF1. The Plant
Cell 14, 2915–2927.
Astrom H. 1992. Acetylated alpha-tubulin in the pollen tube microtubules. Cell Biology International Report 16, 871–881.
Astrom H, Sorri O, Raudaskoski M. 1995. Role of microtubules in
the movement of the vegetative nucleus and generative cell in tobacco
pollen tubes. Sexual Plant Reproduction 8, 61–69.
Bove J, Vaillancourt B, Kroeger J, Hepler PK, Wiseman PW,
Geitmann A. 2008. Magnitude and direction of vesicle dynamics in
growing pollen tubes using spatiotemporal image correlation spec-
506 | Cai and Cresti
troscopy and fluorescence recovery after photobleaching. Plant
Physiology 147, 1646–1658.
Cai G, Bartalesi A, Del Casino C, Moscatelli A, Tiezzi A,
Cresti M. 1993. The kinesin-immunoreactive homologue from Nicotiana tabacum pollen tube: biochemical properties and subcellular
localization. Planta 191, 496–506.
Cai G, Ovidi E, Romagnoli S, Vantard M, Cresti M, Tiezzi A.
2005. Identification and characterization of plasma membrane proteins
that bind to microtubules in pollen tubes and generative cells of
tobacco. Plant Cell Physiology 46, 563–578.
Cai G, Romagnoli S, Moscatelli A, Ovidi E, Gambellini G,
Tiezzi A, Cresti M. 2000. Identification and characterization of a novel
microtubule-based motor associated with membranous organelles in
tobacco pollen tubes. The Plant Cell 12, 1719–1736.
Camacho L, Malho R. 2003. Endo/exocytosis in the pollen tube
apex is differentially regulated by Ca2+ and GTPases. Journal of
Experimental Botany 54, 83–92.
Cardenas L, Lovy-Wheeler A, Kunkel JG, Hepler PK. 2008. Pollen
tube growth oscillations and intracellular calcium levels are reversibly
modulated by actin polymerization. Plant Physiology 146, 1611–1621.
Del Casino C, Li Y-Q, Moscatelli A, Scali M, Tiezzi A, Cresti M.
1993. Distribution of microtubules during the growth of tobacco pollen
tubes. Biology of the Cell 79, 125–132.
Dhonukshe P, Laxalt AM, Goedhart J, Gadella TWJ, Munnik T.
2003. Phospholipase D activation correlates with microtubule reorganization in living plant cells. The Plant Cell 15, 2666–2679.
Dowd PE, Coursol S, Skirpan AL, Kao T-h, Gilroy S. 2006.
Petunia phospholipase C1 is involved in pollen tube growth. The Plant
Cell 18, 1438–1453.
Dunn S, Morrison EE, Liverpool TB, Molina-Paris C, Cross RA,
Alonso MC, Peckham M. 2008. Differential trafficking of Kif5c on
tyrosinated and detyrosinated microtubules in live cells. Journal of Cell
Science 121, 1085–1095.
Fan X, Hou J, Chen X, Chaudhry F, Staiger CJ, Ren H. 2004.
Identification and characterization of a Ca2+-dependent actin filamentsevering protein from lily pollen. Plant Physiology 136, 3979–3989.
Feijo JA, Sainhas J, Hackett GR, Kunkel JG, Hepler PK. 1999.
Growing pollen tubes possess a constitutive alkaline band in the clear
zone and a growth-dependent acidic tip. Journal of Cell Biology 144,
483–496.
Cardenas L, Lovy-Wheeler A, Wilsen KL, Hepler PK. 2005. Actin
polymerization promotes the reversal of streaming in the apex of pollen
tubes. Cell Motility and Cytoskeleton 61, 112–127.
Foissner I, Grolig F, Obermeyer G. 2002. Reversible protein
phosphorylation regulates the dynamic organization of the pollen tube
cytoskeleton: effects of calyculin A and okadaic acid. Protoplasma
220, 1–15.
Chen CY, Wong EI, Vidali L, Estavillo A, Hepler PK, Wu HM,
Cheung AY. 2002. The regulation of actin organization by actindepolymerizing factor in elongating pollen tubes. The Plant Cell 14,
2175–2190.
Fu Y, Wu G, Yang Z. 2001. Rop GTPase-dependent dynamics of tiplocalized F-actin controls tip growth in pollen tubes. Journal of Cell
Biology 152, 1019–1032.
Cheung AY, Wu HM. 2004. Overexpression of an Arabidopsis formin
stimulates supernumerary actin cable formation from pollen tube cell
membrane. The Plant Cell 16, 257–269.
Gardiner JC, Harper JDI, Weerakoon ND, Collings DA, Ritchie S,
Gilroy S, Cyr RJ, Marc J. 2001. A 90 kDa phospholipase D from
tobacco binds to microtubules and the plasma membrane. The Plant
Cell 13, 2143–2158.
Cheung AY, Q-h Duan, Costa SS, de Graaf BHJ, Di Stilio VS,
Feijo J, Wu H-M. 2008. The dynamic pollen tube cytoskeleton: live
cell studies using actin-binding and microtubule-binding reporter
proteins. Molecular Plant 1, 686–702.
Condeelis JS. 1974. The identification of F actin of the pollen tube
and protoplast of Amaryllis belladonna. Experimental Cell Research 88,
435–439.
Cresti M, Blackmore S, van Went JL. 1992. Atlas of sexual
reproduction in flowering plants. Berlin Heidelberg: Springer-Verlag.
Cresti M, Pacini E, Ciampolini F, Sarfatti G. 1977. Germination
and early tube development in vitro of Lycopersicum peruvianum
pollen: ultrastrucutral features. Planta 136, 239–247.
Cresti M, VanWent JL, Willemse MTM, Pacini E. 1976. Fibrous
masses and cell and nucleus movement in the pollen tube of Petunia
hybrida. Acta Botanica Neerlandica 25, 381–383.
Gardiner JC, Marc J. 2003. Putative microtubule-associated proteins from the Arabidopsis genome. Protoplasma 222, 61–74.
Geitmann A, Li Y-Q, Cresti M. 1995. The role of cytoskeleton and
dictyosome activity in the pulsatory growth of Nicotiana tabacum and
Petunia hybrida pollen tubes. Botanica Acta 109, 102–109.
Golomb L, bu-Abied M, Belausov E, Sadot E. 2008. Different
subcellular localizations and functions of Arabidopsis myosin VIII..
BMC Plant Biology 8, 3.
Goode BL, Drubin DG, Barnes G. 2000. Functional co-operation
between the microtubule and actin cytoskeletons. Current Opinion in
Cell Biology 12, 63–71.
Gossot O, Geitmann A. 2007. Pollen tube growth: coping with
mechanical obstacles involves the cytoskeleton. Planta 226,
405–416.
de Graaf BH, Cheung AY, Andreyeva T, Levasseur K,
Kieliszewski M, Wu HM. 2005. Rab11 GTPase-regulated membrane
trafficking is crucial for tip-focused pollen tube growth in tobacco. The
Plant Cell 17, 2564–2579.
Hashimoto K, Igarashi H, Mano S, Takenaka C, Shiina T,
Yamaguchi M, Demura T, Nishimura M, Shimmen T, Yokota E.
2008. An isoform of Arabidopsis myosin XI interacts with small
GTPases in its C-terminal tail region. Journal of Experimental Botany
59, 3523–3531.
de Win AH, Pierson ES, Timmer C, Lichtscheidl IK, Derksen J.
1998. Interactive computer-assisted position acquisition procedure
designed for the analysis of organelle movement in pollen tubes.
Cytometry 32, 263–267.
Helling D, Possart A, Cottier S, Klahre U, Kost B. 2006. Pollen
tube tip growth depends on plasma membrane polarization mediated
by tobacco PLC3 activity and endocytic membrane recycling. The
Plant Cell 18, 3519–3534.
Organelle movement in pollen tubes | 507
Herrero M. 2001. Ovary signals for directional pollen tube growth.
Sexual Plant Reproduction 14, 3–7.
Heslop-Harrison J, Heslop-Harrison Y. 1988. Organelle movement
and fibrillar elements of the cytoskeleton in the angiosperm pollen
tube. Sexual Plant Reproduction 1, 16–24.
Heslop-Harrison J, Heslop-Harrison Y. 1989a. Cytochalasin
effects on structure and movement in the pollen tube of Iris. Sexual
Plant Reproduction 2, 27–37.
Heslop-Harrison J, Heslop-Harrison Y. 1989b. Myosin associated
with the surface of organelles, vegetative nuclei and generative cells in
angiosperm pollen grains and tubes. Journal of Cell Science 94, 319–
325.
Lancelle SA, Hepler PK. 1991. Association of actin with cortical
microtubules revealed by immunogold localization in Nicotiana pollen
tubes. Protoplasma 165, 167–172.
Lee YJ, Szumlanski A, Nielsen E, Yang Z. 2008. Rho-GTPasedependent filamentous actin dynamics coordinate vesicle targeting
and exocytosis during tip growth. The Journal of Cell Biology 181,
1155–1168.
Lee YRJ, Liu B. 2004. Cytoskeletal motors in Arabidopsis. Sixty-one
kinesins and seventeen myosins. Plant Physiology 136, 3877–3883.
Li JF, Nebenführ A. 2007. Organelle targeting of myosin XI is
mediated by two globular tail subdomains with separate cargo binding
sites. Journal of Biological Chemistry 282, 20593–20602.
Heslop-Harrison J, Heslop-Harrison Y. 1990. Dynamic aspects of
the apical zonation in the angiosperm pollen tube. Sexual Plant
Reproduction 3, 187–194.
Li Y, Zee SY, Liu YM, Huang BQ, Yen LF. 2001. Circular F-actin
bundles and a G-actin gradient in pollen and pollen tubes of Lilium
davidii. Planta 213, 722–730.
Heslop-Harrison J, Heslop-Harrison Y. 1991. The actin cytoskeleton in unfixed pollen tubes following microwave-accelerated DMSOpermeabilization and TRITC-phalloidin staining. Sexual Plant Reproduction 4, 6–11.
Lisboa S, Scherer GE, Quader H. 2008. Localized endocytosis in
tobacco pollen tubes: visualisation and dynamics of membrane
retrieval by a fluorescent phospholipid. Plant Cell Reports 27, 21–28.
Heslop-Harrison J, Heslop-Harrison Y, Cresti M, Tiezzi A,
Moscatelli A. 1988. Cytoskeletal elements, cell shaping and movement in the angiosperm pollen tube. Journal of Cell Science 91, 49–
60.
Huang S, Blanchoin L, Chaudhry F, Franklin-Tong VE,
Staiger CJ. 2004. A gelsolin-like protein from Papaver rhoeas pollen
(PrABP80) stimulates calcium-regulated severing and depolymerization
of actin filaments. Journal of Biological Chemistry 279, 23364–23375.
Liu G-Q, Cai G, Del Casino C, Tiezzi A, Cresti M. 1994. Kinesinrelated polypeptide is associated with vesicles from Corylus avellana
pollen. Cell Motility and the Cytoskeleton 29, 155–166.
Lovy-Wheeler A, Cardenas L, Kunkel JG, Hepler PK. 2007.
Differential organelle movement on the actin cytoskeleton in lily pollen
tubes. Cell Motility and the Cytoskeleton 64, 217–232.
Lovy-Wheeler A, Wilsen KL, Baskin TI, Hepler PK. 2005.
Enhanced fixation reveals the apical cortical fringe of actin filaments as
a consistent feature of the pollen tube. Planta 221, 95–104.
Huang S, Jin L, Du J, Li H, Zhao Q, Ou G, Ao G, Yuan M. 2007.
SB401, a pollen-specific protein from Solanum berthaultii, binds
to and bundles microtubules and F-actin. The Plant Journal 51,
406–418.
Lu L, Lee YRJ, Pan R, Maloof JN, Liu B. 2005. An internal motor
kinesin is associated with the Golgi apparatus and plays a role in
trichome morphogenesis in Arabidopsis. Molecular Biology of the Cell
16, 811–823.
Hwang JU, Gu Y, Lee YJ, Yang Z. 2005. Oscillatory ROP GTPase
activation leads the oscillatory polarized growth of pollen tubes.
Molecular Biology of the Cell 16, 5385–5399.
Messerli M, Robinson KR. 1997. Tip localized Ca2+ pulses are
coincident with peak pulsatile growth rates in pollen tubes of Lilium
longiflorum. Journal of Cell Science 110, 1269–1278.
Jiang S, Ramachandran S. 2004. Identification and molecular
characterization of myosin gene family in Oryza sativa genome. Plant
Cell Physiology 45, 590–599.
Miller DD, Scordilis SP, Hepler PK. 1995. Identification and
localization of three classes of myosins in pollen tubes of Lilium
longiflorum and Nicotiana alata. Journal of Cell Science 108, 2549–
2563.
Joos U, van Aken J, Kristen U. 1995. The anti-microtubule drug
carbetamide stops Nicotiana sylvestris pollen tube growth in the style.
Protoplasma 187, 182–191.
Kohno T, Chaen S, Shimmen T. 1990. Characterization of the
translocator associated with pollen tube organelles. Protoplasma 154,
179–183.
Kohno T, Ishikawa R, Nagata T, Kohama K, Shimmen T. 1992.
Partial purification of myosin from lily pollen tubes by monitoring with in
vitro motility assay. Protoplasma 170, 77–85.
Kohno T, Shimmen T. 1987. Ca2+-induced fragmentation of actin
filaments in pollen tubes. Protoplasma 141, 177–179.
Kohno T, Shimmen T. 1988. Accelerated sliding of pollen tube
organelles along Characeae actin bundles regulated by Ca2+. Journal
of Cell Biology 106, 1539–1543.
Lancelle SA, Cresti M, Hepler PK. 1987. Ultrastructure of cytoskeleton in freeze-substituded pollen tubes of Nicotiana tabacum. Protoplasma 140, 141–150.
Monteiro D, Castanho CP, Rodrigues C, Camacho L, Quader H,
Malho R. 2005a. Modulation of endocytosis in pollen tube growth by
phosphoinositides and phospholipids. Protoplasma 226, 31–38.
Monteiro D, Liu Q, Lisboa S, Scherer GEF, Quader H, Malho R.
2005b. Phosphoinositides and phosphatidic acid regulate pollen tube
growth and reorientation through modulation of [Ca2+]c and membrane
secretion. Journal of Experimental Botany 56, 1665–1674.
Moscatelli A, Cai G, Ciampolini F, Cresti M. 1998. Dynein heavy
chain-related polypeptides are associated with organelles in pollen
tubes of Nicotiana tabacum. Sexual Plant Reproduction 11, 31–40.
Moscatelli A, Ciampolini F, Rodighiero S, Onelli E, Cresti M,
Santo N, Idilli A. 2007. Distinct endocytic pathways identified in
tobacco pollen tubes using charged nanogold. Journal of Cell Science
120, 3804–3819.
Moscatelli A, Del Casino C, Lozzi L, Cai G, Scali M, Tiezzi A,
Cresti M. 1995. High molecular weight polypeptides related to dynein
508 | Cai and Cresti
heavy chains in Nicotiana tabacum pollen tubes. Journal of Cell
Science 108, 1117–1125.
Nebenführ A, Staehelin LA. 2001. Mobile factories: Golgi dynamics
in plant cells. Trends in Plant Science 6, 160–167.
Ni CZ, Wang HQ, Xu T, Qu Z, Liu GQ. 2005. AtKP1, a kinesin-like
protein, mainly localizes to mitochondria in Arabidopsis thaliana. Cell
Research 15, 725–733.
Parton RM, Fischer-Parton S, Trewavas AJ, Watahiki MK. 2003.
Pollen tubes exhibit regular periodic membrane trafficking events in the
absence of apical extension. Journal of Cell Science 116, 2707–2719.
Parton RM, Fischer-Parton S, Watahiki MK, Trewavas AJ. 2001.
Dynamics of the apical vesicle accumulation and the rate of growth are
related in individual pollen tubes. Journal of Cell Science 114, 2685–
2695.
Pierson ES. 1988. Rhodamine-phalloidin staining of F-actin in pollen
after dimethylsulfoxide permeabilization. a comparison with the conventional formaldeyde preparation. Sexual Plant Reproduction 1, 83–
87.
Pierson ES, Derksen J, Traas JA. 1986. Organization of microfilaments and microtubules in pollen tubes grown in vitro or in vivo in
various angiosperms. European Journal of Cell Biology 41, 14–18.
Pierson ES, Lichtscheidl IK, Derksen J. 1990. Structure and
behaviour of organelles in living pollen tubes of Lilium longiflorum.
Journal of Experimental Botany 41, 1461–1468.
Pierson ES, Miller DD, Callaham DA, Shipley AM, Rivers BA,
Cresti M, Hepler PK. 1994. Pollen tube growth is coupled to the
extracellular calcium ion flux and the intracellular calcium gradient:
effect of BAPTA-type buffers and hypertonic media. The Plant Cell 6,
1815–1828.
Pierson ES, Kengen HMP, Derksen J. 1989. Microtubules and
actin filaments co-localize in pollen tubes of Nicotiana tabacum L. and
Lilium longiflorum Thunb. Protoplasma 150, 75–77.
Poulter NS, Vatovec S, Franklin-Tong VE. 2008. Microtubules are
a target for self-incompatibility signaling in Papaver pollen. Plant
Physiology 146, 1358–1367.
Romagnoli S, Cai G, Cresti M. 2003. In vitro assays demonstrate
that pollen tube organelles use kinesin-related motor proteins to move
along microtubules. The Plant Cell 15, 251–269.
Romagnoli S, Cai G, Faleri C, Yokota E, Shimmen T, Cresti M.
2007. Microtubule- and actin filament-dependent motors are distributed on pollen tube mitochondria and contribute differently to their
movement. Plant Cell Physiology 48, 345–361.
Tiezzi A, Moscatelli A, Cai G, Bartalesi A, Cresti M. 1992. An
immunoreactive homolog of mammalian kinesin in Nicotiana tabacum
pollen tubes. Cell Motility and the Cytoskeleton 21, 132–137.
Tiezzi A, Pierson ES, Theunis CH, Ciampolini F, Cai G,
Bartalesi A, Cresti M. 1991. The motile apparatus of sperm cells in
angiosperms: correlations with lower plants, gymnosperms and
animals. In: Baccetti B, ed. Comparative spermatology 20 years after,
Vol. 75. Raven Press, 1017–1020.
Tirlapur UK, Cai G, Faleri C, Moscatelli A, Scali M, Del CC,
Tiezzi A, Cresti M. 1995. Confocal imaging and immunogold electron
microscopy of changes in distribution of myosin during pollen hydration, germination and pollen tube growth in Nicotiana tabacum L.
European Journal of Cell Biology 67, 209–217.
Vidali L, Hepler PK. 1997. Characterization and localization of profilin
in pollen grains and tubes of Lilium longiflorum. Cell Motility and the
Cytoskeleton 36, 323–338.
Vidali L, McKenna ST, Hepler PK. 2001. Actin polymerization is
essential for pollen tube growth. Molecular Biology of the Cell 12,
2534–2545.
Vidali L, Yokota E, Cheung AY, Shimmen T, Hepler PK. 1999.
The 135 kDa actin-bundling protein from Lilium longiflorum pollen is
the plant homologue of villin. Protoplasma 209, 283–291.
Wei L, Liu B, Li Y. 2005. Distribution of a kinesin-related protein on
Golgi apparatus of tobacco pollen tubes. Chinese Science Bullettin 50,
2175–2181.
Wickstead B, Gull K. 2007. Dyneins across eukaryotes: a comparative genomic analysis. Traffic 8, 1708–1721.
Wilsen K, Lovy-Wheeler A, Voigt B, Menzel D, Kunkel J,
Hepler P. 2006. Imaging the actin cytoskeleton in growing pollen
tubes. Sexual Plant Reproduction 19, 51–62.
Wu G, Gu Y, Li S, Yang Z. 2001. A genome-wide analysis of
Arabidopsis Rop-interactive CRIB motif-containing proteins that act as
Rop GTPase targets. The Plant Cell 13, 2841–2856.
Xiang Y, Huang X, Wang T, Zhang Y, Liu Q, Hussey PJ, Ren H.
2007. ACTIN BINDING PROTEIN 29 from Lilium pollen plays an
important role in dynamic actin remodeling. The Plant Cell 19, 1930–
1946.
Yokota E, McDonald AR, Liu B, Shimmen T, Palevitz BA. 1995.
Localization of a 170 kDa myosin heavy chain in plant cells.
Protoplasma 185, 178–187.
Yokota E, Muto S, Shimmen T. 1999. Inhibitory regulation of higherplant myosin by Ca2+ ions. Plant Physiology 119, 231–240.
Rutten TL, Knuiman B. 1993. Brefeldin A effects on tobacco pollen
tubes. European Journal of Cell Biology 61, 247–255.
Yokota E, Shimmen T. 1994. Isolation and characterization of plant
myosin from pollen tubes of lily. Protoplasma 177, 153–162.
Steer MW, Steer JM. 1989. Pollen tube tip growth. New Phytologist
111, 323–358.
Yokota E, Tominaga M, Mabuchi I, Tsuji Y, Staiger CJ, Oiwa K,
Shimmen T. 2005. Plant villin, lily P-135-ABP, possesses G-actin
binding activity and accelerates the polymerization and depolymerization of actin in a Ca2+-sensitive manner. Plant Cell Physiology 46,
1690–1703.
Tang XJ, Hepler PK, Scordilis SP. 1989a. Immunochemical and
immunocytochemical identification of a myosin heavy chain polypeptide in Nicotiana pollen tubes. Journal of Cell Science 92, 569–
574.
Tang XJ, Lancelle SA, Hepler PK. 1989b. Fluorescence microscopic localization of actin in pollen tubes: comparison of actin
antibody and phalloidin staining. Cell Motility and the Cytoskeleton 12,
216–224.
Zheng ZL, Yang Z. 2000. The Rop GTPase switch turns on polar
growth in pollen. Trends in Plant Science 5, 298–303.
Zonia L, Munnik T. 2008. Vesicle trafficking dynamics
and visualization of zones of exocytosis and endocytosis in tobacco
pollen tubes. Journal of Experimental Botany 59, 861–873.