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Journal of Experimental Botany, Vol. 54, No. 380,
Plant Reproductive Biology Special Issue, pp. 103±113, January 2003
DOI: 10.1093/jxb/erg003
The actin cytoskeleton is a target of the
self-incompatibility response in Papaver rhoeas
C. J. Staiger1 and V. E. Franklin-Tong2,3
Department of Biological Sciences, Purdue University, West Lafayette, IN 47907-1392, USA
School of Biosciences, University of Birmingham, Edgbaston B15 2TT, UK
Received 26 April 2002; Accepted 11 July 2002
The integration of signals received by a cell, and
their transduction to targets, is essential for all cellular responses. The cytoskeleton has been identi®ed
as a major target of signalling cascades in both animal and plant cells. Self-incompatibility (SI) in
Papaver rhoeas involves an allele-speci®c recognition between stigmatic S-proteins and pollen, resulting in the inhibition of incompatible pollen. This
highly speci®c response triggers a Ca2+-dependent
signalling cascade in incompatible pollen when a
stigmatic S-protein interacts with it. It has been demonstrated recently that SI induces dramatic alterations in the organization of the pollen actin
cytoskeleton. This implicates the actin cytoskeleton
as a key target for the SI-stimulated signals. The
cytological alterations to the actin cytoskeleton that
are triggered in response to SI are described here
and there seem to be several stages that are distinguishable temporally. Evidence was obtained that Factin depolymerization is also stimulated. The current
understanding that the actin cytoskeleton is a target
for the signals triggered by the SI response is discussed. It is suggested that these F-actin alterations
may be Ca2+-mediated and that this could be a mechanism whereby SI-induced tip growth inhibition is
achieved. The potential for actin-binding proteins to
act as key mediators of this response is discussed
and the mechanisms that may be responsible for
effecting these changes are described. In particular,
the parallels between sustained actin rearrangements
during SI and in apoptosis of animal cells are considered.
Key words: Actin binding proteins, actin cytoskeleton, Ca2+
signalling, pollen, self-incompatibility, signal transduction.
The cytoskeleton is a major target of signalling events in
plant cells. Both the microtubule and the actin ®lament
cytoskeleton in plants are known to rearrange when
numerous, diverse external stimuli are applied (Nick,
1999; Staiger, 2000). A major focus of plant cytoskeleton
research is the identi®cation of signals that regulate actin
dynamics (Staiger et al., 2000). In recent years there have
been several studies in plant systems that describe actin
rearrangements in response to speci®c, de®ned physiological stimuli. These include responses of stomatal guard
cells to abscisic acid or light; root hairs responding to Nod
factors from Rhizobium bacteria; and pollen tubes responding to self-incompatibility proteins (Eun and Lee, 1997;
CaÂrdenas et al., 1998; Miller et al., 1999; Geitmann et al.,
Although many of the components responsible for
regulating actin dynamics are well-de®ned in animal
cells and yeast, considerably less is known for plants.
Several actin-binding proteins (ABPs) have been identi®ed
in plants, but their role in vivo is not yet well-de®ned.
However, only a few ABPs which could regulate ®lament
dynamics in pollen tubes have been identi®ed and
characterized, namely, pro®lin, ADF/co®lin and 135ABP (Yokota and Shimmen, 2000; Hepler et al., 2001;
McCurdy et al., 2001). Furthermore, although the ABPs
have been implicated in mediating certain cytological
changes, their involvement in speci®c physiologically
relevant situations has not been demonstrated. No study on
To whom correspondence should be addressed. Fax: +44 (0)121 414 5925. E-mail: [email protected]
ã Society for Experimental Biology 2003
104 Staiger and Franklin-Tong
plants has attempted to characterize the effect that
signalling stimuli have on actin dynamics or, more
speci®cally, the state of actin polymerization.
Tip growing cells provide a good system whereby the
signals responsible for mediating changes to the actin
cytoskeleton may be studied. In plants, pollen tubes
represent a well-characterized cell where tip growth has
been studied in detail. Tip growth involves targeted vesicle
secretion and several signalling components have been
identi®ed as being associated with, or necessary for, pollen
tube growth. These include a tip-based cytosolic free Ca2+
([Ca2+]i) gradient, Ca2+-dependent protein kinase, Rho-like
small GTP-binding proteins, and cAMP (Estruch et al.,
1994; Pierson et al., 1996; Messerli and Robinson, 1997;
Zheng and Yang, 2000; Moutinho et al., 2001). Although it
is thought that the control of pollen tube growth involves a
complex interplay between the cytoskeleton and signalling
cascades, there is currently little direct evidence for this. It
is well established that an intact actin cytoskeleton is
essential for pollen germination and tip growth.
Furthermore, it has been demonstrated that both actin
organization and ®lamentous (F-)actin levels are involved/
important for pollen tube tip growth (Gibbon et al., 1999;
Geitmann and Emons, 2000; Fu et al., 2001; Vidali et al.,
2001). Despite this wealth of knowledge relating to these
individual components, direct causal links between these
components have not yet been established.
The self-incompatibility (SI) response in the ®eld
poppy, Papaver rhoeas L., provides a good example of a
speci®c and genetically controlled system that employs a
signal-mediated inhibition of tip growth. SI involves
highly speci®c `self' recognition of pollen to prevent
self-fertilization and consequent inbreeding. In this species, SI is determined by a single, multi-allelic S gene that
is gametophytically controlled (Lawrence et al., 1978). S
allele-speci®c self-incompatibility (S) proteins are secreted by the stigma and interact with pollen, resulting in
the rapid inhibition of growth of `self' or incompatible, but
not compatible pollen. It is proposed that the stigmatic Sproteins act as signal molecules that interact with a pollen
S-speci®c receptor that has yet to be identi®ed. The
identi®cation and cloning of the stigmatic S gene (Foote
et al., 1994; Walker et al., 1996; Kurup et al., 1998) and
the availability of an in vitro system for SI (Franklin-Tong
et al., 1988) has allowed the dissection of the signalling
cascade triggered by the SI response. Possibly the ®rst
events are S-speci®c increases in [Ca2+]i (Franklin-Tong
et al., 1993, 1997), which involves in¯ux (Franklin-Tong
et al., 2002; Rudd and Franklin-Tong, 2003). This is
thought to trigger the Ca2+-dependent phosphorylation of a
pollen protein, p26 (Rudd et al., 1996) and other signalling
events, including a putative programmed cell death
signalling cascade (Rudd and Franklin-Tong, 2003).
More recently, it has been demonstrated that the SI
response induces rapid alterations in actin cytoarchitecture
in incompatible pollen tubes (Geitmann et al., 2000;
Snowman et al., 2000b). Recent advances in the understanding of the actin cytoskeleton as a target for the SI
signalling cascade(s) are reviewed here, together with a
discussion of potential ABPs that may be involved in
alterations induced by SI.
Actin in normally growing pollen tubes
The F-actin cytoskeleton has been visualized in ®xed
pollen tubes using rhodamine-phalloidin and either ¯uorescence or confocal microscopy (see Geitmann et al.,
2000, for full details of methodology). m-maleimidobenzoyl N-hydroxysuccinimide (MBS) ester has been used,
which gives good quality actin con®gurations which are
not dissimilar in quality to those obtained using rapid
freeze ®xation (Doris and Steer, 1996; Miller et al., 1999).
Figure 1a shows an image of a typical poppy pollen tube
grown in vitro and subsequently ®xed and visualized using
this method. Typically, there are three main `zones' of Factin in the pollen tube: long, even arrays of longitudinal
actin ®lament bundles in the shank of the tube; a subapical
meshwork of F-actin in a `basket-like' con®guration and,
at the tip, an actin-depleted zone. This is very similar to the
actin con®guration which has been described for both ®xed
and living pollen tubes using both phalloidin and a GFPmouse::talin fusion protein (Miller et al., 1996; Kost et al.,
1998; Fu et al., 2001).
SI induces S-speci®c alterations to the F-actin
Several dramatic alterations to the F-actin cytoskeleton are
triggered by the induction of the SI response, which was
achieved by the addition of S-proteins to incompatible
pollen growing in vitro (Geitmann et al., 2000; Snowman
et al., 2000a, b). These are illustrated in Fig. 1. The ®rst
detectable changes were extremely rapid, being detected at
between 30 s and 2 min. Figure 1b shows that the
distinctive subapical `basket-like' con®guration had lost its
organization and actin bundles appeared to be displaced
into the apical region. This very rapidly gave the appearance of a big `blob' of F-actin in the apex of the pollen
tube, as illustrated in Fig. 1c. This tip-localized actin is a
well-recognized feature of tip growing cells that have
stopped elongating (Lancelle et al., 1997; de Ruijter et al.,
1999; Miller et al., 1999). The appearance of this marker
corresponds to the time-period when growth is arrested
(A Geitmann, AMC Emons, VE Franklin-Tong, unpublished data).
Very early in the SI response, pronounced phalloidin
labelling of F-actin adjacent to the plasma membrane was
detected (Fig. 1b, c). Optical sectioning, using confocal
microscopy, revealed this more clearly, as shown in
Fig. 1d. Whether this represents an increase in cortical
Self-incompatibility response in poppy 105
Fig. 1. Alterations in the actin cytoskeleton induced by the SI response. All images show rhodamine-phalloidin staining of F-actin using wide®eld ¯uorescence microscopy, except (d), which shows a confocal optical section. The scale bar in (i) is 10 mm. (a) Untreated pollen tube showing
normal F-actin con®guration. (b) 2 min SI showing actin at tip and evidence of F-actin at the plasma membrane. (c) 5 min SI showing a large
`blob' of F-actin at the tip and further evidence of cortical F-actin. (d) 2 min SI, single optical section obtained using confocal microscopy shows
both the F-actin at the plasma membrane and loss of F-actin in the cytoplasm. (e) 5 min SI shows loss of actin ®laments in the tip region, with a
speckled F-actin appearance. (f) 10 min SI shows further breakdown of F-actin bundles, and punctate actin structures in the cytosol and at the
cortex. (g) 60 min SI shows larger punctate actin structures. (h) Compatible SI reactions at 20 min and (i) 60 min show no alterations in the Factin organization. Since identical biologically active S-proteins were used in incompatible reactions, this demonstrates S-speci®city.
actin (i.e. movement of actin to this region or new
polymerization), or is simply remaining F-actin that is
more apparent due to the loss of signal in the lumen of the
pollen tube is not known. However, the impression is that
there is an increase in actin in this cortical region. This
needs further investigation. It is of interest that a predominance of cortical actin localization has also been observed
in animal cells undergoing apoptosis (Palladini et al.,
1996; GueÂnal et al., 1997). There is preliminary evidence
that programmed cell death (PCD) is likely to be induced
in the SI response (Jordan et al., 2000; Rudd and FranklinTong, 2003). However, this potential link is, at present,
speculative, and needs to be explored further. Concomitant
with this, the overall intensity of phalloidin staining was
reduced substantially within 2±5 min after challenge (see
Fig. 1d), indicating loss of F-actin.
By 5±10 min after SI challenge, the distinctive longitudinal F-actin bundles were no longer the prominent
feature in the pollen tube shank, and very few actin
®lament bundles were visible in the cytoplasm, particularly
in the region within ~50 mm of the tip. The phalloidin
labelling of F-actin gave a ®ne, speckled appearance, as
shown in Fig. 1e suggesting that severing or depolymerization might have occurred. These ®ne fragments of actin
were detected throughout the cytoplasm, but a distinct
distribution was visualized at the cortex. Further changes
were detected at later time points, and between 10±20 min
the F-actin appear to be organized into larger aggregates,
as shown in Fig. 1f. These larger, punctate foci of actin
were visualized throughout the pollen tube cytoplasm,
although they also displayed a prominent cortical localization. They appeared to increase in size over time and
became more discernible as distinct structures, as illustrated in Fig. 1g. Nucleation is not implied by the use of the
word foci, but is considered to be the best word to describe
these aggregates of actin. Studies indicate that these Factin structures persist for at least 3 h.
Cortical actin patches are a typical feature in yeast and,
in recent years, the ABPs and protein kinases involved in
the formation and dynamics of these structures have been
studied intensely (Pruyne and Bretscher, 2000; Pelham and
Chang, 2001). However, there are very few reports of actin
patch-like structures in other eukaryotic cells. It is not
known whether the punctate foci of F-actin are structurally
or functionally equivalent to these actin patches. With the
exception of the large focal adhesions that occur in pollen
tubes growing in vivo (Pierson et al., 1986; Lord, 1994),
actin aggregates have not been reported previously in
pollen tubes. Furthermore, the few reports of punctate actin
structures in higher plants have a contentious status as they
are not readily reproducible. For example, they were
visualized in bean root hairs undergoing nodulation factor
106 Staiger and Franklin-Tong
Fig. 2. Quanti®cation of F-actin changes induced by SI. The
histogram shows the percentages of pollen tubes displaying normal
distribution of F-actin ®laments (class I), membrane-localized F-actin
®laments (class II) or the membrane-localized fragmented F-actin
`punctate foci' (class III) following an incompatible challenge.
`Others' indicate morphologies that do not ®t these classes. [ã Kluwer
Academic Publishers. From: Snowman et al. (2000). Actin
rearrangements in pollen tubes are stimulated by the selfincompatibility (SI) response in Papaver rhoeas L. In: Staiger CJ,
Baluska F, Volkmann D, Barlow P, eds. Actin: a dynamic framework
for multiple plant cell functions, Dordrecht, The Netherlands: Kluwer
Academic Publishers.]
signalling (CaÂrdenas et al., 1998), but were not detected in
other studies of Nod factor stimulation (Miller et al.,
1999). In lower plants, although a cortical actin patch is
detected during the establishment of polarity in algae
(Braun and Wasteneys, 1998; Alessa and Kropf, 1999),
this appears very different to the numerous actin foci that
have been observed here. The nature of the actin patches
stimulated by the SI response, and identi®cation of the
components that interact with them, clearly need to be
investigated further in order to obtain some understanding
of their functional signi®cance.
The speci®city of the alterations in actin con®guration
was established by the use of several controls. The same
biologically active S-proteins were used to challenge
compatible pollen, thereby demonstrating the absolute Sspeci®city of the actin response because the only difference between the responses was the S genotype of the
pollen. Addition of germination medium and heatdenatured S-proteins that were biologically inactive to
incompatible pollen tubes acted as further controls. No
inhibition of growth and no alterations in the actin
cytoskeleton were observed in any of these situations
(Geitmann et al., 2000; Snowman et al., 2000b). This is
illustrated in Fig. 1h, i, which shows that 20 min and 60
min after a compatible challenge, the actin con®guration
appears normal. These controls demonstrate clearly that
the actin alterations observed are triggered speci®cally by
an incompatible SI response.
Quantitation of the alterations in F-actin organization
during the SI response at different time intervals has
helped to de®ne the timing of SI-induced events. This is
Fig. 3. SI-induced F-actin alterations also occur in pollen grains. All
images show rhodamine-phalloidin staining of F-actin. Images were
obtained using confocal microscopy and are full projections in (a), (c)
and (d), and single optical sections in (b), (d) and (f). Scale bar in (a)
is 10 mm. (a) A normal pollen grain at 20 min has germinated and
shows organized F-actin bundles. (b) An optical section through the
pollen grain in (a) shows discernible cytosolic F-actin. (c) A pollen
grain at 20 min post-SI has retained F-actin organization at the cortex,
but (d) an optical section shows that virtually all cytosolic F-actin has
been lost. (e, f) 60 min post-SI F-actin is organized into punctate foci
that is distributed throughout the cytosol and cortex.
shown in Fig. 2. Pollen tubes were scored according to the
organization of their actin cytoskeleton following SI
challenge. The initial movement of actin bundles at the
tip was not scored, since virtually all of the incompatible
pollen tubes showed this within 1±2 min. Three main
classes of F-actin organization were scored. Class I was
that found in normally growing pollen tubes (as described
earlier). Class II were pollen tubes with a predominance of
F-actin adjacent to the plasma membrane; while class III
were those with membrane-localized punctate foci of
actin. The histogram in Fig. 2 shows that after SI induction,
an increase in class II pollen tubes is detected concomitant
with a decrease in the number of class I pollen tubes.
Subsequently, the number of class II pollen tubes
decreases, and a proportional increase in pollen tubes of
class III is detected. After a compatible challenge 80% of
the pollen tubes counted were in the class I category (data
not shown; Geitmann et al., 2000). These data not only
Self-incompatibility response in poppy 107
show the initial speed of the response, but also establish
that the cortical predominance of F-actin occurs early and
prior to the formation of punctate foci of actin at the cell
Although the SI response can be reproduced in pollen
tubes that have been grown for several hours, this is not
what generally happens in vivo, where inhibition normally
occurs in pollen grains, either before or soon after
germination. This being the case, F-actin con®gurations
have also been examined semi-vivo, by incorporating Sproteins into an agarose-based germination medium, and
in vivo (Geitmann et al., 2000). The incompatible pollen
tubes displayed very similar alterations to the F-actin
con®guration as those challenged in vitro. It was also
examined whether the SI actin response occurs in
ungerminated pollen grains. These SI inductions in pollen
grains gave a similar, but delayed, actin response to those
seen in pollen tubes (Snowman et al., 2002). At 20 min,
normal pollen tubes had germinated, and had a wellorganized F-actin cytoarchitecture, as illustrated in Fig. 3a
and b. By contrast, at 20 min post-SI, much of the
cytoplasmic F-actin was lost, although F-actin staining
adjacent to the plasma membrane was retained, as
illustrated in Fig. 3c and d. Punctate foci of actin were
detectable at 60 min (as shown in Fig. 3e, f) and continued
to develop over several hours. This demonstrates that the
SI-stimulated actin changes can occur in ungerminated
pollen grains. SI in this species, although often occurring
before germination, almost always occurs after polarity has
been established, since deposition of callose is at a single
colpal aperture, rather than at all three. The 20 min delay in
the pollen grain actin response, which corresponds to the
time taken to germinate, suggests that polarity perhaps
needs to be established before the actin cytoskeleton can be
targeted. This suggests that it is polarized tip growth that is
targeted in this response. Further studies are required to
test this hypothesis. Nevertheless, these data provide
convincing evidence that the pollen actin cytoskeleton is
targeted by the SI-induced signalling cascades.
Cessation of growth is not responsible for the
actin alterations
In order to establish whether the SI-induced alterations in
actin con®guration observed were a primary effect stimulated by incompatible S-proteins or were merely a
secondary effect of the consequent arrest of growth, pollen
tubes were subjected to several treatments that caused
inhibition of pollen tube growth. Gd3+, caffeine and
EGTA, in appropriate concentrations can effectively arrest
pollen tube growth (Pierson et al., 1994, 1996). Although
Gd3+ and caffeine caused a displacement of the subapical
basket of actin, and EGTA caused only a very slight
change to the subapical mesh, the general organization of
the long arrays of actin bundles in the pollen tube shank
Fig. 4. Latrunculin B induces changes to the F-actin cytoskeleton in
pollen tubes. Effects of latrunculin B (10 min treatment) on F-actin in
pollen tubes of P. rhoeas. Pollen tubes were treated, ®xed and stained
with rhodamine-phalloidin. Images are single optical sections obtained
using confocal microscopy. Scale bar is 10 mm. (a, b) 100 nM LatB.
were intact and unchanged after these treatments
(Geitmann et al., 2000). Since this non-speci®c inhibition
of growth does not stimulate the gross changes in actin
con®guration observed in the SI response, these data
demonstrate that the SI-induced events are not merely
secondary consequences of inhibition of growth, but are
likely to be stimulated by the SI signalling cascade(s).
Could actin depolymerization be responsible for
some of the response?
These observations suggested that actin depolymerization
might be involved in mediating some of the cytoskeletal
changes stimulated in the SI response. In order to test this
possibility, the actin depolymerizing drugs latrunculin A
(LatA) and latrunculin B (LatB) were used, which have
been shown to bind to monomeric G-actin, thereby
reducing the pool of available actin subunits for polymerization (Spector et al., 1989). This results in actin ®lament
depolymerization; apparently LatB can completely deplete
cellular F-actin (Pendleton and Koffer, 2001). A few
minutes of treatment of pollen tubes with 50 nM LatA
resulted in the characteristic actin con®guration of the
pollen tube F-actin being replaced by a network of diffuse
actin arrays. However, phalloidin staining adjacent to the
plasma membrane remained strong. Treatment of pollen
tubes with 100 nM LatB for 10 min resulted in a signi®cant
loss of strongly-staining actin bundles. Short, weakly
stained actin ®lament bundles were detected throughout the
pollen tube, with a distinct signal adjacent to the plasma
membrane, as shown in Fig. 4a and b. This suggests that
there is a population of F-actin that is perhaps more
resistant to depolymerization most likely because of
interactions with unknown ABPs in this region.
Treatment with 1 mM LatB resulted in further apparent
fragmentation of the F-actin cytoskeleton, giving a speckled appearance.
108 Staiger and Franklin-Tong
These studies indicate that though the effect of LatB has
some similarity to the intermediate stages detected in
incompatible pollen tubes, the alterations to F-actin
induced by lantrunculin depolymerization are subtly
different to the changes in organization induced by SI.
This suggests that, although depolymerization is likely to
play a role in the SI response, other factors are, not
surprisingly, likely to be involved. Since the imaging data
for the SI-induced F-actin changes suggested that there
was likely to be a decrease in F-actin in incompatible
pollen tubes as, qualitatively, the phalloidin signal appears
to be reduced upon S-protein challenge, it was decided to
investigate this possibility further.
Quanti®cation of F-actin levels in pollen
In order to investigate the alterations to actin in incompatible pollen further, a quantitative approach has recently
been used. Although a number of studies have provided
microscopic analysis of alterations to the actin cytoskeleton in response to a variety of stimuli in plant cells, there
are few examples of quanti®cation and biochemical
analysis of the changes. Indeed, currently, there exists
only one published report of quanti®ed changes to the Factin concentration in a plant system, and this is in maize
pollen (Gibbon et al., 1999). This study used an adaptation
of a biochemical method based on the number of ¯uorescent-phalloidin binding sites which has been used to
measure F-actin content in activated neutrophils and yeast
(Howard and Oresajo, 1985b; Lillie and Brown, 1994).
This assay was used here to determine the concentration of
actin in polymeric form in pollen (Snowman et al., 2002).
The levels of F-actin in pollen throughout hydration,
germination and growth for several hours were surprisingly constant. The mean concentration of actin in
polymeric form was 15.4 mM (SE 62.7; n=22). A test
was then done to see if changes in F-actin concentration
could be detected by adding LatB, which will depolymerize F-actin (Gibbon et al., 1999). Pollen challenged for 10
min with 100 nM and 1 mM LatB revealed signi®cant
decreases in F-actin levels of ~60% and 45%, respectively
(Snowman et al., 2002). This demonstrated that the
phalloidin-binding assay was appropriate for measuring
rapid changes in F-actin levels in P. rhoeas pollen.
The SI response involves F-actin
Since the F-actin concentration in pollen could be effectively measured, it was examined whether an incompatible
SI challenge resulted in alterations to this. The previous
observations had suggested that depolymerization might
be stimulated, since there was a loss of phalloidin staining
in the cytoplasm and the F-actin bundles were lost, being
replaced by a speckled array of ®ne F-actin fragments.
However, this could potentially suggest fragmentation or
cleavage rather than depolymerization per se. It was
therefore important to establish what type of actin
alterations are being evoked.
Use of the phalloidin-binding assay has revealed a rapid,
large and sustained decrease in F-actin levels as a
consequence of an incompatible response. Even within 1
min, which was the earliest time-point taken, there was a
signi®cant reduction in F-actin compared to untreated
pollen. By 5 min after SI induction, F-actin levels were less
than 50% that of the controls. F-actin levels decreased
further during the 60 min time period over which these
studies were made (Snowman et al., 2002). These data
clearly demonstrate rapid and sustained F-actin depolymerization that was S-speci®c, since the relevant controls
of compatible pollen treated with the same S-proteins and
also incompatible pollen challenged with germination
medium or inactive S-proteins did not show changes in Factin levels (data not shown). This very large reduction in
F-actin was observed in both incompatible pollen grains
and pollen tubes. The overall F-actin reduction as a
consequence of SI challenge was 74% in tubes and 56% in
grains, compared with the GM controls at 60 min.
One interesting difference between the response in
pollen grains and pollen tubes was that the response in
pollen grains was delayed by ~20 min, which corresponds
to the time taken for polarity to be established. This might
suggest that the SI-induced actin depolymerization acts on
tip growth. This idea is substantiated by the studies with
maize pollen, which demonstrate that differences in
sensitivity to LatB are not due to ungerminated grains
having a more stable actin cytoskeleton than actively
growing pollen tubes (Gibbon et al., 1999).
Increases in cytosolic calcium result in changes
in F-actin
How the changes in the actin cytoskeleton described here
relate to the other SI induced signalling events already
identi®ed is an important question. It is well established
that Ca2+ acts as a second messenger in the SI response.
Increases in [Ca2+]i are probably the ®rst events in this
signalling cascade, and are detectable within a few seconds
of the SI-interaction (Franklin-Tong et al., 1997, 2002).
The ®rst changes to the F-actin cytoskeleton were detected
at the earliest time point possible for ®xed cells, 30 s.
Whether Ca2+ is involved, either directly or indirectly, in
these changes is an important question to answer.
Therefore an investigation as to whether F-actin may be
a target for a Ca2+-dependent signalling cascade has begun.
Treatments known to induce increases in [Ca2+]i in pollen
tubes were used. The Ca2+ ionophore, A23187 and
mastoparan (Franklin-Tong and Franklin, 1993; FranklinTong et al., 1996) were used to see whether arti®cial
increases in [Ca2+]i will stimulate changes to the status of
Self-incompatibility response in poppy 109
F-actin in pollen tubes. Both of these drug treatments
resulted in changes to the actin cytoskeleton that had some
broad similarities to those stimulated by SI. Features
included fragmented and bundled F-actin, small punctate
foci of F-actin, and later, larger punctate foci of F-actin
(data not shown; Snowman et al., 2002). considerable
decreases in F-actin, indicating depolymerization, stimulated by these drugs were also measured (data not shown;
Snowman et al., 2002). Since increases in [Ca2+] can
stimulate actin depolymerization, it suggests that the SIinduced Ca2+ increases target the actin cytoskeleton. Early
studies show that increasing [Ca2+]i arti®cially can disrupt
actin ®laments in pollen tubes (Kohno and Shimmen,
1987, 1988). However, although a gelsolin-like severing
activity was considered, this is rather speculative, as there
is no ®rm evidence for this suggestion.
Changes to the actin cytoskeleton cytoarchitecture stimulated by the SI response are described here, together with
preliminary studies that begin to quantify some of the
biochemical changes to the actin cytoskeleton. Several
alterations to the pollen F-actin cytoskeleton stimulated by
the SI response in P. rhoeas have been demonstrated.
Furthermore, a rapid and considerable F-actin depolymerization in incompatible pollen tubes has recently been
measured. Quantitative measurements of changes to Factin levels in a plant cell by a de®ned biological stimulus
have not previously been reported, although Gibbon et al.
(1999) provided the ®rst such measurements in pollen from
maize. Although there are several studies describing
changes to the plant actin cytoskeleton in response to
physiological stimuli, they are all descriptive (Eun and
Lee, 1997; Kobayashi et al., 1997; CaÂrdenas et al., 1998;
Miller et al., 1999). It is, therefore, not known if these
alterations involve reorganization or changes in the
polymerization status of actin.
Depolymerization of actin during SI
There are surprisingly few examples of actin depolymerization stimulated in response to a biologically-relevant
ligand, as extracellular stimuli generally appear to lead to
the polymerization of actin ®laments (Carlsson et al.,
1979; Howard and Oresajo, 1985a; Hall et al., 1988;
Greenberg et al., 1991; Downey et al., 1992; Becker and
Hart, 1999). The few examples of measurements of actin
depolymerization include the stress response of yeast
(Lillie and Brown, 1994; Yeh and Haarer, 1996), the
collapse of growth cones in response to chemo-repulsive
axonal guidance molecules (Aizawa et al., 2001), and
vasopressin stimulation of toad bladder epithelial cells
(Ding et al., 1991). However, it is striking that depolymerization is generally transient. For example, during the heat
shock response of yeast a 25±50% reduction in F-actin
within 30 s is observed, but levels return to normal within 3
min (Lillie and Brown, 1994). By contrast, SI-induced
pollen depolymerization continues for at least 60 min. This
suggests that the formation of actin punctae is due to actin
reorganization rather than re-polymerization. Since these
studies clearly demonstrate that alterations to the actin
cytoskeleton also continue well after the inhibition of tip
growth, this suggests that SI-induced signalling events also
occur after the inhibition of growth. Other SI studies also
indicate this (Rudd and Franklin-Tong, 2003). It is
proposed that some of these events may function to
make this arrest of growth irreversible.
Actin-binding proteins implicated in the pollen SI
The dramatic and rapid changes of the actin cytoskeleton
after S-protein challenge raise the question of how these
alterations are mediated. Actin-binding proteins (ABPs)
are thought to be responsible for modulating changes in
actin organization and dynamics. Depolymerization could
result from a loss of side-binding proteins, capping of
®lament ends, increased severing activity, or stimulation of
a sequestering protein. However, only a few ABPs have
been identi®ed and characterized in pollen tubes. These are
pro®lin, ADF/co®lin and 135-ABP (Hepler et al., 2001;
McCurdy et al., 2001) Although ADF/co®lin can cause
actin depolymerization and is regulated by Ca2+ indirectly,
increases in [Ca2+]i are expected to inhibit, rather than
activate F-actin depolymerization by ADF (Smertenko
et al., 1998; Allwood et al., 2001). 135-ABP is a villin-like
protein with Ca2+/calmodulin-regulated actin-bundling
activity (Yokota et al., 2000). However, no demonstration
of an actin capping, severing or depolymerizing activity
for 135-ABP has been made yet. Since a villin-like protein,
Drosophila QUAIL, lacks detectable severing and depolymerizing activity (Matova et al., 1999) it is unwise to
assume biochemical properties on the basis of sequence
homology. Furthermore, as there are no data on cellular
[135-ABP] in pollen or accurate measurements of its
af®nity for F-actin, its potential function in SI is currently
impossible to assess. Pollen pro®lin has recently been
shown to have increased actin-sequestering activity at
elevated, but physiological calcium concentrations (Kovar
et al., 2000). This ABP seems a likely contributor to actin
depolymerization during SI, but probably does not act
alone. Detailed analysis of cellular concentrations and
binding constants for these and other yet to be identi®ed
pollen ABPs will be necessary before the actin remodelling
that occurs during SI is fully understood.
A potential link between actin and PCD?
There is good evidence that apoptosis in some animal cell
types (e.g. HL-60 cells) can trigger F-actin depolymeriza-
110 Staiger and Franklin-Tong
tion; moreover, this status is maintained for several hours
(Levee et al., 1996; Korichneva and HaÈmmerling, 1999;
Rao et al., 1999). Thus, triggering of apoptosis is virtually
the only current system where long-term actin depolymerization has been measured. The large actin depolymerization that is sustained for at least 1 h in incompatible pollen
is highly reminiscent of events triggered by apoptosis. This
suggests that a programmed cell death (PCD) signalling
pathway might be triggered by SI in incompatible pollen.
Since there is preliminary evidence that PCD is activated
in pollen by SI (Rudd and Franklin-Tong, 2003), this is not
pure speculation, though further investigations to establish
a ®rm link need to be made.
Activated caspase-3, a key effector of apoptosis, has
been shown to have the capacity to cleave several cellular
proteins in animal cells. Both actin and ABPs are
implicated as targets for activated caspases. Proteolytic
cleavage of actin has been observed in some animal cell
types (Brown et al., 1997; GueÂnal et al., 1997; Korichneva
and HaÈmmerling, 1999; Paddenberg et al., 2001), although
this is not a universal phenomenon (Levee et al., 1996;
Bursch et al., 2000). Furthermore, there is evidence that
several ABPs are also targets for caspase activity
(Kothakota et al., 1997; Janmey, 1998; Umeda et al.,
2001), including gelsolin. The gelsolin cleavage product
can disrupt F-actin in the absence of Ca2+ and may
preferentially sever F-actin rather than bind monomeric
actin (Kothakota et al., 1997). Furthermore, this gelsolin
cleavage product can trigger apoptosis (Kothakota et al.,
Recent results from studies on animal cells indicate that
the regulation of actin polymerization may be an important
apoptotic morphological effector. There is convincing
evidence that F-actin depolymerization may act as a
effector of apoptosis in some cell lines (Janmey, 1998;
Korichneva and HaÈmmerling, 1999; Rao et al., 1999). It
has recently been demonstrated that depolymerization of
actin can activate caspases and cytochrome c release from
mitochondria, thereby stimulating apoptosis (Paul et al.,
2002). However, how depolymerization of the actin
cytoskeleton is achieved and maintained during apoptosis
is currently not known. Whether gelsolin is involved in
these cases is not yet known.
The existence of PCD in plant cells is now widely
accepted (Greenberg, 1997; del Pozo and Lam, 1998; Lam
et al., 2001). Although there are no clear caspase
orthologues in the Arabidopsis genome, a number of
studies indicate that caspase-like activities are triggered by
PCD in plant cells (D'Silva et al., 1998; del Pozo and Lam,
1998; Korthout et al., 2000; Richael et al., 2001). Since
there is evidence that PCD is stimulated in the SI response
(Jordan et al., 2000), with preliminary evidence for a
caspase-like activity stimulated in the SI response (Rudd
and Franklin-Tong, 2003), this might indicate a potential
reason why actin depolymerization is sustained. Thus, it
was postulated that a PCD signalling cascade is triggered
by the SI response and actin depolymerization may
potentially play an active role in the SI response by
stimulating caspase activity. This is, at present, the only
explanation (based on the available literature) for sustained
actin depolymerization observed during SI. Further studies
to explore the evidence for a PCD signalling cascade and
to provide ®rm links between this and the sustained
depolymerization of F-actin are required.
In summary, changes to the actin cytoskeleton during
the SI response have been described. There is evidence for
a rapid, massive and sustained depolymerization of Factin, and that this is induced via increased [Ca2+].
However, surprisingly, the most obvious (and at present,
the only) candidate for this Ca2+-dependent depolymerization, namely, pro®lin, is unlikely to be solely responsible
for the extent of depolymerization. The sustained nature of
the depolymerization is highly unusual and points to a
possible link with a PCD signalling cascade.
The work described here was carried out by Benjamin Snowman
funded by a BBSRC Biochemistry and Cell Biology Committee
studentship and by Anja Geitmann, in collaboration with Anne-Mie
Emons, funded by a Mare Curie Fellowship. This work was funded
by USDA-NRICGP support to CJS and BBSRC support to VEFT.
Thanks are due to BBSRC for a special travel grant that allowed
collaborative work between CJS and VEFT.
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