running head: corresponding author: address: telephone: email

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Running Head:

The pollen tube actin fringe and pectin deposition

Corresponding Author:

Peter K. Hepler

Address:

Biology Department, University of Massachusetts, Amherst, MA 01003

Telephone:

413-545-2083

Email:

[email protected]

Research Area:

Cell Biology

Plant Physiology Preview. Published on July 18, 2014, as DOI:10.1104/pp.114.242974

Copyright 2014 by the American Society of Plant Biologists

www.plantphysiol.orgon February 1, 2018 - Published by Downloaded from Copyright © 2014 American Society of Plant Biologists. All rights reserved.

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Title:

The apical actin fringe contributes to localized cell wall deposition and polarized growth in the lily

pollen tube

Authors:

Caleb M. Rounds,* Peter K. Hepler,* Lawrence J. Winship†

Institutions:

*Department of Biology, University of Massachusetts, Amherst, MA 01003

†School of Natural Science, Hampshire College, Amherst, MA 01002

One sentence summary:

Inhibition of lily pollen tube growth with three different agents, brefeldin A, latrunculin B and KCN

provides evidence that the apical actin fringe contributes to localized pectin deposition and polarized

cell growth.



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Footnote: This work was supported by the National Science Foundation (http://www.nsf.gov/)

(grant no. MCB-0847876 to P.K.H.). Corresponding author: Peter K. Hepler,

[email protected].



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Abstract

In Lilium formosanum pollen tubes pectin, a major component of the cell wall, is delivered through

regulated exocytosis. The targeted transport and secretion of the pectin containing vesicles may be

controlled by the cortical actin fringe at the pollen tube apex. Here we address the role of the actin

fringe using three different inhibitors of growth: brefeldin A (BFA), latrunculin B (LatB), and

potassium cyanide (KCN). BFA blocks membrane trafficking, and inhibits exocytosis in pollen

tubes; it also leads to the degradation of the actin fringe, and the formation of an aggregate of F-

actin at the base of the clear zone. LatB, which depolymerizes F-actin, markedly slows growth, but

allows focused pectin deposition to continue. Of note, the locus of deposition shifts frequently, and

correlates with changes in the direction of growth. Finally KCN, an electron transport chain

inhibitor, briefly stops growth while causing the actin fringe to completely disappear. Pectin

deposition continues, but lacks focus instead being delivered in a wide arc across the pollen tube tip.

These data support a model in which the actin fringe contributes to the focused secretion of pectin

to the apical cell wall, and thus to the polarized growth of the pollen tube.



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Introduction

Pollen tubes provide an excellent model for studying the molecular and physiological processes that

lead to polarized cell growth. Because all plant cell growth results from the regulated yielding of the

cell wall in response to uniform turgor pressure (Winship et al., 2010; Rojas et al., 2011) the cell wall

of the pollen tube must yield only at a particular spot: the cell apex, or tip. To accomplish the

extraordinary growth rates seen in many species, and to balance the thinning of the apical wall due

to rapid expansion, the pollen tube delivers prodigious amounts of wall material, largely

methoxylated pectins, to the tip in a coordinated manner. Recent studies suggest that the targeted

exocytosis increases the extensibility of the cell wall matrix at the tip, which then yields to the

existing turgor pressure, permitting the tip to extend or grow (McKenna et al., 2009; Hepler et al.,

2013). There are many factors that influence exocytosis in growing pollen tubes; in the current study

we investigate the role of the apical actin fringe.

For many years it has been known that an actin structure exists near the pollen tube tip, yet its exact

form has been a matter of some contention (Kost et al., 1998; Lovy-Wheeler et al., 2005; Wilsen et

al., 2006; Cheung et al., 2008; Vidali et al., 2009; Qu et al., 2013). The apical actin structure has been

variously described as a fringe, a basket, a collar or a mesh. Using rapid freeze fixation of lily pollen

tubes followed by staining with anti-actin antibodies, the structure appears as a dense fringe of

longitudinally oriented microfilaments, beginning 1-5 µm behind the apex and extending 5-10 µm

basally. The actin filaments are positioned in the cortical cytoplasm close to the plasma membrane

(Lovy-Wheeler et al., 2005). More recently we have used Lifeact-mEGFP, a probe that consistently

labels this palisade of longitudinally oriented microfilaments in living cells (Vidali et al., 2009) (Fig.

1A left column). For the purposes of this study, we will refer to this apical organization of actin as a

fringe.

Many lines of evidence demonstrate that actin is required for pollen tube growth. Latrunculin B

(LatB), which blocks actin polymerization, inhibits pollen tube growth and disrupts the cortical

fringe at concentrations as low as 2 nM. Higher concentrations are needed to block pollen grain

germination and cytoplasmic streaming (Gibbon et al., 1999; Vidali et al., 2001). Actin binding

proteins, including, ADF-cofilin, formin, profilin and villin, and signaling proteins, such as ROP

GTPases and their effectors (RICs), have also been shown to play critical roles in growth and

actin dynamics. (Fu et al., 2001; Vidali et al., 2001; Allwood et al., 2002; Chen et al., 2002;



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Cheung and Wu, 2004; McKenna et al., 2004; Gu et al., 2005; Ye et al., 2009; Cheung et al., 2010;

Staiger et al., 2010; Zhang et al., 2010a; van Gisbergen and Bezanilla, 2013; Qu et al., 2013).

Our understanding of the process of exocytosis and pollen tube elongation has been influenced by

ultrastructural images of pollen tube tips, which reveal an apical zone dense with vesicles (Cresti et

al., 1987; Heslop-Harrison, 1987; Lancelle et al., 1987; Steer and Steer, 1989; Lancelle and Hepler,

1992; Derksen et al., 1995). It has long been assumed that these represent exocytotic vesicles

destined to deliver new cell wall material. This model of polarized secretion has been challenged in

recent years in studies using FM dyes. Two groups have suggested that exocytosis occurs in a

circumpolar annular zone (Bove et al., 2008; Zonia and Munnik, 2008). However, other studies,

using fluorescent beads attached to the cell surface, indicate that the maximal rate of expansion, and

of necessity the greatest deposition of the cell wall material, occurs at the apex along the polar axis

of the tube (Dumais et al., 2006; Rojas et al., 2011). Similarly our experiments with propidium

iodide (PI) (1992; McKenna et al., 2009; Rounds et al., 2011a) and pectin methyl esterase fused to

GFP (McKenna et al., 2009) show that the wall is thickest at the very tip, and suggest that wall

materials are deposited at the polar axis consistent with the initial model of exocytosis (Lancelle and

Hepler, 1992). Experiments using tobacco pollen and a receptor-like-kinase fused to GFP, also

indicate that exocytosis occurs largely at the apical polar axis (Lee et al., 2008).

Many researchers argue that apical actin is critical for exocytosis (Lee et al., 2008; Cheung et al.,

2010; Qin and Yang, 2011; Yan and Yang, 2012). More specifically, recent work suggests that the

fringe participates in targeting vesicles, and thereby contributes to changes in growth direction.

(Kroeger et al., 2009; Bou Daher and Geitmann, 2011; Dong et al., 2012). In the current report,

using three different inhibitors, namely brefeldin A (BFA), latrunculin B (LatB) and potassium

cyanide (KCN), we test the hypothesis that polarized pectin deposition in pollen tubes requires the

actin fringe. Our data show that during normal growth pectin deposition is focused to the apex

along the polar axis of the tube. However, when growth is modulated different endpoints arise

depending on the inhibitor. With BFA, exocytosis completely stops, and the fringe disappears, with

the appearance of an actin aggregate at the base of the clear zone. LatB, as shown previously (Vidali

et al., 2009), incompletely degrades the actin fringe and leaves a rim of F-actin around the apical

dome. Here we show that in the presence of LatB pectin deposition continues with the focus of this

activity shifting in position frequently as the slowly elongating pollen tube changes direction. With



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KCN, the actin fringe degrades completely, but exocytosis continues, and becomes depolarized with

pectin deposits now occurring across a wide arc of the apical dome. This dome often swells as

deposition continues, only stopping once normal growth resumes. Taken together these results

support a role for the actin fringe in controlling the polarity of growth in the lily pollen tube.

Results

The fringe and the thickened tip are stable yet dynamic

Although the cell wall at the pollen tube tip oscillates in thickness (McKenna et al., 2009), it is

important to recognize, that even at its thinnest point the apical cell wall always remains thicker than

the cell wall along the shank. We monitor the changes in wall thickness using the fluorescent signal

from propidium iodide (PI), with which the pollen tubes have been stained. The efficacy of this

approach is based on the studies of McKenna et al. (2009) showing that the changes in PI

fluorescence are virtually identical to the changes in cell wall thickness measured by light microscopy

during oscillatory growth in the same lily pollen tubes. Subsequently the same group determined in

tobacco pollen tubes that the changes in PI fluorescence closely matched the fluorescent signal

derived from the secretion of pectin methyl esterase fused with GFP (McKenna et al., 2009).

Finally, in more recent work Rounds et al. (2011a) established that PI competes with Ca2+ in binding

to de-methoxylated pectin. We conclude from these studies that PI binds pectin and that it

faithfully indicates the appearance of new cell wall material. PI fluorescence thus serves as a

sensitive marker for pectin deposition in pollen tubes (Fig. 1A right column, Video S2) (Rounds et

al., 2011a).

As a first step in testing our hypothesis that the actin fringe regulates pectin deposition, we asked

whether there was a corresponding change in the fringe that reflected cell wall dynamics during

growth. To image actin in lily pollen tubes expressing Lifeact-mEGFP (Vidali et al., 2009), we used

spinning disc confocal microscopy. The improved time resolution of the spinning disc confocal

microscope relative to the conventional laser scanning confocal microscope permits us to obtain an

image of both the longitudinal filaments along the shank (Fig. 1 A left column, Video S1) and the

apical actin fringe at close time intervals. A kymographic analysis of the pollen tube shown in Fig.



8

1B documents the constancy of the fringe over time. In the kymograph a subtle line runs

immediately behind the tip (Fig. 1B arrow). Its slope closely matches that of the tip itself, indicating

that the fringe maintains a stable position relative to the tip despite the constant growth.

Actin filaments have been shown to be dynamic in plant cells (Blanchoin et al., 2010). To examine

whether the actin fringe is composed of stable filaments, we overlaid the first three images of the

series shown in Figure 1A. Each image was given a different color, and then the images were

superimposed on one another. Areas with coincident actin appear white, whereas areas with no

coincident actin appear in one of the individual colors. Because very few white domains are seen,

we conclude that although the structure of the actin fringe as a whole is stable, the individual

microfilaments are constantly changing.

BFA blocks exocytosis and destroys the actin fringe

The fungal macrocyclic lactone brefeldin A (BFA) blocks the activity of an Arf GEF, ultimately

resulting in inhibition of membrane trafficking (Nebenführ et al., 2002). Previous studies on pollen

tubes have established that the drug halts pollen tube growth and dramatically alters vesicle

trafficking. Recent data in particular indicate that both endo- and exocytosis are affected (Rutten and

Knuiman, 1993; Parton et al., 2003; Hörmanseder et al., 2005; Wang et al., 2005; Zhang et al.,

2010b). None of these studies actually examined deposition, instead focusing on internal membrane

markers and membrane associated proteins. BFA also alters the actin cytoskeleton, specifically

disrupting the apical actin structure in Arabidopsis pollen tubes (Zhang et al., 2010b). Earlier work

on lily pollen tubes showed that the addition of BFA led to the appearance of a cytoplasmic

aggregate behind the tip, called the BFA Induced Aggregate, or BIA (Parton et al. 2001, 2003). It

was further shown that the formation of the BIA depended on an intact actin cytoskeleton, because

anti actin drugs, including cytochalasin and jasplakinolide, either blocked or dispersed the formation

of the BIA. Zhang et al. (2010b) describe a dissipation of apical actin, whereas Parton et al. (2003)

documented the appearance of an actin dependent structure. These two descriptions are in some

ways contradictory. We wished to further investigate the effects of BFA directly on the fringe in lily

pollen tubes.

We asked what happens to the structure and distribution of apical actin and to changes in cell wall

pectin deposition in pollen tubes treated with BFA. As the drug takes effect growth slows and PI



9

fluorescence becomes significantly less intense and more uniform over the apical dome. Although

BFA takes several minutes (e.g., 20 min) to impact cell growth, once the pollen tube had stopped,

marked alterations are clear, which are evident as a substantial decline in the PI intensity at the cell

apex (Fig. 2A compare DIC and fluorescence images before and after BFA induced growth

inhibition, Video S3). Note especially that the intensity along the sides of the pollen tube does not

change significantly when compared to the apex (compare the two traces in Fig. 2B). We interpret

the concurrent changes in PI fluorescence and growth rate to mean that expansion of the apical cell

wall continues for a short time, but the absence of vesicles due to the action of BFA makes

exocytosis of new pectin impossible. The cell wall at the tip becomes more resistant to expansion,

possibly due to the action of PME on existing wall pectins, which leads to greater calcium

crosslinking and ultimately the cessation of growth. These data clearly show that BFA blocks cell

wall deposition, but they also further support the idea that exocytosis is confined largely to the

pollen tube tip.

BFA also exhibits a profound effect on actin organization. When we treat cells expressing Lifeact-

mEGFP with BFA, the apical fringe collapses and actin microfilaments appear as a star-like

aggregation of GFP immediately behind the tip (Fig. 2C, Video S4 and Video S5), in approximately

the same position as the BIA, described by Parton et al. (Parton et al., 2003). While the apical actin

fringe degrades, at least some of the actin cables in the shank of the tube remain and these drive the

streaming that is evident.

LatB leads to focused points of pectin deposition that frequently change position

LatB, which blocks actin polymerization by sequestering G-actin, has been shown to disrupt

localization of RabA4b, a Rab GTPase involved in exocytosis, in both root hairs and pollen tubes

(Preuss et al., 2004; Zhang et al., 2010b). Previous work has also shown that the actin fringe

dissipates in the presence of LatB (Cárdenas et al., 2005). In more recent studies on cells expressing

Lifeact-mEGFP, Vidali et al. (2009) have shown that 2 nM LatB disrupts the actin fringe per se,

however a thin rim of cortical fluorescence arises and spreads across the apical dome (see Fig. 7C,

D, and Supplemental movie S5 in Vidali et al., 2009). While some of the fluorescence in this rim

may be due to Lifeact associated with G-actin, a fraction is due to randomly oriented cortical F-actin

(Vidali et al., 2009). It seems likely that these cortical actin filaments drive the streaming that is

observed within the tube apex. Here we see all organelles, including the amyloplasts, which



10

heretofore had been excluded from the apical dome, moving through the apex obliterating the clear

zone.

When we treated pollen tubes with 2 nM LatB and monitored changes in pectin deposition as

measured by PI fluorescence the pollen tubes exhibited various defects in growth before the clear

zone collapsed and cells stopped growing, in agreement with previous results (Vidali et al., 2001).

Before completely stopping, cells generally began turning frequently and the PI signal tended to

move around the apical region rather than staying centered at the apex. A zone of intense PI

fluorescence often presaged the formation of a new growing tip. Frequently, pollen tubes developed

more than one deposition zone as evidenced by the presence of two focused points of PI

fluorescence in Fig 3, last panel (also see Fig. 7c in Vidali et al., 2001).

KCN reversibly inhibits growth

With a desire to achieve rapid inhibition together with relatively quick and full recovery of pollen

tube growth, we built upon earlier studies showing that inhibitors of the mitochondrial electron

transport chain, in particular potassium cyanide (KCN), were particularly effective (Rounds et al.,

2010; Obermeyer et al., 2013). Accordingly, lily pollen tubes were grown in a flow through chamber,

which allowed the quick addition and removal of KCN. Shortly after the addition of 200 µM KCN

pollen tubes stopped growing (~ 1 min). The KCN was washed out with fresh growth medium

immediately after growth stoppage and within a few minutes the cells began growing again. In Fig.

4A a representative pollen tube is shown before, during and after inhibition (see also Video S6 and

Video S7). The left hand panels show the DIC image whereas the right hand panels show the

fluorescent NAD(P)H signal, where the latter serves as a monitor of mitochondrial electron

transport chain activity (Cárdenas et al., 2006). Notice that the intensity of the signal after wash out is

roughly equivalent to that before inhibition. Fig. 4B provides a quantification of the data for the

same pollen tube. The blue line represents growth and the red line shows the NAD(P)H signal 20

µm behind the tip. Note that after recovery, the growth rate is similar to that before inhibition.

Also, the NAD(P)H signal clearly oscillates after recovery. Taken together these data support the use

of KCN as a quick acting, fully reversible inhibitor of pollen tube growth.

Pectin deposition continues during KCN-dependent growth inhibition



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To monitor apical pectin deposition, we used PI during cyanide-dependent growth inhibition. Fig.

5A shows that the PI signal first sharply drops during growth inhibition, but then increases

dramatically at the cell tip (see also video S8). To quantify the PI signal we measured the intensity

along a midline during the entire movie. This procedure describes the most intense signal at the very

tip of the pollen tube. These data are charted as the red line (Fig. 5B) against the growth rate (in

blue). As expected, before growth inhibition the PI signal oscillates as does the growth rate

(McKenna et al., 2009). Following application of cyanide, both the growth rate and the apical PI

signal dramatically decline. Remarkably, the PI signal almost immediately reverses and begins

trending upward again as indicated by the increase in fluorescence (Fig. 5A, third panel). Shortly

before growth resumes, the PI signal declines somewhat and then, along with the growth rate,

resumes oscillating.

As the PI signal has been shown to measure the amount of material in the cell wall by competing

with Ca2+ for binding to demethoxylated pectins (Rounds et al., 2011a), these data indicate that

pectin deposition continues when growth is inhibited by KCN. This is somewhat surprising as one

would expect exocytosis, and perhaps the maintenance of the actin fringe to be a significant energy

sink, and thus unable to occur in the absence of oxidative phosphorylation (Rounds et al., 2011b).

However, a sufficient supply of ATP may derive from glycolysis and aerobic fermentation, which

increases when the mitochondrial electron transport chain is blocked (Rounds et al., 2010;

Obermeyer et al., 2013). Interestingly despite these noticeable changes in cell wall deposition, the

DIC image does not show a dramatic change in the intracellular morphology; although the cell

swells somewhat, the clear zone does not disappear, and the general cell structure of a normal

uninhibited tube remains (Fig. 4A left column).

The actin fringe dissipates during KCN inhibition

To determine the effect of metabolic inhibition on the actin fringe we subjected pollen tubes

expressing Lifeact-mEGFP to inhibitory levels of KCN as described above, and then monitored the

GFP signal using a spinning disc confocal. The cortical actin fringe is apparent at the beginning of

the experiment (Fig. 6A top, Video S9), but during growth inhibition it disappears, although the

longitudinal actin filaments remain (Fig. 6A middle, Video S9). As growth restarts, the actin fringe

reforms (Fig. 6A bottom, Video S9) and is stable once again as growth recovers. To better visualize

this phenomenon, data from the same experiment shown in Fig. 6A are displayed as a kymograph



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(Fig. 6B). The slope of the line going from the left represents the growth. Under normal conditions,

the fringe can be visualized as a stripe of fluorescence immediately behind the tip. However, when

the pollen tube growth stops, and the line representing the apex becomes nearly vertical, the stripe

indicating the fringe disappears. It is also noteworthy that actin organization overall at the tip is

markedly reduced (Fig. 6B asterisk). Taken together these observations show that the fringe is

completely absent during growth inhibition by KCN; however exocytosis continues (see Fig. 5).

Pectin deposition loses polarity in the presence of KCN

Although pectin deposition continues despite the absence of the actin fringe (Fig. 5), it seems

possible that its directional specificity might be altered. That is, vesicle targeting might not be

focused, or might lose polarity entirely. To test this we analyzed the deposition of pectin under three

conditions: with BFA, with KCN, and without an inhibitor. To quantify deposition, the PI signal

from each pixel along the edge of the pollen tube was measured using a custom graphic analysis

program created with the R statistical analysis package (see Materials and Methods). These data were

then compiled in a matrix consisting of the edge values for each frame in the time series. The matrix

can then be visualized as a kymograph where the Y-axis represents all points along the edge in a

given frame and the X-axis shows time (Fig. 7A). The pollen tube’s apex is set to 0 µm along the Y-

axis. Therefore the signal along one edge of the pollen tube is a positive distance from the apex and

along the other is negative. A kymograph of the PI signal for a single control pollen tube is shown in

Fig. 7A. The pollen tube’s growth can be visualized by noting how the upper and lower edges trend

upward or downward as one’s eye moves from left to right along the X-axis. This reflects the fact

that the pollen tube gets longer and therefore the edge is longer. One can also see that the majority

of deposition occurs near the apex with the highest signal centering at 0 µm throughout the

kymograph. Furthermore, very little change in the signal can be observed 5-8 µm away from the

apex.

When the edge values for a single time point are plotted, with distance from apex on the x-axis and

signal intensity on the y-axis, a Gaussian distribution emerges (Fig. 7B). The highest signal is at the

apex, which drops quickly further from the apex. The point with the highest signal represents the

area of maximal pectin deposition (McKenna et al., 2009; Rounds et al., 2011a). We fit the PI data

to a Gaussian function using a non-linear least squares fitting algorithm within R. From the

Gaussian fit we derived a standard deviation, σ, thus providing a statistical description of the



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dispersion of the PI signal around the tube tip. If we model the pollen tube apex as a circle, 2 X σ

describes an arc of the highest signal along this circle (Fig. 7B and Fig. 7C as well as Materials and

Methods). This arc represents the majority of the PI signal at the apex; that is, it is the area of

greatest deposition. As pollen tubes vary in diameter quite substantially, we chose to quantify PI

dispersion by calculating the subtended angle of this arc (Θ c.f. Fig. 7C). We calculated Θ for each

frame of the pollen tube used for 6A and plotted it against the time (Fig. 7D). This allows us to

compare Θ to the amplitude of PI fluorescence at the apex (Fig. 7D). Notice in particular that the

amplitude of the PI signal oscillates as expected (McKenna et al., 2009; Rounds et al., 2011a), as

does Θ. In untreated pollen tubes Θ oscillates around 62° (+/- 3° S.E. n=10).

We also analyzed the change in Θ and therefore the arc of pectin deposition using BFA and KCN.

We reasoned that because BFA abolishes all exocytosis, Θ should go from the control value of ~62°

to “0°” or an undefined angle. If KCN causes a loss of control over deposition, then the angle

should increase dramatically. We conducted trials using 10 control pollen tubes, 11 treated with

KCN and 5 with BFA, and then analyzed the results as with the control described above.

The BFA treatment shows a dramatic reduction in deposition (Fig. 8A). When quantified along with

the signal at the apex, it is clear that deposition has dramatically slowed or stopped (Fig. 8A and B),

in agreement with our initial observations (Fig. 2). Our analysis of 5 pollen tubes confirms this

observation, and reveals that Θ, which begins at 45-50°, cannot be calculated after treatment with

BFA because the PI signal is low and uniform across the tip.

In the KCN treated pollen tube shown (Fig. 8C and D), the deposition initially falls off as with the

BFA treatment suggesting that at first deposition at the apex halts. Then quite suddenly deposition

begins again, but Θ is changed dramatically. After first stopping, Θ quickly rises to a plateau

significantly above the pre-inhibitor Θ (Fig. 7D). Our analysis showed that the Θ rose from 67°

(+/- 3° S.E. n=11) to 136° (+/- 18° n=11) (Fig. 9A) or roughly twice as high (p<0.001 with t-test).

These data suggest that in the presence of KCN deposition polarity has been altered dramatically,

perhaps even lost entirely.



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Discussion and Conclusions

The results of the present study indicate that the cortical actin fringe plays a major role in controlling

the polarized deposition of pectin, and therefore growth, in the lily pollen tube. In the first

experimental approach we used BFA, a well-established inhibitor of the exocytotic pathway. Here

we show that the deposition of pectin, as indicated by extracellular PI fluorescence, markedly

declines in the absence of exocytosis. Simultaneously we observe that the actin fringe dissipates, and

that a star-shaped aggregate of Lifeact-mEGFP appears in the clear zone behind the tube apex.

These observations suggest to us that the process of exocytosis and the presence of a normal cortical

actin fringe are connected.

The second experimental approach involves the use of LatB as a sensitive inhibitor of actin turnover

and dynamics. By binding to actin monomers LatB prevents formation of F-actin, thus leading to its

degradation, especially of those filaments that are rapidly turning over. The apical actin fringe indeed

turns over quickly and keeps strict pace with the rapidly growing pollen tube, as shown in Fig. 1.

Therefore it is not surprising that this array preferentially degrades in the presence of low

concentrations of LatB. While 2 nM LatB degrades actin structure, nevertheless a cortical array of

actin appears in the apex and as shown by Vidali et al., (2009) is expressed as a rim of actin around

the perimeter of the apical dome. A through focus series reveals that at least some of the Lifeact-

mEGFP fluorescence derives from randomly oriented microfilaments in the plane of the plasma

membrane (Vidali et al., 2009).

In lily pollen tubes treated with LatB, it is particularly interesting that pectin deposition continues,

but remains focused to definite hot spots. However, these focal locations change relatively quickly,

where they define new axes of pollen tube growth. It is possible that a similar process occurs in

tobacco pollen tubes expressing an RNAi or antisense transgene against formin FH5, given their

modified structure of apical actin and their meandering growth pattern (Cheung et al., 2010). For

lily pollen tubes, which change growth direction when treated with LatB, it seems plausible that local

exocytosis occurs in regions where there are breaks in the cortical rim of actin, and that these

momentarily become the focused deposition sites we observe.

The third experimental approach used KCN as a reversible inhibitor of the electron transport chain,

and pollen tube growth. Shortly after the addition of KCN, pollen tube growth stops abruptly.



15

Concomitantly, the apical fringe dissipates. At this juncture, pectin deposition as measured by PI

fluorescence at the apex drops dramatically, though only temporarily. Almost immediately

deposition begins again, but with a significant change. No longer is it focused on the apex; instead

deposition occurs over a broad arc at the pollen tube tip. We have measured the angle subtended by

this arc (Θ) and show that it doubles in the absence of the apical actin fringe (Fig. 8C and D). Given

the absence of the apical actin fringe under these experimental conditions, it is attractive to conclude

that this unique cytoskeletal structure contributes to the polarized wall deposition. Thus during

normal growth when the cortical fringe is present, by moving vesicles to the extreme apex it might

prevent their docking and fusion along the side of the pollen tube, insuring that these events will be

focused at the tube tip (Fig. 8D). However, when the fringe is degraded, e.g., during KCN treatment,

then exocytosis takes place more widely accounting for the loss of polarity in wall deposition. In

addition, the more widely spread infusion of methoxylated pectins will broadly weaken the wall

allowing the cell apex to swell (Fayant et al., 2010); although some swelling is usually observed, it is

not as dramatic as the de-polarization in pectin deposition (Fig. 8D).

We fully recognize that many factors, including physiological processes and signaling pathways

contribute to the formation and activity of the actin fringe, as well as to membrane trafficking.

Among the players, the GTPase ROP has often been characterized as a master regulator (Qin and

Yang, 2011). Through its effector proteins Ric3, Ric4 and ICR1, ROP is thought to control Ca2+

influx, actin dynamics and exocytosis (Fu et al., 2001; Gu et al., 2005; Hwang et al., 2005; Lavy et al.,

2007; Lee et al., 2008). In this scenario active ROP through Ric4 initiates actin polymerization and

accumulation of vesicles at the tip. Through Ric3, active ROP initiates the subsequent Ca2+ influx

and actin depolymerization. Finally through ICR1, ROP leads to the recruitment of the exocyst and

vesicle tethering (Lavy et al., 2007; Qin and Yang, 2011). These results are also consistent with the

idea that apical actin is critical for exocytosis.

While a ROP-dependent sequence may play an important regulatory role it is not the only pathway

that deserves attention. Given the tip focused Ca2+ gradient, which will have profound effects on

the apical actin possibly through specific actin binding proteins such as villin/gelsolin (Staiger et al.,

2010; Zhang et al., 2010a; Qu et al., 2013), it becomes apparent that any mechanism that modulates

intracellular Ca2+ could induce a cascade of events crucial to the targeted delivery of vesicles and

polarized growth. Candidate Ca2+ channels include those that are activated by mechanical forces or

stretch (Dutta and Robinson, 2004), by changes in the membrane potential (Qu et al., 2007; Wang et



16

al., 2004; Wu et al., 2010; Wu et al., 2007; Shang et al., 2005), or by the presence of a ligand such as

reactive oxygen species (Potocký et al., 2007; Wu et al., 2010), cyclic nucleotides (Frietsch et al.,

2007) or amino acids (Michard et al., 2011)(For a review see Hepler et al., 2011). In brief there is a

rich array of factors and mechanisms that could modulate Ca2+, and by extension the structure and

organization of the cortical actin fringe.

Finally the results presented in this study show that during normal pollen tube growth, deposition of

PI stained pectin occurs primarily at the apex of the tube, with a dispersion of ~60° centered at the

polar axis. These results are in close agreement with those obtained through the study of the growth-

dependent displacement of fluorescent particles on the pollen tube surface (Rojas et al., 2011).

These results may also include sufficient breadth to render them in agreement with studies using the

membrane marking FM-dyes (Zonia and Munnik, 2008; Bove et al., 2008). These authors have

argued that exocytosis occurs in an annulus around the polar axis. However, given the rapid

increase in area as one moves away from the polar axis, there will unavoidably be a great need for

additional exocytosis from that seen strictly at the polar axis.

The results from this study provide support for the hypothesis that the cortical actin fringe plays a

pivotal role in controlling the polarized growth of the pollen tube (Fig. 8B). During normal growth

in the presence of the actin fringe deposition is limited to the apical dome. However, when growth

is blocked with BFA or KCN the fringe degrades. BFA, because it inhibits membrane trafficking,

prevents any further exocytosis. By contrast KCN only stops pectin deposition momentarily. Even

in the continued presence of the metabolic inhibitor, deposition resumes, but notably now occurs

over the entire apical dome of the pollen tube. However, when the inhibitor is removed and normal

growth resumes, the actin fringe re-emerges, providing evidence for the close coupling between the

actin fringe and polarized pectin deposition. Finally, with LatB, although the localized actin fringe

degrades, a rim of cortical actin appears, which is spread across the dome of the pollen tube (Vidali

et al., 2009). Under these conditions some pectin deposition continues, which is confined to small

regions. When taken together the results give support to the idea that cortical actin participates in

moving vesicles to the preferred sites of fusion at the extreme tube apex.



17

Materials and Methods: Pollen Tube Growth Conditions

Pollen was grown as described previously (Rounds et al., 2010). Briefly, all pollen was from lily

(Lilium formosanum) stocks stored at –80°C and germinated for 1 to 1.5 h on a rotator at room

temperature in a standard growth medium (LPGM): 7% (w/v) Sucrose, 1.6 mM H3BO3, 0.1 mM

CaCl2, and 15 mM MES buffer adjusted to pH 5.7 with 10 N KOH (a final concentration of K of

approximately 6 mM); all reagents were from Fisher Scientific unless otherwise noted. For

microscopic observations pollen was prepared one of two ways. For widefield epifluorescence

microscopy, a pollen suspension was spread on custom-made well slides with a growth medium

solution containing a final concentration of 0.7% (w/v) low-melting agarose (Sigma-Aldrich). The

immobilized pollen was then covered with fresh growth medium for imaging. Cells were allowed to

recover for at least 0.5 h before imaging or further manipulations. For spinning disc confocal

microscopy cells were cultured in agarose as described above or on slides coated with high molecular

weight (150-300 kD) poly-l-lysine. These slides were first cleaned in a Harrick Plasma cleaner PDC-

001 for 30 seconds. They were then coated in ~50 µL of 1 mg/mL poly-l-lysine/H2O solution and

allowed to dry. The slides were thoroughly rinsed in H2O then allowed to air dry before use.

PI Staining of Pollen Tubes

For PI staining of the primary cell wall, the growth medium covering the immobilized and recovered

cells on microscope slides was replaced with LPGM supplemented with 20 μM PI (Sigma). Cells

were imaged more than 10 min after addition of the stain.

Growth Inhibition with KCN, BFA and LatB

Cells were grown as described above. For inhibition with KCN, a two-tube peristaltic pump (Bio-

Rad) was used to add 200 μM KCN in LPGM. One tube was used to remove the growth media

from the slide under analysis, while the other added media with inhibitor. The rate was set to ~ 0.5

mL min−1 for the duration of the experiment. For experiments with either BFA, or LatB, cells were



18

immobilized in agarose or on poly-l-lysine then a 200 μL of LPGM was added. Cells were allowed to

recover, then 200 μL of LPGM supplemented with either 30 μM BFA or 4 nM LatB was added to

yield a final concentration of 15 μM BFA or 2 nM LatB.

DIC and Epifluorescence Microscopy and Imaging

PI, NADH, and DIC images were acquired using a CCD camera (Quantix Cool Snap HQ; Roper

Scientific) attached to a Nikon TE300 inverted microscope (Nikon Instruments) with a 40x/1.3

numerical aperture oil-immersion objective lens. All the equipment was operated with

MetaMorph/MetaFluor (Molecular Devices) software. A filter wheel system (Λ10-2; Sutter

Instruments), mounted immediately before the CCD camera, controlled the position of a polarizing

filter for DIC or an emission filter for fluorescence imaging. Fluorescence excitation light was

provided by a 175-W ozone-free xenon lamp in a DG-4 switching system (Sutter Instruments).

Transmitted light was provided by a low-voltage halogen lamp. We used the following filter setup

for PI imaging: emission, 495 nm; a 565 nm dichroic long pass; and excitation 509 nm long pass (all

filters from Chroma). Exposure times varied but were generally approximately 10 ms for DIC and

approximately 800 ms for PI unless otherwise stated. We used the following filter setup for

NAD(P)H imaging: 360 nm (10 nm band-pass) as excitation filter, 380 nm dichroic, and 400 nm

long-pass emission filter (all filters were from Chroma). We employed an exposure time of 750 ms

and binned the images using ImageJ before analysis.

Spinning-disc confocal microscopy and imaging

For imaging of Lifeact-mEGFP, slides were prepared as described above and then were mounted on

an inverted microscope (model Ti-E; Nikon) equipped with a spinning disk head (model CSU-X1;

Yokogawa Corporation of America) and a 512 × 512 electron multiplying CCD camera (iXON;

Andor Technology). Images were collected with a 1.4 NA 60x oil immersion objective (Nikon) at

room temperature. 50% laser power was used with the 488 laser. The exposure time varied between

50 and 100 ms. Image acquisition process was controlled by MetaMorph software (Molecular

Devices). To create maximal projections, ImageJ (Abramoff et al., 2004) was used. Lifeact-mEGFP

kymographs were prepared using the multikymograph plugin for Image J with a 5 or 11 pixel-wide

scan along a hand selected midline.



19

Growth Rate and NAD(P)H Fluorescence Measurements

Growth rate was measured using the tip-tracking feature of the MetaMorph software package

(Molecular Devices). The average NAD(P)H fluorescence was measured in a 10 μm2 box centered 5

μm from the pollen tube tip (Cárdenas et al., 2006) using a custom R script (Ihaka and Gentleman,

1996; Rounds et al., 2010).

Bombardment

Plasmid DNA for Lifeact-mEGFP was constructed as described (Vidali et al., 2009). Plasmid DNA

was prepared using alkaline lysis followed by precipitation with PEG and extraction with phenol-

chloroform. DNA was coated onto 1 to 3 mg of 1.1 µm diameter tungsten particles (Bio-Rad

Laboratories) according to the manufacturer's instructions. The coated microprojectiles were

aliquoted onto two macrocarriers (Bio-Rad Laboratories). Pollen was allowed to hydrate in 1 mL of

the appropriate growth media (see below) before being placed on a 25 mm MF-Millipore membrane

(Millipore), which in turn was set on Whatmann paper moistened with pollen growth media. The

macrocarrier assembly was positioned in the top slot of the PDS-1000/He biolistic system and the

sample assembly in the slot below (Bio-Rad). Pollen grains were bombarded twice (once with each

aliquot) using an 1100-psi rupture disc (Bio-Rad). After bombardment, pollen was transferred to a

microcentrifuge tube and incubated for 2 hours at room temperature with constant rotation. Cells

were then immobilized on a microscope slide in growth as described above.

Edge/midline detection and PI fluorescence along the edge

For each tiff image in a sequence, approximate cell edges were first estimated by segmentation and

Canny edge detection in ImageJ (Abramoff et al., 2004). The cell midline was drawn by eye using

smoothing splines then digitized. Approximated edges were digitized then used as input to an R

script that scanned the perimeter of each cell pixel by pixel, extending a sample line perpendicular to

the approximate edge trace (please see supplemental information). Bicubic interpolation along the

sample line provided pixel values that were used to determine the point of the maximum PI signal.

The X and Y locations of the maximum PI signal were used as first estimate of edge location. The

center of the tip was calculated as the intersection of the midline and the edge trace. Then distances

from the tip along the cell edge (S) were calculated for each frame in a sequence as the geometric



20

distances between pairs of (X,Y) points. X and Y values were parameterized as smooth splines vs. S

and new X and Y coordinates for the smoothed edge were interpolated and used to determine the

PI value at each point from the original images. The location of the edge and the value of the PI

fluorescence at each location were saved for future analysis.

Kymographs

Kymographs were plotted using the kymo command in R.

Dispersion

For each image in a series PI fluorescence vs. S (with S=0 as the cell tip) data were first smoothed, a

minimum and maximum value determined, then the data normalized to the maximum PI value. The

resulting data were fit to a Gaussian function using the nls function (non-linear least squares) in R:

��

� �

��

��

PInorm is the normalized PI fluorescence, S is the distance from the tip, µ is the location of the peak,

and σ is the standard deviation, e.g. the square root of the variance, a measure of the dispersion of

the function and of the PI signal.

To compare pollen tubes over time and treatment, we used σ to calculate the angle θ (degrees) as an

index of PI dispersion (Figure 4) as:

� ��

��360

Supplemental Materials

Supplemental Protocol 1: R script for generating kymographic data.



21

Supplemental Protocol 2: R script for analyzing kymographic data to determine Θ

Supplemental Video S1: Lily pollen tube expressing Lifeact-mEGFP

Supplemental Video S2: PI labeling of lily pollen tube.

Supplemental Video S3: PI labeled pollen tube treated with BFA.

Supplemental Video S4: Lily pollen tube expressing Lifeact-mEGFP before BFA inhibition.

Supplemental Video S5: Same pollen tube as supplemental video S4 after BFA inhibition.

Supplemental Video S6: NAD(P)H auto-fluorescence in lily pollen tube treated with KCN.

Supplemental Video S7: DIC image of lily pollen tube treated with KCN.

Supplemental Video S8: PI labeled lily pollen tube treated with KCN.

Supplemental Video S9: Pollen tube expressing Lifeact-mEGFP treated with KCN.

Acknowledgements: We thank Dr. M. Bezanilla and Dr. T.I. Baskin and members of their

respective labs for helpful discussions.



22

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27

Figure Legends

Figure 1: The actin fringe and the thickened pollen tube tip wall are stable, though dynamic

structures during pollen tube growth. A, The left hand column shows a pollen tube transformed

with Lifeact-mEGFP imaged with a spinning disc confocal. Maximal projections from every 15 s are

shown. The second column shows an epifluorescence image of a pollen tube stained with PI. Again

images captured every 15 s are shown. In both instances the scale bar represents 10 μm. B, The data

from the pollen tube in A expressing Lifeact-mEGFP have been subjected to kymograph analysis

using an 11-pixel strip along the image’s midline. C, The first three frames from the pollen tube in A

and B were assigned the colors red blue and green respectively then overlaid. Areas with white show

overlap of all three. The fringe is stable, but most of its constituent actin is not shared between

frames.

Figure 2: BFA reduces PI signal at the pollen tube tip and alters the actin fringe. A, 15 μM BFA was

added to a growing pollen tube stained with 20 μM PI. The top pair of images represents the state

of the pollen tube before the addition of BFA. The first image is DIC, while the second is PI

fluorescence. The second pair represents the same pollen tube after BFA inhibition of exocytosis

and growth. Note the dramatic reduction in PI fluorescence at the apex. B, Data drawn from the

same experiment are shown in A. The red trace represents the PI fluorescence at the cell apex. The

blue trace represents the PI signal 16 μm behind the apex along the cell’s edge. C, A similar

experiment carried out with a pollen tube expressing Lifeact-mEGFP. The image on the left

represents the cell before BFA inhibition and the right hand image after BFA inhibition. The fringe

is prominent on the left and is no longer present on the right; instead a star like formation is clear.

Figure 3: LatB leads to focused points of pectin deposition. Pollen tubes treated with 2 nM LatB

lose their polarity as growth markedly slows. Notably the deposition of pectin, as indicated by PI

fluorescence, no longer remains strictly at the polar axis, but wanders over the apical region. Often

deposition can be observed simultaneously at two locations (last two image pairs). Eventually the

pollen tube completely stops elongating. These effects shown above are typical, but overall there is

considerable variability.

Figure 4: KCN inhibition is fully reversible. Pollen tubes were grown on microscope slides then at

36 s 200 μM KCN was added via a peristaltic pump. At 120 s shortly after growth had stopped,



28

washout of KCN was begun. A, DIC and NAD(P)H signal for a pollen tube going through KCN

inhibition is shown. The first image is from the beginning of the experiment, while the second pair

shows the pollen tube during inhibition; note the high NAD(P)H signal. The final pair shows the

recovered pollen tube with a lower NAD(P)H signal. B, Data drawn from the same experiment are

graphically displayed. The blue trace represents growth rate and the red represents the NAD(P)H

signal from a 10 μm box behind the tip. Note that both signals oscillate both before and after, but

not during KCN inhibition.

Figure 5: PI continues to accumulate in the absence of apical extension. A, 200 μM KCN was

added to the growing pollen tube at 30 s. At 150 s the KCN washout was begun. The images show

PI fluorescence and the scale bar represents 10 μm. B, Data drawn from the experiment shown in A

are graphically displayed. The blue trace represents the growth rate of the pollen tube tip, whereas

the red line represents the PI fluorescence at the apex of the cell. Note that both the growth rate and

PI fluorescence oscillate before and after, though not during, growth inhibition.

Figure 6: The actin fringe dissipates during KCN inhibiton. KCN was added to a growing pollen

tube expressing Lifeact-mEGFP at 49 s and the wash out begun at 126 s. A, The top frame shows

the pollen tube at the beginning of the experiment before KCN addition. A prominent apical fringe

of actin can be seen. The middle frame shows the pollen tube during inhibition, when the cell is not

growing. Longitudinal filaments are still present, but the fringe is absent. The bottom frame shows

the pollen tube after growth recovery; note that the fringe has reappeared. B, A kymograph of a five

pixel wide band down the center of the pollen tube shows the fate of the actin fringe in response to

KCN. When the fringe is present it appears as a diagonal line immediately behind the tip. Arrows

denote where the fringe disappears along with growth cessation and then reappears with the

resumption of growth. The asterisk shows the area where the fringe is completely absent.

Figure 7: Deposition of pectin is restricted to a distinct zone at the pollen tube apex. A, The PI

signal along the edge of a growing pollen tube was collected at each time point in a series. The apex

is assigned 0 µm on the Y-axis. The PI intensity at each point moving away from the tip along the

pollen tube’s edge is then plotted on the Y-axis. The X-axis represents time. This kymograph shows

the changes in fluorescence intensity along the pollen tube’s edge over time. B, The PI signal at a

single time point (60 s) is plotted with the distance from the apex on the X-axis and the intensity on

the Y-axis. A Gaussian function was fit to these data and the σ was calculated. The dashed blue line



29

shows the apex and the dotted blue line shows the distance indicated by σ. C, This schematic shows

the relationship between σ, which defines an arc at the apex and Θ, the angle subtended by twice

this arc. D, Data drawn from the same pollen tube show changes in the amplitude of the apical PI

signal (blue) and Θ (red dashed).

Figure 8: KCN and BFA alter the deposition of pectin at the pollen tube apex. A and B, Results

from a single experiment in which BFA was added to a pollen tube. The kymograph and Θ graphs

were created as described in figure 7. Deposition halts just after 300 s for BFA and Θ becomes

undefined. C and D, Treating pollen tubes with KCN at first causes an apparent loss in deposition,

as seen in the BFA experiment, but then results in a continuous increase in deposition over a

broader section of the apex as shown in D.

Figure 9: KCN and BFA affect both cell wall deposition and actin structure. A, Displays the Θ in

pollen tubes before KCN inhibition (fringe present), during growth stoppage (fringe absent) and

under control conditions. Error bars represent standard error. T-test between tubes before and after

KCN treatment yields p< .001. There was no significant difference between pre-treatment and

control tubes. B, Grey arrows in control (center) or KCN treated pollen tubes show the direction of

vesicles leading to deposition of wall material. The deposition is represented by the gray to black

gradient shown at the apex in control and spreading backwards in the KCN treated pollen tube. In

the image representing the control pollen tube, the actin fringe is represented as black lines at the

base of the clear zone. This fringe is missing from KCN treated pollen tubes, whereas the actin is

reorganized into a star shaped aggregate in BFA treated tubes.



Figure 1: The actin fringe and the thickened pollen tube tip wall are stable, though dynamic structures during pollen tube growth. A, The left hand column shows a pollen tube transformed with Lifeact-mEGFP imaged with a spinning disc confocal. Maximal projections from every 15 s are shown. The second column shows an epifluorescence image of a pollen tube stained with PI. Again images were captured every 15 s are shown. In both instances the scale bar represents 10 μm. B, The data from the pollen tube in A expressing Lifeact-mEGFP have been subjected to kymo-graph analysis using an 11-pixel strip along the image’s midline. C, The first three frames from the pollen tube in A and B were assigned the colors red, blue, and green respec-tively then overlaid. Areas with white show overlap of all three. The fringe is stable, but most of its constituent actin is not shared between frames.

A

B

C



Figure 2: BFA reduces PI signal at the pollen tube tip and alters the actin fringe. A, 15 μM BFA was added to a growing pollen tube stained with 20 μM PI. The top pair of images represents the state of the pollen tube before the addition of BFA. The first image is DIC, while the second is PI fluorescence. The second pair represent the same pollen tube after BFA inhibition of exocytosis and growth. Note the dramatic reduction in PI fluorescence at the apex. B, Data drawn from the same experiment are shown in A. The red trace represents the PI fluorescence at the cell apex. The blue trace represents the PI signal 16 μm behind the apex along the cell’s edge. C, A similar experi-ment carried out with a pollen tube expressing Lifeact-mEGFP. The image on the left represents the cell before BFA inhibition and the right hand image after BFA inhibition. The fringe is promi-nent on the left and is no longer present on the right. Instead a star-like formation is clear.

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Figure 3. LatB leads to focused points of pectin deposition. Pollen tubes treated with 2 nM LatB lose their polarity as growth markedly slows. Notably the deposition of pectin, as indicated by PI fluorescence, no longer remains strictly at the polar axis, but wanders over the apical region. Often deposition can be observed simultaneously at two locations (last two image pairs). Eventually the pollen tube completely stops elongating. These effects shown here are typical, but overall there is considerable variability.



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Time (s)Figure 4: KCN inhibition is fully reversible. Pollen tubes were grown on microscope slides then at 36 s 200 μM KCN was added via a peristaltic pump. At 120 s shortly after growth had stopped, washout of KCN was begun. A, DIC and NAD(P)H signal for a pollen tube going through KCN inhibition is shown. The first image is from the beginning of the experiment, while the second pair shows the pollen tube during inhibition - note the high NAD(P)H signal. The final pair shows the recovered pollen tube with a lower NAD(P)H signal. B, Data drawn from the same experiment are graphically displayed. The blue trace represents growth rate and the red represents the NAD(P)H signal from a 10 μm box behind the tip. Note that both signals oscillate both before and after, but not during KCN inhibition.

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Time (s)Figure 5: PI continues to increase in the absence of apical extension. A, 200 μM KCN was added to the growing pollen tube at 30 s. At 150 s the KCN washout was begun. The images show PI fluores-cence and the scale bar represents 10 μm. B, Data drawn from the experiment shown in A are graphically displayed. The blue trace represents the growth rate of the pollen tube tip whereas the red line represents the PI fluorescence at the apex of the cell. Note that both the growth rate and PI fluorescence oscillate both before and after, though not during, growth inhibition.

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Figure 6: The actin fringe dissipates during KCN inhibiton. KCN was added to a growing pollen tube expressing Lifeact-mEGFP at 49 s and the wash out begun at 126 s. A, The top frame shows the pollen tube at the beginning of the experiment before KCN addition. A prominent apical fringe of actin can be seen. The middle frame shows the pollen tube during inhibition, when the cell is not growing. Longitudinal filaments are still present, but the fringe is absent. The bottom frame shows the pollen tube after growth recovery - note that the fringe has reappeared. B, A kymograph of a five pixel wide band down the center of the pollen tube shows the fate of the actin fringe in response to KCN. When the fringe is present it appears as a diagonal line immediately behind the tip. Arrows denote where the fringe disappears along with growth cessation and then reappears with the resumption of growth. The asterisk shows the time when the fringe is completely absent

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Figure 7: Deposition of pectin is restricted to a distinct zone at the pollen tube apex. A, The PI signal along the edge of a growing pollen tube was collected at each time point in a series. The apex is assigned 0 µm on the Y-axis. The PI intensity at each point moving away from the tip along the pollen tube’s edge is then plotted on the Y-axis. The X-axis represents time. This kymograph shows the changes in fluorescence intensity along the pollen tube’s edge over time. B, The PI signal at a single time point (60 s) is plotted with the distance from the apex on the X-axis and the intensity on the Y-axis. A Gaussian function was fit to these data and the σ was calculated. The dashed blue line shows the apex and the dotted blue line shows the distance indicated by σ. C, This schematic shows the relationship between σ, which defines an arc at the apex and Θ, the angle subtended by twice this arc. D, Data drawn from the same pollen tube shows changes in the amplitude of the apical PI signal (blue) and Θ (red dashed).

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Figure 8: KCN and BFA alter the deposition of pectin at the pollen tube apex. A and B, Results from a single experiment in which BFA was added to a pollen tube. The kymograph and Θ graphs were created as described in figure 7. Deposition halts just after 300 s for BFA and Θ becomes undefined. C and D, Treating pollen tubes with KCN at first causes an appar-ent loss in deposition as seen in the BFA experiment but then results in a continuous increase in deposition over a broader section of the apex as shown in D.

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Figure 9: KCN and BFA affects both cell wall deposition and actin structure. A, Displays the Θ in pollen tubes before KCN inhibition (fringe present), during growth stoppage (fringe absent) and under control conditions. Error bars represent standard error. T-test between tubes before and after KCN treatment yields p< .001. There was no significant difference between pre-treat-ment and control tubes. B, Grey arrows in control (center) or KCN treated pollen tubes show the direction of vesicles leading to deposition of wall material. The deposition is represented by the gray to black gradient shown at the apex in control and spreading backwards in the KCN treated pollen tube. In the image representing the control pollen tube, the actin fringe is repre-sented as black lines at the base of the clear zone. This fringe is missing from KCN treated pollen tubes. whereas the actin is reorganized into a star shaped aggregate in BFA treated tubes.

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running head: corresponding author: address: telephone: email

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