IntroductionThe angiosperm pollen tube, or male gametophyte, is a highlypolarised, rapidly tip-growing plant cell, specialised to delivergenetic material from the site of pollination on the flowerstigma to the site of fertilisation at the ovule (Heslop-Harrison,1987). Combined with their relative ease of experimentalmanipulation, these features make the pollen tube a valuablemodel system for the study of processes associated withpolarity and growth at the single cell level (Feijó et al., 2001;Hepler et al., 2001).

One of the most intriguing aspects of pollen tube growth,which is shared with tip-growing fungal hyphae and plant roothairs, is the phenomenon of periodicity or oscillation in growthrate (Lopez-Francó et al., 1994; Pierson et al., 1996; Wymer etal., 1997). Animal neurons, which also extend by tip growth,do not appear to exhibit such regular periodicity, although theydo undergo growth rate fluctuations triggered by environmentalcues and dependent upon apical fluctuations in intracellularcalcium (Gomez and Spitzer, 1999). Pollen tube growth rateand orientation are also closely linked to intracellular calciumsignals at the tip – most notably the oscillating tip-focused

gradient that exhibits the same periodicity as growth rate(Malhó et al., 2000; Hepler et al., 2001). Several other growthassociated periodic phenomenon in the pollen tube have beennoted including other ion gradients and fluxes at the apex(Messerli and Robinson, 1998; Feijó et al., 2001; Zonia et al.,2002), cell wall banding patterns (Pierson et al., 1995; Li et al.,1996) and dynamics of the apical vesicle accumulation (Partonet al., 2001). Despite recent interest, the significance of growthrate periodicity, the underlying driving force, how it isintegrated with the other mechanisms involved in tip growth aswell as how widespread this phenomenon is in different pollentube species are not well understood.

In our previous studies, using FM4-64 as a marker of theapical vesicle accumulation, we observed an apparentstructural organisation and clear periodicity in the movementsof labelled material comprising the apical vesicle accumulationthat was closely related to the periodicity of growth ratefluctuation (Parton et al., 2001). These findings suggested thepossibility of a regulated periodicity to membrane traffic andvesicle trafficking at the pollen tube tip and prompted us toquestion the relationship between the movements of the apical

2707

The growing pollen tube provides an excellent singlecell model system in which to study the mechanismsdetermining growth regulation, polarity and periodicbehaviour. Previously, using FM4-64, we identified periodicmovements within the apical vesicle accumulation thatwere related to the period of oscillatory growth. Thissuggested a more complex interdependence betweenmembrane traffic, apical extension and periodicity thanpreviously thought. To investigate this a comparison wasmade between normally growing and Brefeldin-A-treated,non-growing, tubes. Brefeldin-A treatment establishedan intriguing, stable yet dynamic system of membraneaggregations in the pollen tube tip that exhibited regularmovements of material with a 5-7 second period comparedwith the normal ~30 second periodicity observed ingrowing tubes. Heat treatment was found to reduce periodlength in both cases. After BFA treatment membrane wasdemonstrated to flow from the extreme pollen tube apexback through a distinct subapical Brefeldin-A-inducedmembrane accumulation. The effects of Brefeldin-A on thedistribution of ER- and Golgi-targeted fluorescent proteins

revealed that ER did not contribute directly to thesystem of membrane aggregations while only certaincompartments of the Golgi might be involved. Theinvolvement of membrane derived from the apicalvesicle accumulation was strongly implicated. Calciummeasurements revealed that Brefeldin-A abolished thetypical tip-focused calcium gradient associated with growthand there were no obvious periodic fluctuations in apicalcalcium associated with the continued periodic Brefeldin-A membrane aggregation associated movements. Ourexperiments reveal an underlying periodicity in the pollentube that is independent of secretion, apical extension andthe oscillating tip-focused calcium gradient normallyassociated with growth, but requires an active actincytoskeleton.

Movies available online

Key words: Oscillating growth, Pollen tube, Brefeldin A, Membranetrafficking

Summary

Pollen tubes exhibit regular periodic membranetrafficking events in the absence of apical extensionRichard M. Parton*, Sabine Fischer-Parton, Anthony J. Trewavas and Masaaki K. WatahikiInstitute of Cell and Molecular Biology, University of Edinburgh, Mayfield Road, Edinburgh EH9 3JU, UK*Author for correspondence (e-mail: richard.parton@ed.ac.uk)This article is dedicated to Allan Gillies (1947-2003), who died recently. His work in organisation and administration within the University of Edinburgh has over many yearscontributed much to the advancement of science through his maintenance of a productive research environment.

Accepted 10 March 2003Journal of Cell Science 116, 2707-2719 © 2003 The Company of Biologists Ltddoi:10.1242/jcs.00468

Research Article

2708

vesicle accumulation, apical extension and periodicity. Suchmovements of vesicles and endomembranes recorded by lightmicroscopy, which may also include vesicle budding andfusion events, we have termed here ‘membrane traffic’ distinctfrom ‘vesicle traffic’ which here specifically implies theinvolvement of vesicle budding and fusion.

Like other tip-growing cells such as fungal hyphae (Fischer-Parton et al., 2000); growth neurones (Andersen and Bi,2000); root hairs (Wymer et al., 1997) and rhizoids (Bartnikand Sievers, 1988; Parton et al., 2000), in pollen tubes thedistinct localised apical vesicle accumulation is associatedwith exocytic delivery of material to the extending apex (Steerand Steer, 1989; Lancelle and Hepler, 1992; Cheung et al.,2002). In the pollen as well as root hairs and fungal hyphaethere are also reports of endocytosis associated with tip growth(O’Driscoll et al., 1993; Derksen et al., 1995; Blackbournand Jackson, 1996; Wymer et al., 1997; Fischer-Parton et al.,2000).

In living pollen tubes of Lilium longiflorum we showed thatBrefeldin-A (BFA) reproducibly leads to the loss of the FM4-64 labelled apical vesicle accumulation which correlates witha rapid, BFA concentration dependent, decline in growth rate,inhibition of secretion and the subsequent appearance of an‘undefined structure’ which accumulated label [see Fig. 9(Parton et al., 2001)]. Previous studies have established thatBFA, a known inhibitor of vesicle production in plant andanimal cells (Ritzenthaler et al., 2002), blocks secretion of cellwall material in tobacco pollen tubes resulting in growth arrest(Rutten and Knuiman, 1993; Geitmann et al., 1996; Ueda etal., 1996).

In mammalian cells BFA has been shown to block COPIcoatomer protein mediated vesicle trafficking by interfering inthe complex between GTPase ADP ribosylation factor 1 (Arf1)and its guanine nucleotide exchange factor Sec7 domain bylocking it in its inactive form with GDP (Robineau et al., 2000;Rohn et al., 2000).

In the present study we initially characterised the dynamicnature and origins of our previously undefined BFA-inducedstructure and noted an apparent periodicity in the membranetrafficking associated with it that had not been reported before.These observations of what appeared to be periodic behaviourassociated with a non-growing tube prompted us to investigatethis phenomenon in relation to the normal expression ofperiodicity in growth rate, membrane trafficking at the apexand the oscillating tip-focused gradient in intracellular calciumassociated with growing tubes. Our findings lead us to proposethat the periodicity exhibited in the vesicle cloud movements(Parton et al., 2001) during normal growth may be theexpression of an underlying periodicity that contributes toperiodicity in growth rates in pollen tubes rather than beingdependent upon it.

Materials and MethodsChemicals and materialsChemicals for culture media were obtained from BDH Chemicals(Poole, Dorset, UK) or Fisher Scientific (Loughborough, UK). Dyesand inhibitors were obtained from Molecular Probes Europe (Leiden,Netherlands) or Calbiochem-Novabiochem LTD (Nottingham, UK)and made up as recommended by the supplier. Solvent concentrationwas kept below 1% (v/v).

Plant materialPollen of Nicotiana tabacum, N. plumbaginifoliaand Arabidopsisthaliana was collected fresh from plants cultured under glass at theUniversity of Edinburgh. L. longiflorum pollen was obtained fromfresh cut flowers obtained locally.

Media used for pollen culture were: as described (Read et al., 1993)for the Nicotianaspecies; modified from that described (Feijó et al.,1999), by the addition of 2 mM CaCl2, for L. longiflorum, andmodified to 0.01% Boric acid, 0.07% CaCl2, 3% PEG 6000, 10%sucrose (w/v) from that described (Hodgkin, 1983) for Arabidopsisthaliana. Growth media and conditions were selected for rapid growthrate and even, straight tube morphology. Batches of cultured tubes orindividuals showing slow or irregular growth were not included foranalysis.

Where culture medium was used at increased concentrationsrelative to normal strength (100%); this was diluted from doublestrength medium (200%) and quoted as percentage strength. Pollenwas imbibed for 10 to 30 minutes in the appropriate liquid mediumbefore transfer to thin layer growth chambers, as described (Parton etal., 2001) and incubated in a humid environment for 3 to 12 hours (at20-25°C).

Targeting and expression of fluorescent proteinsTargeting of GFP to the ER was achieved by expression of mGFP5-ER [based upon the C-terminal ‘HDEL’ ER retrieval signal and N-terminal ER transit peptide (Pelham, 1989; Haseloff et al., 1997;Ridge et al., 1999)] in pollen expression vector pART16. Golgiapparatus targeting was achieved by the expression of GT-EYFP[EYFP-Golgi: based upon the N-terminal 81 amino acid targetingsequence of the Golgi resident, type II membrane protein, human β-1,4-galactosyltransferase (Roth and Berger, 1982; Palacpac et al.,1999), Clontech, Basingstoke, UK] in pART16. The Golgi apparatuswas also targeted with the GONST1-YFP sequence [based upon theArabidopsis Golgi localised GDP-mannose transporter, accessionnumber AJ314836 (Baldwin et al., 2001)] in pART16. Cytoplasmiclocalised GFP expression was achieved with tandem linked solubleGFPs (smGFP) (Davis and Vierstra, 1998) in pART16.

The pART16 pollen expression vector was constructed onpT7Blue-3 cloning vector (Novagen, Darmstadt, Germany). Thepromoter region and 5′UTR were amplified by PCR using SC5-122SN(TAGAGCTCGCGGCCGCAATCGCATCTCATATGTC) and ASC5-1920XH (GGCCTCGAGTGCCCGCTAGCTTGTAGTTGAATC).The OCS terminator and multiple cloning site of pART7 (Gleave,1992) were cut out with NotI and XhoI restriction enzymes. Fragmentsof ZmC5 promoter (Wakeley et al., 1998), 5′ UTR, multiple cloningsite, and OCS terminator were cloned into KpnI, NotI site of pT7Blue-3. The resulting construct was named pART16 and contained the samemultiple cloning site as pART7 but also included kanamycinresistance.

Transient expression of fluorescent protein constructs in pollen wasachieved by biolistic bombardment (with DNA-coated 1 or 1.6 µmgold particles) of wetted pollen as described (Kost et al., 1998).

Subcellular localisation of fluorescent protein tagged constructswas confirmed by transient expression in onion epidermis under the35S promotor (biolistic bombardment), examined by confocalmicroscopy.

Microscopy and imagingDifferential interference contrast (DIC) video imaging at up to 1image/400 millisecond was performed using a Nikon TMD invertedmicroscope, ×40 DIC 0.85 NA objective and Orca-ER cooled CCDcamera (Hamamatsu Photonics, Japan) driven by the ImprovisionOpenlab software 3.04. Images were processed and quantitativelyanalysed in ImageJ V1.27z (National Institutes of Health, USA,http://rsb.info.nih.gov.ij).

Journal of Cell Science 116 (13)

2709Pollen tube periodicity

Confocal microscopy was performed using a Leica TCS SPConfocal Microscope as described (Parton et al., 2001). ×63 Leicawater immersion plan apo (NA 1.2) and ×20 multi-immersion (NA0.75) objectives were used throughout. Images were imported intoImageJ for quantitative analysis or Adobe Premiere 5.1 for generationof video clips. Excitation and emission wavelengths forfluorochromes are given in Table 1. Loading of cells with stains wasachieved by application during the imbibition of pollen grains inliquid medium or by direct addition of dye solutions in 115% liquidmedium to growing tubes on thin gel layers. Inhibitors were applieddirectly to growing pollen tubes on thin gel layers in 70 µl of 115-120% liquid medium.

Fluorescence-recovery-after-photobleaching experiments wereperformed on the confocal imaging system using the 514 nm laserline and the bleach protocol of the TCSNT software.

Heat treatments were imposed using a heated stage against a roomtemperature of 20±1°C; culture medium temperature was constantlymonitored via a thermocouple (type T) temperature probe.

Simultaneous dual channel confocal ratio imaging of cytosolic[Ca2+] was carried out using a mixture of Texas Red (TR)-10,000 kDaDextran (volume marker) and Oregon Green 488 BAPTA (OG-488)-10,000 kDa Dextran (Calcium sensor). A 1:1 (~1 mM each) dye

mixture was injected into pollen tubes using a customised version ofthe pressure probe injection system (Oparka et al., 1991). Quantitativeanalysis and ratio image generation were done in ImageJ.

Results BFA treatment of pollen tubes gives rise to a distinctstructure within the cytoplasm of the tipAs reported previously (Parton et al., 2001), BFA treatmentresults in a particular re-organisation of the cytoplasm at thepollen tube apex, easily seen with FM4-64 staining (Fig. 1A-D). In brief, 15-60 minutes after BFA application (dependingupon concentration), the FM4-64 membrane-staining pattern atthe pollen tube tip (Fig. 1A; Movie 1, http://jcs.biologists.org/supplemental) was disrupted such that the extreme apicalstaining was more diffuse and less well localised, no longerexhibiting the characteristic ‘V’-shaped pattern within theapical clear zone. Even more obvious was the emergence of adistinct, relatively large (5-10 µm diameter in L. longiflorum),intensely stained subapical structure or aggregation ofmembrane material (confirmed by Nile Red membranelabelling, not shown) which we have termed the BFA-inducedaggregation or BIA (Fig. 1B; Movie 2). These events wereremarkably consistent within populations of cells and for thedifferent effective BFA concentrations used between 3.0and 36 µM (concentrations leading to growth arrest andBIA structures in at least 50% of intact tubes). Similarreorganisation was also observed in BFA-treated N.plumbaginifoliapollen tubes (Fig. 1C,D).

Reversibility of BFA effects has previously been reported forplant cell culture (Ritzenthaler et al., 2002); however, in pollentubes and fungal hyphae only partial reversal has been reported(Rutten and Knuiman, 1993; Cole et al., 2000). We found thatwhen L. longiflorumpollen tubes that had been treated for 2-

4 hours with BFA (10 µM) were subjectedto 3 hours of continuous perfusionwith fresh medium, the BFA-inducedrearrangements became less distinct butgrowth did not resume.

Polarity and streaming are altered butnot abolished by BFA treatmentIn order to characterise how the dynamicorganisation of the pollen tube tip wasaltered by BFA treatment, a picture of the

Fig. 1.BFA effects the organisation of livingpollen tube tips. Confocal images of FM4-64distribution in L. longiflorum(A,B) and N.plumbaginifolia(C,D) before and 30-60minutes after 3.6 µM BFA treatment.(E-F) Confocal images of mitofluor stainingbefore and after BFA treatment in L.longiflorum.(G-H) Schematic of cytoplasmicmovements in (G) growing and (H) BFA-treated pollen tube, based upon imagesequences. (I) Projected time series of an FM4-64-stained, BFA-treated tube showing tracks ofmovement. Bars, 15 µm (A,B,E,F) or 10 µm(C,D); corresponding bright field imagesshown at one-third size.

Table 1. Excitation and emission wavelengths offluorochromes

Fluorochrome Excitation* nm Emission† nm

FM4-64 514 or 488 600-700Mitofluor 514 or 488 >520Syto 16 514 550-600smGFP 488 515-550YFP 488 or 514 530-600Texas Red 568 590-700Oregon Green 488 488 492-544

*100 mW Argon ion laser.†Tunable emission filter of the Leica TCS SP confocal.

2710

apical zonation and streaming pathways of growingand BFA treated non-growing L. longiflorumpollentubes was built up.

The ‘reverse-fountain’ streaming path is wellknown for pollen tubes, different organelles can beobserved to follow slightly different trajectories andthere is an obvious ‘non streaming’ region within acone-shaped region of the apex corresponding to theapical vesicle accumulation (Heslop-Harrison, 1987;Parton et al., 2001). Mitofluor Green staining wasused as a convenient marker of the streaming pathfollowed by mitochondria (Fig. 1E; Movie 3). Thestructurally closely related dye Mitotracker Greenhas been used previously to label mitochondria inplant cells (Coelho et al., 2002) and the distributionof mitochondria in pollen tubes was independentlyconfirmed by staining mitochondrial DNA with Syto16 (not shown). The paths of lipid storage dropletsand amyloplasts could easily be followed by DICmicroscopy.

In BFA-treated tubes, although vigorous reverse-fountain cytoplasmic streaming continued it could beseen to have shifted further from the tube apex tobehind the BIA. Mitochondria, which in growingtubes follow a streaming pathway very close to thetip, along the flanks of the apical clear zone (Fig.1E,G; Movie 3), after BFA treatment showed adistribution restricted to no further than the site of theBIA (Fig. 1F,H; Movie 4). The situation was the samefor lipid storage droplets and amyloplasts (Fig. 1H).Yet cytoplasmic movements in front of the BIA werenot completely abolished. Within the region, whichwould in a growing tube (Fig. 1G; Movies 1, 3) beoccupied by the apical vesicle accumulation,movement of clumped material could be seenassociated with the BIA (Fig. 1I; Movies 2, 4, 5).These clumps of material (BIA-associated membraneaggregations) periodically ‘materialised’ andmigrated from the extreme apex towards the BIAwith which they appeared to merge.

Flow of material can be tracked through the BFA-induced membrane aggregationThe movement of BIA associated aggregations was tracked, byFM4-64 staining, from the tip towards the BIA along ‘track-like’ strands projecting out from the BIA itself (Fig. 1B,H,I;Movies 2, 5; and see below). Movement of material throughthe BIA itself was investigated by FRAP analysis (Fig. 2).Material was shown to be displaced from the anterior side ofthe BIA away from the tip by the subapical displacement ofbleached spots (Fig. 2A). The recovery of fluorescence at thelocation of the bleached areas appeared to be by incorporationof new stained membrane from the BIA-associatedaggregations into the BIA and displacement of bleachedmembrane away from the tip. The displaced bleached area alsoshowed slight recovery in fluorescence, probably by diffusion(Fig. 2A′). Displacement of material through the BIA wasdistinguished from simply diffusion of free dye by followingthe progress of two separate but adjacent bleached spots, whichremained distinctly separate as they migrated away from the

Journal of Cell Science 116 (13)

Fig. 2.Movement of material into and through the BIA of L. longiflorumpollentubes followed by confocal FRAP of FM4-64. (A) Bleaching within the BIA;(B) bleaching of the apical region. Sites of photobleaching indicated on brightfield images. Time is in seconds after bleaching. (A′) Fluorescence intensity atthe original location of the bleach site and ‘following’ the displaced region ofbleached fluorescence (see insert diagrams). (B′) Fluorescence intensity at anapical bleach site and within the BIA. Bars, 15 µm.

Fig. 3. A functional actin cytoskeleton is required to maintain theBIA and associated movements. Effects of (A) 5.0 µM cytochalasinD and (B) 2.0 µM Jasplakinolide on BFA (3.6 µM)-treated L.longiflorumpollen tubes labeled with FM4-64. Times are in seconds.Bars, 15 µm.

2711Pollen tube periodicity

tip. Recovery was slow, suggesting an absence of mixing andlimited free diffusion of dye within the BIA (Fig. 2A).

Photobleaching within the extreme apex supported theobservation that this was the source of stained membranetrafficked through the BIA (Fig. 2B). Following the apicalbleach and transient reduction in apical fluorescence there wasa clear transient drop in the fluorescence of the BIA thatsubsequently showed a general increase. Superimposed uponthe general increase in fluorescence there also appeared to bea periodic fluctuation in the brightness of the BIA (Fig. 2B′).It should also be noted at this point that the overall size ofthe BIA did not appear to steadily increase – suggestingthat material was also lost (discussed later). The periodicmovements were not halted by the bleach procedure.

Organisation and dynamics of the BIA are actindependentIn pollen tube tip growth the actin cytoskeleton is involved incytoplasmic streaming and vesicle transport (Hepler et al.,2001). The involvement of actin in the periodic movements ofmembrane (BIA-associated aggregations) between the extremeapex and subapical Brefeldin-A-induced structure wasinvestigated with drugs known to affect the actin cytoskeleton.

Cytochalasin-D applied to normally growing L.longiflorumpollen tubes at relatively low concentrations (1.5µM) rapidly arrests growth and disperses the apical FM4-64staining pattern (Parton et al., 2001). A similar treatment withCytochalasin D applied to BFA-treated cells had little effecton the staining pattern. Concentrations of 5.0 µM and higherwere required before obvious effects were noticed. At theseconcentrations the movement of material to the subapicalBIA was rapidly arrested and the structure itself dissipated(Fig. 3A).

The actin cytoskeleton ‘stabilising’ drug Jasplakinolide (Ouet al., 2002) at only 2 µM rapidly blocked the periodictrafficking of material to the BIA and disrupted the generalapical zonation but did not directly dissipate the membraneaggregation (Fig. 3B). Only with extended times afterapplication of Jasplakinolide (>5 minutes) did there appear tobe some dissociation of the BIA, by which time cytoplasmicstreaming had effectively halted. At 2 µM Jasplakinolide alsoblocked tip growth of normally extending pollen tubes.

The BIA incorporates markers of the apical vesicleaccumulation but not the ERHaving established the trafficking of material between the apexand BIA it was important to determine its nature. That BFAinterferes in vesicle trafficking and secretion in animal and

plant cells is now well documented, it was therefore reasonableto presume that the BFA induced apical reorganisation was theresult of endomembrane system disruption.

Fluorescent labels of membranes (FM4-64), lipid (Nile Red,not shown) and the cytosol (targeted GFP; 10,000 kDa dextran-dye conjugates; Figs 1, 4) showed by dye accumulation, or inthe case of the cytosolic markers, by dye exclusion, thatthe BIA consists of lipid as membranes. The associatedaggregations appeared to be of similar nature as theirmovements could be followed with the various labels.

In previous studies into BFA effects on pollen tubes, amembrane re-organisation was described in the apical regionand attributed to redistribution of ER and Golgi membranes onthe basis of EM analysis (Rutten and Knuiman, 1993;Geitmann et al., 1996). The origins of the BFA-inducedaggregations observed here in L. longiflorumpollen tubes wereinvestigated in vivo using ER and Golgi-targeted fluorescentproteins (Figs 5-7).

In untreated, rapidly growing tubes, mGFP5-ER (Haseloff

Fig. 4.Consecutive confocal images (interval 1.5 seconds) of a BFA(3.6 µM)-treated tube expressing cytoplasmic targeted GFP. Bar, 15µm.

Fig. 5.ER and Golgi localisation in growing L. longiflorumpollentubes shown by targeted fluorescent protein expression.(A-B) mGFP5-ER in (A) growing tube, sequential images (image/3seconds) showing the dynamic nature of ER; (B) onion epidermis,showing the typical reticulate ER structure. (C-E′) GT-EYFP: (C)Sequential images (image/2 seconds) of a growing tube revealingmovement of targeted structures; (D) high expression levels; (E) inonion epidermis; (E′) effects of BFA on labeling in onion epidermis.(F-G) GONST1-YFP expressed in growing L. longiflorumpollentubes: (F) Sequential images (as for C); (G) high expression levelsshowing labelling of the apical vesicle accumulation. Inserts showbright field images at one-third size. Bars, 15 µm (A,C,D,F,G);100µm (B,E).

2712

et al., 1997) expression clearly marked a region extending fromthe periphery of the apex back into more subapical regions(Fig. 5A). In median section there was less dense labellingwithin the ‘V-shaped’ apical clear zone and almost totalexclusion from the extreme apical (~3 µm) lens-shaped region,which corresponds to the apical vesicle accumulation aslabelled by FM4-64. More towards the apex, the mGFP5-ERdoes not label a clear tubular structural organisation, as haspreviously been encountered with ER labelling in tip-growingcells (McCauley and Hepler, 1990; Cheung et al., 2002).Further back, distinct interconnected strands could be seen.Time-course analysis and rapid focusing up and down showedthat mGFP5-ER localised to a dynamic 3D organisation (notshown), very different from the peripheral ER normallyencountered with ER labelling in epidermal cells, but inagreement with EM images of ER distribution in the pollentube tip (Pierson and Cresti, 1992; Lancelle and Hepler, 1992),fungal hyphae (Grove, 1978) and fern protonemata (Dyerand Cran, 1976). Very similar mGFP5-ER localisation wasobserved in tobacco pollen tubes (not shown). Expression of

mGFP5-ER in onion epidermis showed the typical reticulatecortical-ER distribution (Fig. 5B).

GT-EYFP (Clontech) expression (Fig. 5C) produced a brightpunctate pattern corresponding to roughly 1-2 µm diameterstructures associated with a weak background fluorescence ofa similar distribution to mGFP5-ER. The brightly labelledstructures were scattered throughout the subapical region ofpollen tubes but were effectively excluded from the wholeapical hemisphere. The distribution and movement of thebright structures differed significantly from the labelling ofmitochondria by Mitofluor Green (Fig. 1E). Movement of theGT-EYFP structures was slower and more stilted (based uponexamination of time-sequence data) than that of mitochondria.At high levels of GT-EYFP expression, growth was slightlyreduced and more sensitive to laser scanning while the level ofbackground ‘ER-like’ localisation was much higher (Fig. 5D).Expressed in onion epidermis, GT-EYFP clearly localised tothe ER and to small roughly spherical structures thatcorrespond to the expected distribution of Golgi bodies (Fig.5E-E′). Partial distribution of other Golgi-targeted fluorescent

proteins into the ER has been reported previously(Brandizzi et al., 2002; Saint-Jore et al., 2002).

The GONST1-YFP fusion protein, which targets to theGolgi in plant cells (Baldwin et al., 2001), produced avery similar punctate fluorescence to GT-EYFP in L.longiflorum pollen tubes (Fig. 5F). GONST1-YFPexpression lacked the ER-like background of GT-EYFPbut included a faint labelling of the V-shaped region ofthe apical vesicle accumulation. Labelling associatedwith the apical vesicle accumulation region was muchclearer with high levels of expression (Fig. 5G). Growthrate and morphology were generally unaffected by thisconstruct.

BFA treatment resulted in drastic redistribution of themGFP5-ER (Fig. 6A) leaving no sign of the labelling atthe periphery of the extreme apex. Redistributioncorrelated with the change in apical zonation observed inbrightfield and with FM4-64 staining. The redistributedmGFP5-ER was largely restricted to behind the site ofthe BIA. No labelling of either the BIA or associatedaggregations by mGFP5-ER was seen (Fig. 6A, >30minutes). This was confirmed by co-labelling with FM4-64 that showed these aggregations to be present (notshown). However, it was generally possible to make outthe staining of two fine strands originating from theextended region of dense subapical staining andprojecting towards the apex (Fig. 6A, 40 minute timepoint). These strands corresponded to the tracking lanes

Journal of Cell Science 116 (13)

Fig. 6. (A-C) Differences in ER and Golgi involvement in the BIA andassociated aggregates revealed by fluorescein protein expression in living L.longiflorumpollen tubes treated with 3.6 µM BFA. (A) mGFP5-ERredistribution (times in minutes). Arrowheads indicate fine structuresprojecting into the apex in the main picture and contrast adjusted insert.(B-C) Redistribution of Golgi markers GT-EYFP and GONST1-YFP (timesafter applying BFA in minutes). In all cases, before the 15-minute time-point tubes were still extending so images have been aligned from the tips.Images for time-points 40 minutes and longer are contrast adjusted toincrease brightness relative to earlier time points. Arrowheads in C indicatethe BIA. Inserts show bright field images at one-third size. Bars, 15 µm.

Fig. 7.GT-EYFPdistribution after 40minutes BFA (3.6 µM)treatment shows similarityto BFA effects on mGFP5-ER. The zoomed regionshows labeled ‘projections’extending from the site ofthe BIA towards the apex(arrowheads). Bar, 15 µm.

2713Pollen tube periodicity

of material passing from the apex to the BIApreviously described (Fig. 1H). Little clearindication of periodic trafficking events occurringcould be seen from the mGFP5-ER labelling;however, there was some evidence of periodicextension and retraction of the labelled ‘arm like’projections, described above, which correlated withthe movements of FM4-64-stained material.

BFA treatment of GT-EYFP-expressing pollentubes led to changes in the distribution, size andapparent numbers of the bright 1-2 µm structures,but even after growth arrest did not result in theircomplete disappearance (Fig. 6B). Soon after BFAapplication the bright structures could be seen toclump; subsequently, individual structures appearedenlarged. Redistribution of the weak background‘ER-like’ labelling was identical to that observedwith the mGFP5-ER. As with mGFP5-ER, no clearlabelling of the apical membrane aggregations or theBIA was seen, yet staining was again seen in finestrands that extended from the region of densesubapical staining towards the apex (Fig. 7).

The effects of BFA on the punctate structures ofGONST1-YFP expressing pollen tubes were thesame as for GT-EYFP (Fig. 6C). However with theGONST1-YFP construct, label appeared to beredistributed to the BIA and associated aggregations. Fromexamination of time-series images material was seen to trackbetween these compartments as described for FM4-64labelling (Fig. 6C; 60 minute time point).

Movements associated with the BIA exhibit a regularperiodicityFollowing the characterisation of the BFA-inducedphenomenon, in which we established that there is trafficking ofmembranes from the apex to a subapical membrane aggregationdependent upon an active actin cytoskeleton, we investigated theintriguing periodicity apparent in the trafficking of material.

Rapid time-course imaging using DIC confirmed that eventsoccurred with a definite period (Fig. 8A; Movie 5). Withimaging rates of 0.5-1 frames per second (FPS) it was possibleto show that in non-growing BFA-treated L. longiflorumpollen

tubes material entered the BIA with a period of 5-7 seconds(Table 2) sustained for at least 6 hours after their initialobservation 30 minutes to 1 hour after BFA treatment (Fig.8A′).

Table 2. Periodic behaviour of BFA-treated pollen tubes Period length Min/max period Number

Conditions (seconds)† (seconds) of tubes

3 µM/25°C 6.2±0.7 5.3/7.4 710 µM/20°C* 6.5±0.9 5/7.4 510 µM/30°C* 5.1±0.7 4.2/6 510 µM/25°C 6.2±0.8 4.8/7.4 720 µM/25°C 6.2±0.9 4.7/7.6 735 µM/25°C 6.5±0.8 5.8/7.5 5

*Paired data.†Data represent the mean±s.d.

Fig. 8.Periodic ‘trafficking events’ at the tip of BFA-treated (1 hour, 3.6 µM) L. longiflorumpollen tubes. (A-A′) DIC bright field image sequence and correspondingplot recording movement of material along the pollentube axis as pixel intensity changes in a ‘samplingwindow’ (highlighted in A). A single trafficking event isindicated in red. (B) As in A but with FM4-64 labelling.(B′) Periodic movements of FM4-64-stained materialalong the pollen tube axis plotted as pixel intensitychanges (gray circles) in a ‘sampling window’ highlightedin the insert (white rectangle) over time, each movementevent is marked with a vertical bar. The insert image isfalse-coloured to reveal the pattern of relative stainingintensity: red, high fluorescence; blue, low fluorescenceintensity. FM4-64 signal intensity within the BIA (blackrectangle in insert image) is also plotted (black squares).The two plots were aligned by a –2.5 second shift in theBIA intensity plot. Bars, 15 µm.

2714

Having established the sampling rate required to report theperiodicity, BFA-treated L. longiflorumpollen tubes stainedwith FM4-64 were imaged by low resolution confocalmicroscopy (1.5-3 FPS; Fig. 8B) in order to quantifyfluorescence intensity within the BIA. This approach revealedthat staining intensity of the BFA structure increased withfusion of the incoming membrane from the trafficking eventsbut subsequently dropped between fusion events giving riseto a periodic fluctuation in signal. The rise in signal in theBIA appeared to be concurrent with the fusion event;however, the way in which the two parameters weremeasured, with regions of interest at two different locations,introduced an apparent lag in fluorescence increase relativeto the recording of the movement of material (Fig. 8B′ andinsert diagram). In the example shown in Fig. 8B′, the plotof movement was shifted by 2.5 seconds relative to the plotof intensity to compensate for this artefact. Note in someplaces the occurrence of shouldered peaks on the intensitytrace. This is due to the two ‘sides’ of the approaching loopof material (Fig. 8B) arriving slightly out of sync.Throughout, the BIA did not visibly increase in size withincreasing numbers of fusion events. Coupled with thefluctuating fluorescence intensity (above) and FRAP data(Fig. 2) this confirms the loss of stained membrane from, orpassage through the BIA.

Growth rate fluctuations are consistently of longer periodthan that associated with the BIAThe period of trafficking events after BFA treatment was notcomparable with the regular growth fluctuations exhibited byL. longiflorumpollen tubes (~30 second period), which are ofsimilar magnitude to growth fluctuations recorded for otherspecies we tested under optimised growth conditions (Table 3;Fig. 9; see Materials and Methods) or the periodic movementsof ~25-40 seconds of the apical vesicle accumulation withinthe cytoplasm of the tip (Parton et al., 2001).

We investigated ways of perturbing the period of growth ratefluctuation without arresting growth and found that whilemoderate temperature shift had no significant effect, anincrement of 10°C could reproducibly raise growth rate andreduce period duration in L. longiflorumpollen tubes (paireddata t-test, P=0.05; Table 3; Fig. 10A). Heat treatmentconsistently increased apical diameter (Fig. 10B) and thetendency to apical rupture. Applying similar heat treatment toBFA-treated tubes also decreased period duration (paired datat-test, P=0.005; Table 2; Fig. 10C; Movie 6). The effects of

different BFA concentrations, between 3.0 and 35 µM, werealso directly compared; however no significant difference inperiod was found (t-test, P=0.05; Table 2).

Journal of Cell Science 116 (13)

Table 3. Periodic growth behaviour of pollen tubesGrowth rate†,‡ Period length Min/max period Number

Species (µm/minute) (seconds)‡ (seconds) of tubesL. longiflorum 15.5±5.2 30.4±8.6 16.5/41.3 12L. longiflorum20°C* 10.6±2.5 22.6±4.9 18.7/30.8 5L. longiflorum30°C* 16.5±2.7 16.4±3.1 13.2/21.4 5N. plumbaginifolia 6.1±0.9 41.8±12.6 24.8/73.5 12N. tabacum 5.6±1.3 32.4±6.6 21.3/45.3 8A. thaliana21°C 4±0.8 39±7.8 29.2/48 9

*Paired data.†Growth at standard temperature 25±1°C, unless otherwise stated.‡Data represent the mean±s.d.

Lilium l ong iflorum

Ave. growth rate 11 µm /min

Dis

tan

ce g

row

n -

µ m

0

10

20

30

40

50

0

5

10

15

20

25

Period ~ 35 sec Gro

wth

rat

e -

µm/m

in

A

Nicotiana plum baginifo lia

Ave. growth rate 7.36 µm/min

0

10

20

30

40

50

4

6

8

10

Gro

wth

rat

e -

µm/m

in

Dis

tan

ce g

row

n -

µm

Period ~ 45 sec

B

0

10

20

30

0

2

4

6

8

Gro

wth

rat

e -

µm/m

in

Dis

tan

ce g

row

n -

µ m Nicotiana tabacum

C

Arabidopsis thaliana

Ave. growth rate 4.05 µm/min

0 50 100 150 200 250 300

0

10

20

30

2.5

3.5

4.5

5.5

Time (s)

Gro

wth

rat

e -

µm/m

in

Dis

tan

ce g

row

n -

µ m

Period ~ 45-50 sec

D

Ave. growth rate 4.0 µm/minPeriod ~ 40 sec

Fig. 9.Periodic growth fluctuations for different pollen speciesdetermined from bright field image sequences. Distance grown andinterval growth rate (between consecutive images) are plotted againsttime.

2715Pollen tube periodicity

The periodic movements associated with the BIA are notdependent upon a corresponding calcium signalAs has been previously reported (Feijó et al., 2001), innormally growing cells a clear tip-focused gradient in [Ca2+]ccould be observed that fluctuated with the same period as thegrowth rate oscillation, whereas in BFA-treated cells, calciumimaging failed to show an obvious tip-focused calciumgradient (n=5) (Fig. 11). There was also no obvious fluctuationin [Ca2+]c at the extreme apex or more globally within the apexthat we could relate to either the ~30 second periodicity seenwith normal growth or the 5-7 second periodicity in BIA-associated aggregation movement (which continued as normalin the presence of dye; Fig. 11A; Movie 7). Note that regionsof interest were sampled and ratios calculated for time-coursedata at different locations within the apical region, but avoidingareas corresponding directly to the BIA and associatedaggregates where dye signal was poor due to exclusion (seeinsert image in Fig. 11A).

Discussion At present it is not fully understood how growth processes areorganised at the pollen tube tip to produce rapid growth withperiodic fluctuations in rate. Our previous work (Parton et al.,2001) suggested that determination of periodicity in growthrate fluctuation perhaps involves regulated membrane andregulated vesicle trafficking rather than exclusively thefluctuating ion movements that have received most attention.Here we have addressed this hypothesis by interfering with thenormal processes of vesicle trafficking and apical extension todetermine whether periodicity in membrane trafficking ismerely a consequence of periodic fluctuations in apicalextension.

BFA as a tool to dissociate tip growth, vesicle traffickingand periodicityThis study has made use of the ability of BFA to block tipgrowth and secretory activity in pollen tubes in a rapid andreproducible manner at relatively low concentrations (Ruttenand Knuiman, 1993; Geitmann et al., 1996; Ueda et al., 1996;Parton et al., 2001) while allowing other activities to continue.We did not find any significant difference in the final BFA-induced membrane aggregations over an ~10-foldconcentration range (Table 2). Unlike previous work on BFAeffects in pollen tubes, we have focused upon the dynamicbehaviour of living treated tubes in comparison with that ofnormal growing tubes.

While the mode of action of BFA in mammals is generallyaccepted, the situation in plants is more contentious (seeIntroduction) with variation observed in BFA effects indifferent studies of plant cells. One major argument is that BFAhas multiple sites of action or non-specific effects with higherconcentrations and longer treatment times (Driouich andStaehelin, 1997; Satiat-Jeunemaitre et al., 1996; Ritzenthaleret al., 2002; Saint-Jore et al., 2002). A recent re-analysis ofBFA effects (Ritzenthaler et al., 2002) in tobacco culture cellssuggests similar effects to those on mammalian cells. It ispossible that exact BFA effects depend upon the predominantvesicle trafficking processes operating, for example, in tip-growing pollen tubes this would be the secretion of materials

for growth. In this respect it is significant that pollen tubes ofboth dicots and monocots show similar BFA effects. Geldneret al. discuss multiple ARF-GEF’s in Arabidopsis, some ofwhich are predicted to be BFA insensitive, some sensitive(Geldner et al., 2003). They speculate that the overall BFAeffect on a cell is dependent upon the relative involvement ofBFA sensitive and insensitive ARF-GEF’s.

In our pollen tube work the low concentrations required, fastaction, reproducibility (Parton et al., 2001) and speciesindependence of BFA effects all argue that we are dealing witha fairly specific mode of action.

Non-growing pollen tubes can still exhibit polarisedorganisation and periodicityA distinctly polar cytological organisation and periodic growthrate fluctuations appear to be features exhibited by several tip-growing cell types (López-Franco et al., 1994; Wymer et al.,1997; Hepler et al., 2001). However, it is not understoodexactly how or why periodicity might be important or themechanism by which periodicity is established and maintained(Feijó et al., 2001). Indeed, regular periodicity is not

Fig. 10.Temperature alters periodic behaviour in L. longiflorumpollen tubes. (A) A 10°C rise shortens the period of growth pulsesand (B) increases tube diameter. (C) A 10°C rise shortens the periodof trafficking events associated with the BIA. Bar, 20 µm.

2716

established in Lilium (Messerli and Robinson, 1997) or N.tabacum(Zonia et al., 2002) pollen tubes until a certain age orlength is achieved and has not been recorded for all pollenspecies (Pierson et al., 1995). Nevertheless, periodicity ingrowth rate is clearly apparent during the rapid growth phaseof several pollen species, including Lilium, Nicotiana,Agapanthus(Camacho et al., 2000) and Arabidopsis. Mostresearch on pollen tube periodicity has focused upon howfluctuating ion entry and concentrations (most notably ofcalcium) relates to growth rate (Messerli et al., 2000;Holdaway-Clarke et al., 1997). Less attention has been paid tothe periodic nature of cytoplasmic movements (Zonia et al.,2002; Parton et al., 2001).

With BFA treatment we have a situation where, althoughapical extension is halted, expression of both polarised andperiodic behaviour continues. The periodic movements weobserve within the apical cytoplasm are clearly not a directconsequence of the periodic fluctuations in apical extension.Furthermore, our findings show that periodicity in the pollentube is not dependent upon rounds of active secretion of cellwall materials or weakening/tightening of the cell wall as thetip is extended, which are associated with cycles of stretchingof the PM (Holdaway-Clarke et al., 1997; Messerli et al.,2000). It is possible that periodic trafficking of membrane mayactually underlie the expression of growth rate periodicity.While Golgi-derived material is clearly implicated and plasmamembrane might be involved we were unable to determinewhether the phenomenon we have found has any links toendocytic activity.

Intriguingly, the periodicity of membrane traffickingexhibited by BFA-treated pollen tubes is of a significantlydifferent frequency from the periodicity associated with normaltip growth, in apical extension rate and the movements of the

apical vesicle cloud (5-7 seconds compared with ~30 seconds).However, we find it hard to imagine that the establishment ofthe observed periodicity is purely an artefact of BFA treatment.We think it more likely that the movements we see are relatedto those exhibited by the apical vesicle accumulation duringnormal growth and that BFA treatment has led to disruption offeed back mechanisms by which the normal period length isregulated (Feijó et al., 2001).

We have determined that period length in the pollen tube canbe modified experimentally. With a 10°C raise in temperaturewe were able to convincingly reduce the period length ofgrowth rate fluctuation in L. longiflorum pollen tubes andsimilar experimental manipulation also reduced period lengthin the BFA-associated phenomenon. These findings aresignificant in that they prove that we are dealing with a ‘simple’oscillatory mechanism, rather than a timing or ‘clockmechanism’ (Kippert and Hunt, 2001) that would need to betemperature compensated (i.e. not perturbed by changes intemperature).

BFA-treated tubes exhibit dynamic activities that can berelated to features of normal growthOur investigations of the BIA and movement of associatedaggregates by time-lapse and FRAP analysis show that we arenot observing simply the cycling of membrane aggregates inthe cytoplasmic flow but are following the occurrence of bothbulk membrane translocation and some degree of vesicletrafficking between membrane pools – features that we canrelate to the vesicle cloud movements observed during normalgrowth.

The path followed by the BIA-associated aggregatescorresponds to the bulk membrane flow seen with FM4-64

Journal of Cell Science 116 (13)

Fig. 11.Calcium ratio imaging of non-growing,BFA-treated (A) and growing (B) L. longiflorumpollen tubes. (A) Plots show the typicalmovement of material into the subapicalmembrane aggregation (closed circles; c.f. Fig.2) and the apical intracellular calciumconcentration (open squares: ratio OG 488/TR).Images are (top to bottom): a Texas Redfluorescence image (the BIA and associatedaggregates appears dark); the correspondingprojected TR image time-series displayed as asingle ‘negative’ image to show the path ofmovement of material; the corresponding ratioimage. The areas sampled to produce the plots(rectangles) and location of the BIA (circled)and associated aggregates (arrowhead) areindicated. (B) Untreated pollen tube: plots showthe typical oscillatory fluctuation in apicalintracellular calcium concentration duringgrowth (open squares: ratio OG 488/TR; thearea sampled is shown on the insert image) andcorresponding fluctuation in growth rate (closedcircles). Insert images show OG 488/TR ratioscorresponding to a consecutive minima andmaxima in the tip-focused cytosolic calciumgradient. Bars, 10 µm.

2717Pollen tube periodicity

staining of the apical vesicle accumulation of growing tubes(Parton et al., 2001). Furthermore this movement from theextreme apex in BFA-treated tubes was actin dependent(cytochalasin and jasplakinolide sensitive), as are the apicalvesicle accumulation movements of growing tubes (Hepler etal., 2001). The organisation and movement of the BIA andassociated material are consistent with our proposal of astructural organisation to the apical vesicle accumulationoccupying that region in a growing tube. Previous reportssuggest that BFA does not affect the actin cytoskeleton in plantcells (Satiat-Jeunemaitre et al., 1996).

The possibility of vesicle trafficking in the presence of BFAis supported by the occurrence of BFA-independent trafficking(Klausner et al., 1992) and coat protein activity (Jackson andCasanova, 2000). The fact that cytochalasin D effectivelydissipated the subapical BFA-induced structure suggests that itis not simply a mass of fused membrane but a gathering ofsmaller components with a structural organisation maintainedby actin cytoskeleton.

The nature of the trafficked material links the periodictrafficking of the BIA to the periodic movements of theapical vesicle accumulation Studies of BFA-effects on plant cells generally describeendomembrane rearrangements or ‘BFA compartments’. Theterm BFA compartment was first used to describe the re-organisation of the Golgi, without ER involvement, that occursin maize and onion roots after BFA treatment (Satiat-Jeunemaitre et al., 1996). However, a range of endomembranere-arrangements have been recorded that include variouscombinations between the ER, Golgi, secretory vesicles orendocytic vesicles (Driouich and Staehelin, 1997; Farquharand Hauri, 1997; Sanderfoot and Raikhel, 1999; Batoko et al.,2000; Geldner et al., 2001; Ritzenthaler et al., 2002).Membrane aggregations noted from previous studies on BFAin pollen tubes have generally been attributed to ER:Golgifusions with complete loss of the Golgi (Rutten and Knuiman,1993; Geitmann et al., 1996).

While what we report here appears to be similar to the above,in that there is re-organisation of the Golgi, without ERinvolvement, we continue to see a punctate distribution ofGolgi markers after BFA-treatments, which suggests Golgi arenot completely dissipated (Fig. 6B,C). In fact, the effects ofBFA on our membrane labels FM4-64 and GONST1-YFPsuggest that the membrane aggregations of BFA-treated pollentubes involve membranes contributing to the vesicleaccumulation of the pollen tube tip. In untreated cells, FM4-64, which labels neither the ER nor Golgi to any obviousextent, preferentially locates at the site of the apical vesicleaccumulation that is dissipated by BFA (Parton et al., 2001).The GONST1-YFP fusion protein appears to label the Golgi,in agreement with GT-EYFP, and additionally associates withthe apical vesicle accumulation (Fig. 5F,G), even reportingfluctuating movements similar to those of FM4-64 duringgrowth. GT-EYFP, while labelling both the Golgi and ER, doesnot associate with the apical vesicle accumulation, suggestingthat this construct is restricted to earlier Golgi compartmentsthan GONST1-YFP. After BFA treatment, of the two Golgimarkers, GONST1-YFP alone labels the BIA. An origin fromsecretory vesicle membranes is not inconsistent with previous

studies in pollen tubes (Geitmann et al., 1996; Ueda et al.,1996) that report pectin-containing membrane aggregates.

The likely development of the BIA from secretory vesiclesand membranes of the apical vesicle accumulation provides afurther plausible link between the behaviour of the BIA andthat of the apical vesicle cloud. Taken together with theparallels in what we see with the apical vesicle accumulationmovements of growing tubes and the trafficking events of BFA-treated tubes – despite the difference in the actual period ofmovement this suggests that in BFA-treated cells we are indeedseeing a periodicity in membrane trafficking pathways, relatedto the movements of the apical vesicle accumulation duringgrowth, which is revealed not to be simply a consequence ofthe periodicity in apical extension rate. It seems far less likelythat the ~7 second period represents periodic behaviournormally occurring in growing tubes yet unrelated to tip growthand ‘masked’ in the absence of BFA. What our experimentsare unable to distinguish is whether we are dealing withmembrane components of endocytic or exocytotic traffic.

Oscillatory behaviour in the pollen tube tip appears to beindependent of the oscillating tip-focused intracellularcalcium gradient normally associated with growthCalcium imaging revealed that BFA abolished the typicaloscillating tip-focused calcium gradient normally associatedwith growth. Although it has already been established thatdisruption of the tip-focused oscillating calcium gradientaccompanies tip growth arrest in pollen tubes (Pierson et al.,1996), our findings are interesting in that, in the absence ofboth the typical tip-focused calcium gradient or any obviousdetectable periodic fluctuation in apical calcium, there shouldstill occur regular periodic movements associated with theBIA.

The results from BFA treatment suggest that the normallyobserved oscillating tip-focused calcium gradient is not theunderlying basis of periodicity but possibly a consequence ofa more fundamental oscillator, yet they do not contradict theunderstanding of the strong relationship between secretion andthe calcium signal (Roy et al., 1999) or the effects of calciumon growth orientation (Malhó and Trewavas, 1996). Ourmeasurements do not, however, eliminate the possibility ofcontinued calcium fluxes at the apex or of intracellularcalcium signals of lower magnitude or confined more closelyto the membrane that we were unable to detect. Previously, Liet al. showed the dependence of tip growth, calcium entry andthe tip-focused gradient upon a Rop-GTPase in the Rho familyof small GTP-binding proteins (Li et al., 1999). Rho-GTPasesare known to interact with the cytoskeleton but also affectvesicle trafficking (Farquhar and Hauri, 1997). Theimportance of Rop-GTPase in pollen tube tip growth providesan indication of a molecular mechanism by which periodicityin cytoplasmic movements or vesicle trafficking couldunderlie oscillatory calcium signalling and oscillatory growthfluctuation.

ConclusionThe complexity of tip-growth is evident and the significance ofperiodicity intriguing (Feijó et al., 2001). Although periodicgrowth rate fluctuations do not appear to be essential to pollen

2718

tube apical extension, they are more wide spread than firstthought. The ‘conventional’ model for pollen tube growth hascentred upon the tip-focused calcium gradient co-ordinatingand directing polarity, orientation and growth rate (Pierson etal., 1996; Malhó and Trewavas, 1996). Recent work has begunto question this (Hepler et al., 2001; Zonia et al., 2001). Ourcurrent hypothesis, to explain how the periodicity in the apicalvesicle accumulation and the periodicity associated with theBIA are related to the periodicity of growth, is that there is anunderlying structure to the apical vesicle accumulation and itsmovements that could in some way be involved in organisingor regulating vesicle traffic and consequently extension at thetip (Parton et al., 2001). Here we have presented a novelperspective on the periodicity of pollen tube tip growthby demonstrating the existence of periodic behaviour incytoplasmic movements and membrane trafficking, and in theabsence of an obvious tip focused [Ca2+]c gradient, secretionor apical extension. These findings highlight the need to furtherinvestigate the significance of periodic movements of the apicalvesicle accumulation – the contribution of the elusiveendocytic component of membrane traffic to these movementsstill remains to be determined.

This work was supported by BBSRC postdoctoral fellowships (toR.M.P., M.K.W. and S.F.-P.) and funding from the COSMIC imagingfacility, University of Edinburgh (to R.M.P.). Thanks to Paul Dupreeand Michael Handford (University of Cambridge, UK), and Patrick J.Hussey (University of Durham, UK) for providing constructs. Thanksalso to Chris Hawes for useful advice, to Chiang-Shiong Loh(University of Singapore) for measurements of Nicotianagrowth ratesand Tony Collins (Babraham Institute, Cambridge) for advice onImageJ.

ReferencesAndersen, S. S. L. and Bi, G. (2000). Axon formation: a molecular model

for the generation of neural polarity. Bioessays22, 172-179.Baldwin, T. C., Handford, M. G., Yuseff, M.-I., Orellana, A. and Dupree,

P. (2001). Identification and characterisation of GONST1: a Golgi-localisedGDP-mannose transporter in Arabidopsis. Plant Cell13, 2283-2295.

Bartnik, E. and Sievers, A. (1988). In vivo observations of a sphericalaggregate of endoplasmic reticulum and of Golgi vesicles in the tip of fastgrowing Chara rhizoids. Planta176, 1-9.

Batoko, H., Zheng, H. Q., Hawes, C. and Moore, I. (2000). A Rab1 GTPaseis required for transport between the endoplasmic reticulum and Golgiapparatus and for normal Golgi movement in plants. Plant Cell 12, 2201-2217.

Blackbourn, H. D. and Jackson, A. P. (1996). Plant clathrin heavy chain:Sequence analysis and restricted localisation in growing pollen tubes. J. CellSci. 109, 777-786.

Brandizzi, F., Snapp, E., Roberts, A., Lippincott-Schwartz, J. and Hawes,C. (2002). Membrane protein transport between the ER and Golgi in tobaccoleaves is energy dependent but cytoskeleton independent: evidence fromselective photobleaching. Plant Cell14, 1293-1309.

Camacho, L., Parton, R. M., Trewavas, A. J. and Malhó, R. (2000). [Ca2+]cdistribution and oscillations in pollen tubes: comparison of different dyes,dye forms and loading methods using confocal imaging. Protoplasma212,162-173.

Cheung, A. Y., Chen, Y.-h., Glaven, R. H., de Graaf, B. H. J., Vidali, L.,Hepler, P. K. and Wu, H.-m. (2002). Rab2 GTPase regulates vesicletrafficking between the endoplasmic reticulum and the Golgi bodies and isimportant to pollen tube growth. Plant Cell14, 945-962.

Coelho, S. M., Taylor, A. R., Ryan, K. P., Sousa-Pinto, I., Brown, M. T.and Brownlee, C. (2002). Spatiotemporal patterning of reactive oxygenproduction and Ca2+ wave propagation in fucus rhizoid cells. Plant Cell14,2369-2381.

Cole, L., Davies, D., Hyde, G. J. and Ashford, A. E. (2000). Brefeldin Aaffects growth, endoplasmic reticulum, Golgi bodies, tubular vacuole

system, and secretory pathway in Pisolithus tinctorius. Fungal GeneticsBiol. 29, 95-106.

Davis, S. J. and Vierstra, R. D. (1998). Soluble, highly fluorescent variantsof green fluorescent protein (GFP) for use in higher plants. Plant Mol. Biol.36, 521-528.

Derksen, J., Rutten, T., Lichtscheidl, I. K., Dewin, A. H. N., Pierson, E. S.and Rongen, G. (1995). Quantitative analysis of the distribution oforganelles in tobacco pollen tubes: implications for exocytosis andendocytosis. Protoplasma188, 267-276.

Driouich, A. and Staehelin, L. A. (1997). The plant Golgi apparatus:Structural organization and functional properties. In The Golgi Apparatus(ed. E. G. Berger and J. Roth), pp. 276-299. Basel, Switzerland: Birkhäuser.

Dyer, A. F. and Cran, D. G. (1976). The formation and ultrastructure ofrhizoids on protonemata of Dryopteris borreriNewm. Ann. Bot. 40, 757-765.

Farquhar, M. G. and Hauri, H.-P. (1997). Protein sorting and vesicular trafficin the Golgi apparatus. In The Golgi Apparatus(ed. E. G. Berger and J.Roth), pp. 65-116. Basel, Switzerland: Birkhäuser.

Feijó, J. A., Sainhas, J., Hackett, G. R., Kunkel, J. G. and Hepler, P. K.(1999). Growing pollen tubes possess a constitutive alkaline band in theclear zone and a growth-dependent acidic tip. J. Cell Biol. 144, 483-496.

Feijó, J. A., Sainhas, J., Holdaway-Clarke, T., Cordeiro, M. S., Kunkel, J.G. and Hepler, P. K. (2001). Cellular oscillations and the regulation ofgrowth: the pollen tube paradigm. BioEssays23, 86-95.

Fischer-Parton, S., Parton, R. M., Hickey, P. C., Dijksterhuis, H. A.,Atkinson, H. A. and Read, N. D. (2000). Confocal microscopy of FM4-64as a tool for analysing endocytosis and vesicle trafficking in living fungalhyphae. J. Microsc. 198, 246-259.

Geitmann, A., Wojciechowicz, K. and Cresti, M. (1996). Inhibition ofintracellular pectin transport in pollen tubes by monensin, brefeldin A andcytochalasin D. Bot. Acta109, 373-381.

Geldner, N., Friml, J., Stierhof, Y.-D., Jürgens, G. and Palme, K. (2001).Auxin transport inhibitors block PIN1 cycling and visicle trafficking. Nature431, 425-428.

Geldner, N., Anders, N., Wolters, H., Keicher, J., Kornberger, W., Muller,P., Delbarre, A., Ueda, T., Nakano, A. and Jürgens, G. (2003). TheArabidopsis GNOM ARF-GEF mediates endosomal recycling, auxintransport, and auxin-dependent plant growth. Cell 112, 219-230.

Gleave, A. P. (1992). A versatile binary vector system with a t-DNAorganizational-structure conducive to efficient integration of cloned DNAinto the plant genome. Plant Mol. Biol. 20, 1203-1207.

Gomez, T. M. and Spitzer, N. C. (1999). In vivo regulation of axonextension and pathfinding by growth-cone calcium transients. Nature397,350-355.

Grove, S. N. (1978). The cytology of hyphal tip growth. In The FilamentousFungi, Vol. 3, Developmental Mycology(ed. J. E. Smith and D. R. Berry),pp. 28-50. New York: Wiley.

Haseloff, J., Siemering, K. R., Prasher, D. C. and Hodge, S. (1997).Removal of a cryptic intron and subcellular localization of green fluorescentprotein are required to mark transgenic Arabidopsis plants brightly. Proc.Natl. Acad. Sci. USA94, 2122-2127.

Hepler, P. K., Vidali, L. and Cheung, A. Y. (2001). Polarized cell growth inhigher plants. Annu. Rev. Cell Dev. Biol. 17, 159-187.

Heslop-Harrison, J. (1987). Pollen germination and pollen-tube growth. Int.Rev. Cytol. 107, 1-76.

Hodgkin, T. (1983). A medium for germinating Brassica pollen in vitro.Eucarpia, Cruciferae Newsletter8, 62-63.

Holdaway-Clarke, T. L., Feijó, J. A., Hackett, G. R., Kunkel, J. G. andHepler, P. K. (1997). Pollen tube growth and the intracellular cytosoliccalcium gradient oscillate in phase while extracellular calcium influx isdelayed. Plant Cell9, 1999-2010.

Jackson, C. L. and Casanova, J. E. (2000). Turning on ARF: the Sec7 familyof guanine-nucleotide-exchange factors. Trends Cell Biol. 10, 60-67.

Kippert, F. and Hunt, P. (2001). Ultradian clocks in eukaryotic microbes:from behavioural observation to functional genomics. Bioessays22, 16-22.

Klausner, R. D., Donaldson, J. G. and Lippincott-Schwartz, J. (1992).Brefeldin A: Insights into the control of membrane traffic and organellestructure. J. Cell Biol. 116, 1071-1080.

Kost, B., Spielhofer, P. and Chua, N. H. (1998). A GFP-mouse talin fusionprotein labels plant actin filaments in vivo and visualizes the actincytoskeleton in growing pollen tubes. Plant J. 16, 393-401.

Lancelle, S. A. and Hepler, P. H. (1992). Ultrastructure of freeze-substitutedpollen tubes of Lilium longiflorum. Protoplasma167, 215-230.

Journal of Cell Science 116 (13)

2719Pollen tube periodicity

Li, Y.-Q., Zhang, H.-Q., Pierson, E. S., Huang, F.-Y., Linskens, H. F.,Hepler, P. K. and Cresti, M. (1996). Enforced growth-rate fluctuationcauses pectin ring formation in the cell wall of Lilium longiflorumpollentubes.Planta 200, 41-49.

Li, H., Lin, Y., Heath, R. M., Zhu, M. X. and Yang, Z. (1999). Control ofpollen tube tip growth by a Rop GTPase-dependent pathway that leads totip-localised calcium influx. Plant Cell11, 1731-1742.

López-Franco, R., Bartnicki-Garcia, S. and Bracker, C. E. (1994).Pulsed growth of fungal hyphal tips. Proc. Natl. Acad. Sci. USA91, 12228-12232.

Malhó, R. and Trewavas, A. J. (1996). Localised apical increases of cytosolicfree calcium control pollen tube orientation. Plant Cell8, 1935-1949.

Malhó, R., Camacho, L. and Moutinho, A. (2000). Signalling pathways inpollen tube growth and reorientation. Ann. Bot. 89, 59-68.

McCauley, M. M. and Hepler, P. K. (1990). Visualisation of the endoplasmicreticulum in living buds and branches of the moss Funaria hygrometricabyconfocal laser scanning microscopy. Development109, 753-764.

Messerli, M. A. and Robinson, K. R. (1997). Tip localized Ca2+ pulses arecoincident with peak pulsatile growth rates in pollen tubes of Lilliumlongiflorum. J. Cell Sci. 110, 1269-1278.

Messerli, M. A. and Robinson, K. R. (1998). Cytoplasmic acidification andcurrent influx follow growth pulses of Lilium longiflorumpollen tubes. PlantJ. 16, 87-91.

Messerli, M. A., Creton, R., Jaffe, L. F. and Robinson, K. R. (2000).Periodic increases in elongation rate precede increases in cytosolic Ca2+

during pollen tube growth. Dev. Biol. 222, 84-98.O’Driscoll, D., Hann, C., Read, S. M. and Steer, M. W. (1993). Endocytotic

uptake of fluorescent dextrans by pollen tubes grown in vitro. Protoplasma175, 126-130.

Oparka, K. J., Murphy, R., Derrick, P. H. and Prior, D. A. M. (1991).Modification of the pressure-probe technique permits controlledintracellular microinjection of fluorescent probes. J. Cell Sci. 98, 539-544.

Ou, G. S., Chen, Z. L. and Yuan, M. (2002). Jasplakinolide reversiblydisrupts actin filaments in suspension-cultured tobacco BY-2 cells.Protoplasma219, 168-175.

Palacpac, N. Q., Yoshida, S., Sakai, H., Kimura, Y., Fujiyama, K., Yoshida,T. and Seki, T. (1999). Stable expression of human beta 1,4-galactosyltransferase in plant cells modifies N-linked glycosylation patterns.Proc. Natl. Acad. Sci. 96, 4692-4697.

Parton, R. M., Dyer, A. F., Read, N. D. and Trewavas, A. J. (2000). Apicalstructure of actively growing fern rhizoids examined by DIC and confocalmicroscopy. Ann. Bot. 85, 233-245.

Parton, R. M., Fischer-Parton, S., Watahiki, M. K. and Trewavas, A. J.(2001). Dynamics of the apical vesicle accumulation and the rate of growthare related in individual pollen tubes. J. Cell Sci. 114, 2685-2695.

Pelham, H. R. B. (1989). Control of protein exit from the endoplasmicreticulum. Ann. Rev. Cell Biol. 5, 1-23.

Pierson, E. S. and Cresti, M. (1992). Cytoskeleton and cytoplasmicorganisation of pollen and pollen tubes. Int. Rev. Cytol. 140, 73-125.

Pierson, E. S., Li, Y. Q., Zhang, H. Q., Willemse, M. T. M., Linskens, H.F. and Cresti, M. (1995). Pulsatory growth of pollen tubes: Investigationof a possible relationship with the periodic distribution of cell wallcomponents. Acta Bot. Neerl. 44, 121-128.

Pierson, E. S., Miller, D. D., Callaham, D. A., van Aken, J., Hackett, G.and Hepler, P. K. (1996). Tip-localized calcium entry fluctuates duringpollen tube growth. Dev. Biol. 174, 160-173.

Read, S. M., Clarke, A. E. and Basic, A. (1993). Stimulation of growth ofcultured Nicotiana tabacumW38 pollen tubes by poly ethylene glycol andCu(II) salts. Protoplasma177, 1-14.

Ridge, R. W., Uozumi, Y., Plazinski, J., Hurley, U. A. and Williamson, R.E. (1999). Developmental transitions and dynamics of the cortical ER ofArabidopsiscells seen with green fluorescent protein. Plant Cell Physiol.40, 1253-1261.

Ritzenthaler, C., Nebenführ, A., Movafeghi, A., Stussi-Garaud, C., Behnia,L, Staehelin, L. A. and Robinson, D. G. (2002). Reevaluation of the effectsof brefeldin A on plant cells using tobacco bright yellow 2 cells expressingGolgi-targeted green fluorescent protein and COP1 antisera. Plant Cell14,237-261.

Robineau, S., Chabre, M. and Antonny, B. (2000). Binding site of Brefeldin-A at the interface between the small G protein ADP-ribosylation factor 1(ARF-1) and the nucleotide-exchange factor Sec7 domain. Proc. Natl. Acad.Sci. USA97, 9913-9918.

Rohn, W. M., Rouille, Y., Waguri, S. and Hoflack, B. (2000). Bi-directionaltrafficking between the trans-Golgi network and the endosomal/lysosomalsystem. J. Cell Sci. 113, 2093-2101.

Roth, J. and Berger, E. G. (1982). Immunocytochemical localisation ofgalactosyle transferase in HeLa cells: codistribution with thiaminepyrophosphatase intrans-Golgi cisternae. J. Cell Biol. 93, 223-229.

Roy, S. J., Holdaway-Clarke, T. L., Hackett, G. R., Kunkel, J. G., Lord,E. M. and Hepler, P. K. (1999). Uncoupling secretion and tip growth inlily pollen tubes: evidence for the role of calcium in exocytosis. Plant J. 19,379-386.

Rutten, T. and Knuiman, B. (1993). Brefeldin A effects on tobacco pollentubes. Eur. J. Cell Biol. 61, 247-255.

Saint-Jore, C. M., Evins, J., Batoko, H., Brandizzi, F., Moore, I. andHawes, C. (2002). Redistribution of membrane proteins between the Golgiapparatus and endoplasmic reticulum in plants is reversible and notdependent on cytoskeletal networks. Plant J. 29, 661-678.

Sanderfoot, A. A. and Raikhel, N. V. (1999). The specificity of vesicletrafficking: Coat proteins and SNAREs. Plant Cell11, 629-641.

Satiat-Jeunemaitre, B., Cole, L., Bourett, T., Howard, R. and Hawes, C.(1996). Brefeldin A effects in plant and fungal cells: something new aboutvesicle trafficking? J. Micros. 181, 162-177.

Steer, M. W. and Steer, J. M. (1989). Tansley review no 16: Pollen tube tipgrowth. New Phytol. 111, 323-358.

Ueda, T., Tsukaya, H., Anai, T., Hirata, A., Hasegawa, K. and Uchimiya,H. (1996). Brefeldin A induces the accumulation of unusual membranestructures in elongating pollen tubes of Niciotiana tabacumL. J. PlantPhysiol. 149, 683-689.

Wakeley, P. R., Rogers, H. J., Rozycka, M., Greenland, A. J. and Hussey,P. J. (1998). A maize pectin methylesterase-like gene, ZmC5, specificallyexpressed in pollen. Plant Mol. Biol. 37, 187-192.

Wymer, C. L., Bibikova, T. N. and Gilroy, S. (1997). Cytoplasmic freecalcium distributions during the development of root hairs of Arabidopsisthaliana. Plant J. 12, 427-439.

Zonia, L., Cordeiro, S. and Feijó, J. A. (2001). Ion dynamics andhydrodynamics in the regulatin of pollen tube growth. Sex. Plant Reprod.14, 111-116.

Zonia, L., Cordeiro, S., Tupy, J. and Feijó, J. A. (2002). Oscillatory chlorideefflux at the pollen tube apex has a role in growth and cell volume regulationand is targeted by inositol 3,4,5,6-tetrakisphosphate. Plant Cell 14, 2233-2249.