ArabidopsisCAP1 – a key regulator of actin organisation ...jcs.biologists.org/content/joces/120/15/2609.full.pdf · establishes CAP1 as a fundamental facilitator of actin dynamics

2609Research Article

IntroductionCyclase-associated protein (CAP) was identified in S.cerevisiae as an interactor of adenylate cyclase (AC) (Field etal., 1990). Mutations in CAP/SRV2 not only affect theregulation of AC by Ras (Fedor-Chaiken et al., 1990; Shima etal., 2000) but also cause actin organisational phenotypes(Vojtek et al., 1991). Investigations into the biochemicalactivity of CAP in the context of the actin cytoskeleton hasdefined CAP as an actin-binding protein (ABP) capable ofassociating with monomeric actin and facilitating actintreadmilling (Balcer et al., 2003; Mattila et al., 2004). The C-terminus of S. cerevisiae CAP is required in vivo and in vitrofor the majority of cytoskeletal functions (Gerst et al., 1991;Mattila et al., 2004), while the N-terminus regulates ACactivation in vivo. The functional division between signallingand actin organisation has led to CAP being considered abifunctional protein.

CAP is conserved over a wide range of organisms. Cross-species complementation experiments have shown thatheterologous CAP can consistently complement S. cerevisiaeCAP-dependent cytoskeletal functions but not AC activation.The N-terminus of S. cerevisiae CAP is required to expose ACbinding sites to Ras (Shima et al., 2000). S. pombe also requiresCAP for AC activity (Kawamukai et al., 1992), but S. pombeAC is not activated by the Ras pathway. CAP in S. pombe mustfacilitate AC activation in a novel fashion and, consequently,the N-terminus of S. pombe CAP cannot complement S.

cerevisiae cap mutants or vice-versa (Kawamukai et al., 1992).CAP isoforms from other species are also unable tocomplement S. cerevisiae AC activation (Matviw et al., 1992;Vojtek and Cooper, 1993; Yu et al., 1994; Zelicof et al., 1993)and have been argued to operate in their own species-specificsignalling pathways (Hubberstey and Mottillo, 2002). Thecross-species association of apparently independent signallingand cytoskeletal activities might reflect an as yet unidentifiedfunctional integration of the two roles (Vojtek and Cooper,1993).

In addition to S. cerevisiae and S. pombe, CAP mutants havebeen identified and characterised in Drosophila (Baum et al.,2000; Benlali et al., 2000), in Dictyostelium (Noegel et al.,1999) and in mammals, where RNAi suppression of CAPfunction has been performed (Bertling et al., 2004).Phenotypes shared by these mutants are reductions in polarisedcell morphology and cell motility coinciding withdisorganisation of actin-rich structures. At the level of tissueorganisation the cap phenotypes reveal a requirement for CAPin multicellular developmental signalling pathways. InDictyostelium, CAP is required to perpetuate the cAMP relaysignal to organise fruitbody formation (Noegel et al., 2004),and in Drosophila CAP is essential for Hedgehog-mediated eyedevelopment (Benlali et al., 2000).

Homologues of CAP have been identified in plants (Barreroet al., 2002; Kawai et al., 1998). The single Arabidopsisisoform has been shown to have the ability to bind actin and

Maintenance of F-actin turnover is essential for plant cellmorphogenesis. Actin-binding protein mutants reveal thatplants place emphasis on particular aspects of actinbiochemistry distinct from animals and fungi. Here weshow that mutants in CAP1, an A. thaliana member of thecyclase-associated protein family, display a phenotype thatestablishes CAP1 as a fundamental facilitator of actindynamics over a wide range of plant tissues. Plantshomozygous for cap1 alleles show a reduction in statureand morphogenetic disruption of multiple cell types. Pollengrains exhibit reduced germination efficiency, and cap1pollen tubes and root hairs grow at a decreased rate and toa reduced length. Live cell imaging of growing root hairsreveals actin filament disruption and cytoplasmicdisorganisation in the tip growth zone. Mutant cap1 alleles

also show synthetic phenotypes when combined withmutants of the Arp2/3 complex pathway, which furthersuggests a contribution of CAP1 to in planta actindynamics. In yeast, CAP interacts with adenylate cyclasein a Ras signalling cascade; but plants do not have Ras.Surprisingly, cap1 plants show disruption in plantsignalling pathways required for co-ordinated organexpansion suggesting that plant CAP has evolved to attainplant-specific signalling functions.

Supplementary material available online athttp://jcs.biologists.org/cgi/content/full/120/15/2609/DC1

Key words: Actin, CAP, Arabidopsis

Summary

Arabidopsis CAP1 – a key regulator of actinorganisation and developmentMichael J. Deeks1,*, Cecília Rodrigues1,2,*, Simon Dimmock1,*, Tijs Ketelaar1, Sutherland K. Maciver3,Rui Malhó2 and Patrick J. Hussey1,‡

1The Integrative Cell Biology Laboratory, School of Biological and Biomedical Sciences, Durham University, South Road, Durham, DH1 3LE, UK2Universidade de Lisboa, Faculdade Ciências, Instituto Ciência Aplicada e Tecnologia, Lisbon, Portugal3Centre for Integrative Physiology, School of Biomedical Sciences, University of Edinburgh, George Square, Edinburgh, EH8 9XD, UK*These authors contributed equally to this work‡Author for correspondence (e-mail: [email protected])

Accepted 15 May 2007Journal of Cell Science 120, 2609-2618 Published by The Company of Biologists 2007doi:10.1242/jcs.007302

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to complement the cytoskeletal defects of CAP-deficient yeast(Barrero et al., 2002), which suggests that plant CAP proteinshave the potential to regulate the actin cytoskeleton, but theendogenous role of CAP in plant cells has remaineduncharacterised.

The plant actin network is required for a variety of processesincluding the regulation of transpiration, pathogen defenceresponses, and (most visibly) growth and development(reviewed by Hussey et al., 2006). Disruption of actinpolymerisation by drugs (Baluska et al., 2001), and by someloss-of-function, gain-of-function and misexpression actinmutants (Gilliland et al., 2002; Kandasamy et al., 2002;Nishimura et al., 2003) results in dwarf plants with restrictedand uncoordinated cell expansion phenotypes. Sequenced plantgenomes contain homologues of many ABPs, some of whichhave been shown to modulate actin behaviour in planta. Withthe exception of AIP1 (Ketelaar et al., 2004a), most plant ABPmutants and suppression constructs affect the morphogenesisof only a variable subset of cell types. The tissue-specificnature of formin phenotypes (Deeks et al., 2005; Ingouff et al.,2005; Yi et al., 2005) and profilin (McKinney et al., 2001) canbe considered to be a symptom of large gene families with thepotential for genetic redundancy, but the relatively mildphenotypes of components of the Arp2/3 complex (Mathur etal., 2003) together with the unexpectedly severe ArabidopsisAIP1 phenotype suggests that plants place functional emphasisupon individual classes of ABPs in a pattern that differs fromanimals and fungi. Here, we show that the Arabidopsishomologue of CAP (CAP1) is essential for the development ofmultiple cell types and that null mutant phenotypes of thesetissues correlate with actin organisational defects. Moreover,deactivation of CAP1 alters the growth behaviour of multipleorgans in a novel fashion resulting in curled inflorescences andmeandering roots consistent with CAP1 contributing to thefunction of plant-specific signalling pathways.

ResultsDisruption of CAP1 affects plant developmentThe biological role of the actin-binding protein CAP1 wasinvestigated through the characterisation of T-DNA insertionalleles SALK_112802 (designated cap1-1) and GABI-KAT453G08 (cap1-2; Fig. 1). Plants homozygous for the insertionalleles were identified among segregating populations usinggenomic PCR. RT-PCR designed to amplify full-length CAP1demonstrated an absence of CAP1 transcript in cDNAgenerated from cap1-1 homozygote and cap1-2 homozygoteRNA templates (Fig. 2A). No truncated CAP1 mRNA wasdetected in mutant plants.

Plants homozygous for either cap1-1 or cap1-2 showed aconsistent co-segregating pleiotropic phenotype (n=70 and 65,respectively). The absence of the phenotype in heterozygotesand F1 plants derived from backcrosses defines the cap1-1 andcap1-2 alleles as recessive. F1 plants from crosses betweencap1-1 and cap1-2 homozygotes confirmed allelism. Allidentified aspects of the phenotype are present in both cap1-1and cap1-2 homozygote plants. The influence of maternalgenotype on seedling and plant growth behaviour wasnegligible, as plants descended from outcrosses with cap1homozygote plants as the maternal parent appeared normal.Moreover, cap1 homozygote plants descended fromheterozygote parents exhibited all phenotypic aspects.

Mutant plants homozygous for cap1 alleles have severelyreduced stature (Fig. 2B). Rosette diameters of mutant plantsmeasured at 22 days after germination (DAG) are reducedcompared with wild-type controls (20.7, 14.3 and 15.3 mm forWT, cap1-1 and cap1-2, respectively; n>30 for all lines)although the mean number of rosette organs is equal. Rootgrowth is also impaired in cap1 seedlings, with a 44%reduction in primary root length compared with wild-typeplants after a 5-day growth period on vertical plates. Wild-typeand cap1 plants grown in parallel initiated inflorescencessimultaneously but differed in rates of inflorescence growth(Fig. 2B). At 35 DAG wild-type, cap1-1 and cap1-2inflorescences measured a mean height of 139.9, 89.9 and 90mm, respectively. Inflorescences of cap1 plants produce floralbuds at a slower mean rate than wild-type inflorescences,contributing to height differences. Epidermal peels taken fromsynchronous stem internodes of cap1 and wild-typeinflorescences show a reduction in cell elongation (Fig. 2C,D).

Mutant cap1 pollen grains show reduced fertilityComparison of microarray expression analysis experimentshighlights maturing pollen grains as a major site of CAP1expression. The viability of pollen with mutant cap1 alleleswas assessed in vitro. Pollen grains and the tubes they produceprovide a convenient model to study highly polar growthprocesses. Pollen derived from mutant plants showed areduction in the rate of germination after 24 hours ofincubation in growth medium when compared to wild-typepollen (Fig. 3A-C). The growth rates of tubes successfullyproduced by mutant pollen grains were compared with wild-type tubes 5 hours after the initiation of germination (Fig. 3E)and were found to grow at a mean speed of approximately 1.0�m per minute, almost one-third of the rate of wild-typegrowth (2.8 �m per minute). After 24 hours of growth in vitromutant pollen tubes do not reach the same terminal lengths aswild-type tubes (Fig. 3D).

The growth phenotype of cap1 pollen tubes was confirmedin vivo using pollen grains germinated on intact flowers.

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cap1-1

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Fig. 1. Location of the T-DNA inserts in Arabidopsis CAP1(At4g34490). Translated sections of exons of Arabidopsis CAP1 arerepresented by boxes, and introns are represented by horizontal lines.T-DNAs are not drawn to scale. Primer 1 (CAP28F) combined withprimer 2 (CAP28R) was capable of amplifying CAP1 cDNA fromazygous plants but not from cap1-1 or cap1-2 homozygote plants(see Fig. 2). Products could be amplified with primers 1 and 2combined with T-DNA primers (3 and 4, respectively) using theappropriate homozygote plant genomic template, but not from acDNA template, which suggests the absence of processed CAP1:T-DNA fusion transcripts in homozygote mutant plants.

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Emasculated wild-type stigmas were fertilised with eitherwild-type or cap1 pollen and after a minimum of 4 hours weredissected, fixed and stained with aniline blue to assess pollenviability. After 4 hours, wild-type pollen tubes had traversedmost of the style, whereas mutant pollen tubes had yet topenetrate the stigma (Fig. 4). Mutant tube penetration wasbeginning to occur by 5.5 hours, by which time pioneeringwild-type tubes were in contact with ovules (Fig. 4). Flowersobserved 24 hours after pollination showed that mutant pollentubes were capable of eventually reaching ovules after asufficient growth period.

If the poor viability of mutant pollen is due to the debilitatingeffect of the male gametophyte inheriting a cap1 mutant allele,then the transmission frequency of cap1 alleles withinsegregating populations is likely to be reduced. The frequencyof F2 cap1-1 and cap1-2 homozygotes from F1 heterozygoteparents is low (3% compared with an expected value of 25%).Progeny of heterozygote plants were not observed to have anincrease in mortality to explain the absence of homozygotes,which suggests that a fertility problem was causing the lowfrequency of cap1 mutants. Genotyping of F1 plants generatedby crosses between wild-type plants and cap1 heterozygotesshows that the fertility of cap1 pollen is reduced: wild-typeplants fertilised with pollen from cap1 heterozygote plantsshow a cap1 transmission frequency of 1.2% (n=85). Bycontrast, the reciprocal cross shows a transmission frequencyof 44% (n=41). This indicates that pollen grains with a cap1genotype display a fertilisation handicap when competingagainst wild-type pollen generated by the same parent, as anovule is nearly 50 times more likely to be fertilised by a pollengrain inheriting a wild-type allele.

F-actin is disrupted in cap1 mutantsMutant cap1 lines were crossed with plants carryingGFP:FABD2, a construct consisting of the second actin-binding domain of fimbrin fused to GFP under the control ofthe CaMV 35S promoter, to identify possible actin cytoskeletaldisruption associated with the developmental abnormalities ofcap1 mutants. Observing the pollen tube cytoskeleton withGFP:FABD2 was found to be prohibited as the CaMV 35Spromoter does not stimulate expression within thegametophyte. Instead, we compared the actin arrays of roothairs, a sporophytic tissue used as a model for tip growth. Roothairs of cap1 homozygotes are severely shortened, bulbous,waved and occasionally branched when compared with wild-type root hairs grown in equivalent conditions (Fig. 5A,B).Mean growth speed is reduced to 0.36 �m per minute formutant hairs compared with 0.79 �m per minute for wild-typehairs.

Elongating wild-type root hairs contain a population oflongitudinally oriented actin cables within the shank of the hairthat disperse approximately 10 �m from the growing tip (Fig.5C). Dynamic F-actin populations at the tip are essential fordirecting and facilitating Golgi-derived vesicle fusion to theplasma membrane of the growth zone. Induced stabilisation ofF-actin and subsequent invasion of the apical clear zone byactin bundles has been found to correlate with cessation ofgrowth (Ketelaar et al., 2004a; Ketelaar et al., 2004b). Growinghairs lacking CAP1 contain short F-actin bundles thatcongregate at the cell cortex (Fig. 5D). These F-actin bodiesoften appear as aggregates rather than defined bundles (Fig.

5C,D). Frequently, the cortical F-actin aggregates extend to thevery tip of growing cap1 root hairs (Fig. 5E,F). The absenceof organised long actin cables coincides with an apparent

Fig. 2. RT-PCR shows that CAP1 is absent in plants homozygous foralleles cap1-1 and cap1-2 (A). The full-length CAP1 transcript canbe amplified from plants with a wild-type CAP1 allele (1), but notfrom plants homozygous for either cap1-1 or cap1-2 (2). Controlindividuals are azygous plants from populations segregating cap1-1or cap1-2. Primer combinations are illustrated in Fig. 1. All plantswere successful templates for amplifying the control GAPCtranscript (upper gel). Plants homozygous for cap1 alleles showgrowth deficiencies when compared with wild-types (B), the mostobvious of which is a reduction in the rate of inflorescencedevelopment (plants were photographed at 53 DAG; bar, 5 cm).Epidermal peels taken from primary inflorescences between thesecond and third developing silique at 42 DAG demonstrate thatwild-type cells (C) are further elongated than equivalent cap1 cells(D). Individual primary inflorescences chosen for comparison boreequal numbers of mature lateral organs. Wild-type GFP:FABD2primary inflorescence epidermis cells (E) have a parallelarrangement of fine actin cables along the axis of cell expansion.Mutant epidermis (F) contains shorter F-actin bundles poorly alignedwith respect to the axis of growth. Bars, 200 �m.

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reduction in long range transport. Labeling of mitochondriausing mitotracker red showed that the wild-type pattern ofreverse fountain streaming of organelles (supplementarymaterial Movie 1) is not apparent in immature cap1-1 andcap1-2 root hairs (supplementary material Movie 2). Growingmutant root hairs often contain large zones of cytoplasmbetween the growing tip and central vacuole that are depletedin F-actin relative to the accumulations at the cortex (Fig. 5D).

Morphological abnormalities associated with F-actindisruption also occur in cap1 cell types where expansion islocalised to ‘diffuse’ zones of cell wall. The epidermal cellsof cap1 inflorescences are shorter with respect to thelongitudinal axis of the inflorescence than wild-type cells(Fig. 2C,D) and contain a relatively sparse population ofpoorly aligned F-actin bundles. Trichome cells of the leafepidermis also grow in a diffuse manner (Schwab et al., 2003)

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Fig. 3. Wild-type pollen grains incubated for 24 hours invitro in the presence of pollinated stigmas germinate anddevelop tubes (A). Pollen from cap1-1 and cap1-2 plantsshows a visibly lower frequency of germination andreduced tube development (B). Analysis of largernumbers of pollen grains (C) shows that the germinationrate of mutant pollen grains is less than half that of wild-type grains (n>800 for all genotypes). The mean lengthof pollen tubes after 24 hours (D) is similarly reduced(n>180). The growth speed of pollen tubes wascompared at 5 hours after exposing the pollen grains togermination medium (E). The mean speed of mutantgrains is reduced to nearly a third that of wild-type(n=32 for wild-type, n=17 for cap1).

Fig. 4. The in vitro growth behaviour of pollen grains of different genotypes was confirmed in vivo. Flowers were pollinated and afterincubation were dissected and stained with aniline blue. After 4 hours (left panel) large numbers of wild-type pollen grains have germinatedand produced intensely staining callose deposits. Many wild-type pollen tubes have grown the length of the style tissue and are beginning toenter the ovary. A minority of cap1 pollen grains show signs of germination at this time point, and only auto-fluorescence from vascular tissueis visible within the style. Stigmas stained at 5.5 hours (centre panel) after pollination with wild-type pollen contain a significant number ofpollen tubes developing callose plugs (white arrows) within the ovary, and some tubes are contacting ovules. A greater proportion of mutantpollen grains are germinating but tube growth is still retarded. Stigmas stained after 24 hours of pollen tube growth (right panel) shows thatsome mutant pollen tubes do eventually contact ovules. Bars, 200 �m.

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and are sensitive to disruptions in actin turnover(Mathur et al., 1999; Szymanski et al., 1999).Arabidopsis leaf trichome cells exposed to actindepolymerising drugs or produced by plantshomozygous for null alleles of components ofthe Arp2/3 complex and associated signallingpathway display a ‘distorted’ phenotypeconsisting of bloating and twisting of trichomestalks and branches. Trichomes from cap1plants display a weak distorted phenotype: cap1trichome branches are mildly twisted, and stalkinter-branch zones are often excessivelyelongated (Fig. 6A,B). The angle betweentrichome branches is also affected in a largeproportion of trichomes that otherwise wouldhave a wild-type appearance (Fig. 6A-C).Comparison of developing trichomes expressingGFP:FABD2 identified frequent excessiveaccumulation of F-actin in the core ofelongating branches (Fig. 6D,E) that does notresemble the cohesive network of longitudinallyaligned cables observed in the wild-type. Thisunusual F-actin array correlates with brancheswith diameters broader than those of wild-typebranches of a comparable age. The phenotype isprevalent in trichomes between developmentalstages 4 and 5, where branches are undergoingrapid expansion, but excessive F-actinaccumulation within central cytoplasmic regionscan be identified in trichomes as young asdevelopmental stage 2. The central bundles ofcap1 trichomes show some resistance to theaction of the actin-deploymerising druglatrunculin B (data not shown), which possiblyexplains the absence of enhanced sensitivity ofcap1 trichome morphogenesis to latrunculin Btreatment (see supplementary material Fig. S2).The redistribution of F-actin in cap1 mutanttrichomes is the reverse of microfilamentredistribution in mutant root hairs, whichsuggests fundamental differences in thecytoskeletal organisation of tip growing anddiffusely growing plant cells.

Synthetic phenotypes are produced between CAP1alleles and the Arp2/3 complex pathwayThe cap1 mutant alleles were crossed with previouslycharacterised null alleles of SCAR2 and ARP2 (Basu et al.,2005; Le et al., 2003) to search for novel syntheticphenotypes that would reveal new functional roles for CAP1and the Arp2/3 pathway during plant development. Nullalleles of the Arp2/3 activator SCAR2 display a weakdistorted trichome phenotype (Basu et al., 2005; Zhang et al.,2005). The scar2-1/cap1-1 and scar2-1/cap1-2 doublehomozygotes show an enhanced trichome phenotype withincreased trichome distortion greater than either singlemutant (Fig. 7A), which suggests that the actin-bindingproteins SCAR2 and CAP1 act in parallel to control trichomebranch expansion. No further synthetic phenotypes weredetected in other tissues. The distorted shapes of arp2-1/cap1-1 and arp2-1/cap1-2 trichomes are similar to those of

arp2 single mutants making ARP2 epistatic with respect toCAP1 during trichome development. This is expected asmutants of Arp2/3 complex components resemble trichomesgrown in the presence of high concentrations of actindepolymerising drugs, and therefore may represent an actinphenotypic zenith of morphological distortion. Doublemutants of either arp2-1/cap1-1 or arp2-1/cap1-2 showed agreater reduction in plant stature and a more severe inhibitionof root hair elongation (Fig. 7B). The root hairs of the doublemutant are arrested at the stage of bulge expansion at thesurface of the root epidermis and do not initiate tip growth.Drug studies have previously shown that bulge formation isnot as dependent upon the actin cytoskeleton as the later stageof tip growth (Baluska et al., 2000) explaining the arrest ofthe ABP double at this stage. Under standard growthconditions arp2 root hair development has previously beendescribed as normal (Mathur et al., 2003) and no

Fig. 5. When compared with wild-type root hairs (A), cap1 root hairs (B) are short,bulbous, and occasionally waved or branched (Bars, 200 �m). Growing wild-typehairs expressing GFP:FABD2 (C) have longitudinal actin cables within theproximal cytoplasm aligned with the axis of growth. F-actin forms a diffusedynamic network at the growing distal end of the hair that regulates vesicle fusionto the tip. In cap1 growing hairs (D) the diffuse tip network is replaced by F-actinaggregates (brightly labelled by GFP:FABD2), which can be observed at the verytip of the hair. Bars, 20 �m. Long actin bundles are absent from the central regionsof the cytoplasm and instead F-actin can be found in shorter accumulationsrestricted largely to the cell cortex. Imaging of the very tip of these growing roothairs shows the presence of GFP:fimbrin in bright aggregates at the cortex (F) inzones normally free of F-actin (E; bars, 5 �m).

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morphological abnormalities were identified in arp2-1 singlemutant root hairs grown as controls in parallel with doublemutants. Despite the absence of severe synergisticphenotypes in the double mutants, the additive arp2/cap1 roothair phenotype shows that the Arp2/3 pathway has afundamental role in root hair tip growth that is only revealedin a cap1 background.

CAP mutants are affected in co-ordination of organgrowthIn addition to being retarded in length, cap1 inflorescences curlduring bolting and exhibit alterations in the direction ofexpansion that create ‘kinks’ in the stem (Fig. 8A comparedwith Fig. 8B). The changes in growth angle occur at nodes,creating corners at points of lateral organ development.Zigzagging of the inflorescence has been reported ingravitropic mutants, but cap1 inflorescences remaingravitropic. Also unlike gravitropic mutants, cap1 secondaryinflorescences regularly achieve 360 degree loops relative tothe vector of primary inflorescence expansion (Fig. 8B). Thelooping process begins with secondary inflorescence headsbending downwards the gravity vector. At any one moment intime 37% of cap1 inflorescence heads are at an angle lowerthan the gravitational horizon (n=307). In the sameenvironmental conditions only 1% of wild-type inflorescenceheads grow at an angle below the same threshold (n=279).6.5% of cap1 inflorescence heads over a period of 7 daysachieved a complete 360 degree rotation relative to the axis oftheir own stem. The inflorescence rotation rarely exceeds onecomplete loop and is a temporary phenomenon; affected

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Fig. 6. ESEM images of rosette leaves demonstrates that the majorityof Columbia ecotype wild-type leaf trichomes have three branches(A). Mutant trichomes (B) show abnormal branch angles, twistedbranches, and expanded inter-branch zones [e.g. see trichomelabelled (i)]. Bars, 500 �m. The most prevalent aspect of the mutanttrichome phenotype is an increased variation in branch angles. Ahistogram (C) divided into bins of 10 degrees from 0 to 240 showsthat the majority of wild-type three-prong branches are separated byan angle of approximately 120°. Trichomes of mutant plants show awider spread of angles with extremes of 23 and 220 degrees. Thenumber of measurements was 90 for all lines and all trichomes weretaken from the sixth rosette leaf at 16 DAG. Imaging of GFP:FABD2in wild-type trichomes (D) and cap1 trichomes (E) aged betweendevelopmental stages 4 to 5 shows that F-actin in mutant trichomesaccumulates within the core of expanding branches (E; bars, 20 �m).

Fig. 7. Mutants homozygous for both cap1-1/cap1-2 and scar2-1show an enhancement of the distorted phenotype greater than eithersingle mutant (A), with increased bulging and twisting of branchesand inter-branch zones (ii). Mutants homozygous for both cap1-1/cap1-2 and arp2-1 have root hairs that do not progress beyondbulges on the surface of trichoblasts (B), unlike the cap1 singlemutants, which initiate tip growth. Root hairs of an arp2-1homozygote at the same magnification are shown as a control. Bars,200 �m.

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inflorescences uncurl hours to days after loop completion(supplementary material Movie 4). The pedicles of floralorgans are also susceptible to curling (Fig. 8B) but thesedistortions are permanent. Time-lapse recording of wild-typeand cap1 plants revealed that cap1 inflorescences do notundergo rotational circumnutation movements (supplementarymaterial Movies 3, 4) but instead cap1 inflorescence-headsoscillate at irregular intervals within the vertical plane. Ananalogous phenotype can be observed in roots; cap1 roots areunable to grow in a straight line across the horizontal surfaceof agar medium (Fig. 8D,F), yet remain gravitropic.Microscopic analysis did not reveal twisting of epidermal cellfiles within affected organs, and the chirality of inflorescencecurls occurs randomly between individual plants and betweeninflorescences of the same plant. These aspects of the cap1phenotype indicate a loss of coordination in organ expansion.Bending of organs is achieved in plants by simultaneousdifferential expansion of opposed cell layers. Loss ofcircumnutation movements and initiation of novel curlingmotion can result from either corruption of growth signals or

the interpretation of these signals in target tissues indicating aninvolvement of CAP in as yet unknown plant signallingpathways.

DiscussionEndogenous CAP1 is required for Arabidopsis actinorganisationDisruption of the CAP1 gene causes phenotypes in multipletissues that correlate with disturbances in the actincytoskeleton. Elongating epidermal cells, tip growing cells, andtrichomes show severe morphological abnormalities andunusual aggregation of F-actin. We have shown that theArabidopsis CAP homologue is an actin monomer bindingprotein (supplementary material Fig. S1), and recent work hasshown that CAP1 directly accelerates the exchange of ADP forATP by actin monomers (Staiger and Blanchoin, 2006;Chaudhry et al., 2007) filling a functional space in actinbiochemistry left by the absence of plant profilin nucleotideexchange activity. Drug studies have long shown that cap1plant phenotypes are compatible with a suppression of actinbiochemistry – sequestering, capping, or stabilising actin (withlatrunculin B, cytochalasin D, or jaspokinolide, respectively)produces a similar suite of defects. A previous study has shownthat the overexpression of plant CAP disassembles F-actinarrays in vivo and causes severe growth defects (Barrero et al.,2002), which is surprising when considering that theoverexpression of CAP in other organisms does not result ingross phenotypic abnormalities. We have established that thispotential for CAP1 to influence actin biochemistry is affirmedby the function of endogenous plant CAP in actin-dependentgrowth processes.

Reconciling CAP1 biochemistry with the CAP1phenotypeThe visible disruption to the Arabidopsis actin cytoskeletonresulting from absence of CAP1 consists of an F-actin re-arrangement into short bundles or aggregates that retain anunusual position within their respective cells types. Shortbundles congregate to form a dense actin ‘core’ in expandingtrichome branches while in growing root hairs actin bundlesdiminish and withdraw to the cell cortex. The formations of F-actin adopted in cells lacking functional CAP vary fromorganism to organism. In S. cerevisiae the appearance of theF-actin arrays of dividing cells undergoes only a subtlealteration, as both actin patches and cables remain intact. Actinpatch distribution is perturbed (Field et al., 1990) and the ASH1mRNA polar marker is not anchored correctly after beingtransported along actin cables (Baum et al., 2000) possiblyindicating a subtle cable defect. Animal cells with knockeddown CAP amass large arrays of stable F-actin and losedynamic F-actin arrays associated with lamellipodia (Baum etal., 2000; Benlali et al., 2000; Bertling et al., 2004; Rogers etal., 2003). These re-arrangements can be interpreted asevidence supporting biochemical observations that some CAPisoforms can sequester actin monomers in vitro and thusprevent excessive polymerisation (Freeman et al., 1995;Gieselmann and Mann, 1992; Gottwald et al., 1996). Plant F-actin does not homogenously accumulate in the absence ofCAP as cap1 root hairs appear devoid of the organised bundlesthat amass in the shank of wild-type root hairs behind thegrowing tip. These observations suggest a more complex role

Fig. 8. Wild-type inflorescences (A) remain relatively straight duringbolting. Inflorescences from cap1 mutants exhibit curls and kinks atnodes (B). Some young inflorescences perform almost a completerotation during early expansion. Pedicles supporting flowers orgrowing siliques are also curled. The root systems of wild-typeplants (C) grown on the surface of solidified agar medium extendradially. Root systems of cap1 plants (D) fail to efficiently colonisethe agar surface. Comparison of an individual wild-type root andassociated lateral roots (E) with a cap1 root and its associated laterals(F) shows that cap1 roots are excessively curled and looped. Bars,2 mm.

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for CAP1 in the turnover of actin filaments, possibly relatingto the observed in vitro activity of accelerating actin monomernucleotide exchange either directly (Moriyama and Yahara,2002; Chaudhry et al., 2007) or through interplay with ADF(Mattila et al., 2004; Chaudhry et al., 2007).

The critical state of actin dynamics depends upon thebalance of actin-binding protein activity. Overexpression ofboth plant CAP1 and AIP1 (Barrero et al., 2002; Ketelaar etal., 2007) mimic the phenotypic effects of reduced expressionof these respective proteins (this study) (Ketelaar et al.,2004a). A sub-nominal level of CAP1 protein is likely toreduce the in vivo concentration of ATP-actin monomers,while an increase in CAP1 concentration or activity couldenhance CAP1 sequestration of actin monomers from the G-actin pool. Any imbalance in ABP activity impacts upon theefficacy of actin turnover and, consequently, on cytoskeletal-driven cell growth.

Arabidopsis CAP1 is required for signallingThe curling of cap1 inflorescences and roots indicates a rolefor CAP1 in coordinating the expansion of tissues. The curlingof cap1 organs is distinct from the twisting observed in mutantssuch as spiral or lefty, which show defects in microtubuleorganisation as the cap1 curls have no consistent chirality, andthe turns of roots are not associated with visible twisting ofepidermal cell files. However, the inflorescence curlingphenomenon has a common pattern of behaviour: cap1inflorescences always initiate a curl by turning towards thegravity vector. Recovery to a vertical position closes the curl,and the process is then rapidly reversed over the course of afew hours to re-straighten the inflorescence. Such movementrequires the simultaneous differential expansion of many cellfiles, suggesting the involvement of an intercellular signal thatis either misinterpreted or misdirected in the absence of CAP1.

The interpretation of mutant phenotypes in understandingthe signalling role of CAP is complicated by the multipleconsequences of cytoskeletal disturbance. Clonal analysis ofDrosophila eye discs shows a requirement for CAP insignalling processes to organise photoreceptor differentiation(Benlali et al., 2000). Rather than directly transducing a signalin the manner of SRV2, Drosophila CAP was hypothesised toperturb hedgehog signalling by causing morphologicalabnormalities across the surface of the eye disc leading to thephysical disruption of morphogen distribution. Thereforedissection of the plant CAP signalling phenotype shouldalways be considered in light of the effects of actin disruptionon morphogenesis and other basal cell processes such asexocytosis and endocytosis. Attempting to separate signallingand cytoskeletal activities through domain analysis must alsobe approached with caution. In S. cerevisiae, where thisanalysis was first performed, the ‘cytoskeletal’ phenotypeswere never fully complemented using only the C-terminus(Gerst et al., 1991) and recently the transduction of signalsfrom Ras to adenylate cyclase within a cell-death pathway wasfound to be more reliant upon the C-terminus of SRV2 than theN-terminus (Gourlay and Ayscough, 2006).

The two biological activities of CAP have long beenconsidered independent functions, but evidence isaccumulating that interaction with actin monomers could becoupled with participation in signalling pathways. InDictyostelium, CAP is required to perpetuate the cAMP

chemotactic signal (Noegel et al., 2004) and to respond to thesame signal by stimulating cytoskeletal based motility (Noegelet al., 1999). Recently, SRV2 of yeast was shown to have aremarkable association with the cytoskeleton during Ras-mediated signalling to apoptotic-like pathways (Gourlay andAyscough, 2006). Suppression of actin dynamics leads to Rasactivation, which in turn activates adenylate cyclase. One of theconsequences of this pathway is further actin rearrangements,possibly via PKA effectors downstream from cAMP signalling(Gourlay and Ayscough, 2006). CAP is needed to transduce thesignal from Ras to adenylate cyclase, and the actin-bindingdomain of CAP is required for this process. In this instanceCAP appears to offer an input to the pathway dependent uponthe actin-binding domain, even feasibly acting as some formof sensor to the free G-actin pool. As the effectors of thesignalling pathway are very likely to include other actin-binding proteins, the cytoskeletal phenotypic effects of CAPknockouts in yeast are dependent on CAP signalling activities,totally integrating the two roles and confusing any phenotypicdistinction.

The concept of integration invites a model for CAPfunction based upon feedback from the actin cytoskeleton.Latrunculin B treatment is known to exaggerate thegravitropically stimulated bending of roots (Hou et al., 2004)through an uncharacterised mechanism. The roots of cap1plants also respond in an exaggerated fashion togravistimulation (P.J.H., unpublished). The meanderingbehaviour of cap1 roots and the curling of inflorescencesmight result from an overcompensation to internal cues aimedat maintaining a controlled angle of tissue expansion. CAP1could feasibly be required to monitor and respond to thestatus of the G-actin pool in expanding cells. Rapid dynamicscould provoke the perpetuation of an intercellularcompensatory signal via CAP1 to expand other cell files inan antagonistic manner and stimulate direct intracellularsuppression of actin turnover.

ConclusionIn conclusion, Arabidopsis CAP1 is essential for healthygrowth, but unlike AIP1 its deactivation is not lethal to plantlife. The cap1 mutants could reveal aspects of the in vivobehaviour of other ABPs, both through the observation of F-actin formation in affected tissues and through double-mutantanalysis. The accumulation of F-actin at the cortex of affectedcells suggests the presence of so far unidentified F-actinmachinery at sites of intense exocytosis. Arabidopsis cap1 isalso required for the coordination of tissue expansion in themanner of a component of an intercellular signalling pathway.

Materials and MethodsRecombinant protein purificationFor actin-binding studies, the full-length transcript of CAP1 was cloned into aGateway derivative of pGEX-4T-1 to form a translational fusion with GST at theN-terminus of CAP1. Recombinant protein was expressed in E. coli strain BL21DE3 pLysS Rosetta 2. Cultures were grown to OD600 0.8 and induced using IPTG(final concentration 1 mM) at 37°C for 2.5 hours or overnight at 14°C. Cellsexpressing GST-tagged proteins were resuspended in PBS, pH 7.3 and lysed usinga freeze-thaw cycle. Cell supernatant was incubated with glutathione sepharose 4Baccording to the manufacturer’s instructions (Amersham, UK) and washed threetimes with PBS.

Actin purificationRabbit skeletal muscle actin was purified as described previously (Spudich andWatt, 1971), and as later modified (Winder et al., 1995). Briefly, rabbit muscle

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acetone powder was mixed with buffer G (2 mM Tris pH 8.0, 0.2 mM ATP, 0.5 mMDTT, 0.2 mM CaCl2, 1 mM NaN3). After a 30 minute incubation and spin, thesupernatant was filtered. MgCl2 was added to a final concentration of 2 mM andKCl was added to 0.8 M. After polymerisation the F-actin was pelleted for 2 hoursat 50,000 g. Actin was resuspended in G-buffer and dialysed for 2 days, thencentrifuged at 50,000 g. The top two-thirds of the supernatant was gel-filtered usingsephacryl S300 to isolate actin monomers.

ADP-actin was generated from purified ATP-actin monomers on the day of useby incubation with yeast 20 U/ml hexokinase and 1 mM glucose in G-buffer for 3hours (Pollard, 1986). 0.1 M ADP (Sigma) stock solution was also treated withhexokinase and glucose to remove ATP contaminants.

Actin-binding assaysFor actin-depletion assays, G-actin was added to 225 �l of depletion buffer (10 mMTris pH 7.5, 0.2 mM CaCl2, 0.5 mM DTT, 0.2 mM ADP, 1 mM MgCl2, 100 mMKCl) to make a final solution of 3 �M. 25 �l (12.5 �l bead volume) of glutathionesepharose beads (Amersham) coated with either GST or GST-CAP1 wereimmediately added to the actin solution to make a 9 �M bait protein suspension.Beads were incubated with the actin for either 5 minutes or 30 seconds with gentleagitation. Following incubation, beads were briefly spun to the bottom of the tubeand 100 �l of supernatant removed and mixed with 2� SDS loading buffer. Sampleswere run on an 8% polyacrylamide SDS gel and stained using Coomassie solution.Native gels were polymerised at a final concentration of 10% Protogelacrylamide/bisacrylamide mix (National Diagnostics). 0.2 mM ADP (pre-treatedwith 20 U/ml hexokinase and 1 mM glucose) was present in both gel and runningbuffer (25 mM Tris, 200 mM glycine, 0.5 mM DTT). Recombinant GST-CAP1 andGST were removed from glutathione beads using elution buffer (10 mM reducedglutathione, 50 mM Tris-HCl, pH 8.0) and dialysed for 5 hours with 4 changes ofG-buffer. Combinations of ADP-actin (final concentration 1 �M), GST-CAP1 (5�M) and GST (5 �M) were assembled in G-buffer with a total volume of 20 �l andincubated on ice for five minutes before loading onto the native gel.

Plant linesArabidopsis seed was sterilised using 5% bleach (BDH) for 25 minutes with gentleagitation followed by 4 washes with water. Seeds were plated on to half-strengthMurashige and Skoog salts (Sigma) with 0.8% plant agar. After germination allplants were grown either on compost or half MS plates in 16 hours light (at 22°C)and 8 hours dark (at 18°C). Salk T-DNA lines (Alonso et al., 2003) were createdby SIGNAL (the Salk Institute Genomic Analysis Laboratory) and supplied byNASC (Nottingham, UK). GABI KAT line 453G08 (Rosso et al., 2003) was createdand supplied by the Max Planck Institute for Plant Breeding Research (Cologne,Germany).

Pollen assaysWild-type and cap1 pollen was germinated in vitro as previously described(Krishnakumar and Oppenheimer, 1999). 100 �l aliquots of agarose pollengermination medium [1 mM CaCl, 1 mM Ca(NO3)2, 1 mM MgCl2, 0.01% boricacid, 18% sucrose, 0.5% agarose, pH 6.0] were solidified on microscope slides toproduce a smooth coating. A 5 �l drop of liquid germination medium (withoutagarose) was applied to the centre of the slide, and mature wild-type or cap1 pollenwas released into the liquid from open pollen sacs. Two mature pollinated stigmaswere placed on the slide within 5 mm of the samples. Samples were left either for5 hours or overnight in a closed humid environment within standard growth roomconditions. After application of coverslips, pollen tubes were observed using a ZeissAxioskop microscope with 40� objective, and images were captured using a videocamera (Roper Scientific) controlled by Openlab 3 software (Improvision, UK).

For in vivo growth assays, wild-type and cap1 pollen was used to fertilise WTstigmas. After a specific period of germination within standard growth roomconditions, stigmas were prepared as described (Jiang et al., 2005) with minormodifications. Fertilised carpels were dissected longitudinally to bisect the septum.The dissected tissue was fixed in a 3:1 ethanol:acetic acid solution for 2 hours. Thesamples were then washed with water and incubated with 10 M NaOH for 2 hours.Tissue was subsequently washed 3 times with water and 3 times with 100 mMK2HPO4-KOH, pH 11. Samples were incubated in the dark for 2 hours in 0.1%aniline blue in 100 mM K2HPO4-KOH, pH 11 before being mounted in glyceroland observed with a Zeiss LSM510 confocal microscope under 405 nm blue diodelaser excitation.

ImagingInflorescence epidermal peels were imaged using an Eclipse TE300 invertedmicroscope (Nikon, Japan) with Orca camera (Hamamatsu, Japan). All imagingof GFP:FABD2 fluorescence was performed using a Zeiss LSM510 confocalmicroscope with 40� objective. For mitochondrial imaging, growing root hairswere labelled with 250 nM mitotracker red CMXros (Invitrogen). For microscopicanalysis of growing root hairs, seedlings were grown in ‘biofoil sandwiches’, asdescribed previously (Ketelaar et al., 2004a). Images of various organs weretaken using a SZH10 stereomicroscope (Olympus) and video camera (RoperScientific).

GenotypingDNA from plants was prepared as described by (Edwards et al., 1991). All PCRexperiments used RedTaq (Bioline) polymerase in accordance with manufacturer’sinstructions. The CAP1 wild-type allele was amplified using primers:CAP1-1 For: 5�-CTCCTGACTTCGCCATC-3�CAP1-1 Rev: 5�-GCAACCTGAACAAGTAACTATA-3�CAP1-2 For: 5�-CTCGTACCAGTAAACCGGCCTT-3�CAP1-2 Rev: 5�-CCGTGAAAACAACACCCACTTT-3�.

The amplification of the T-DNA insertion alleles was achieved with primers:CAP1-1 Rev and LBb1 (5�-GCGTGGACCGCTTGCTGCAACT-3�)CAP1-2 Rev and cap1-2TDNA (5�-CCCATTTGGACGTGAATGTAGA-3�).

RT-PCRTotal RNA was isolated from homozygous mutant and azygous plants using theRNeasy Plant Mini Kit (Qiagen). The total RNA was DNase treated with RNase-free DNase (Promega) following the manufacturer’s instructions. The cDNA first-strand was synthesised using 5 pmol of oligo(dT) 12-18mer and 200 units ofSuperscriptTMII RNaseH-Reverse Transcriptase (Invitrogen), according to themanufacturer’s instructions. The RNA was removed by addition of 2 units ofRibonuclease H (Promega) and incubation at 37°C for 20 minutes. 1 �l of thereaction mixture was used as a template for the PCR reaction with BioRed Taq DNApolymerase (Bioline).CAP28F: 5�-GGAATCCATATGGAAGAGGATTTGATTAAGCGCCTT-3�CAP28R: 5�-GGAATCCATATGTTAGGCACCTGAATGCGAGACCGGTGTTG-3�.

Control primers to Arabidopsis glyceraldehyde-3-phosphate dehydrogenase Cwere:GAPCFOR: 5�-CACTTGAAGGGTGGTGCCAAG-3�GAPCREV: 5�-CCTGTTGTCGCCAACGAAGTC-3�.

Environmental scanning electron microscope (ESEM)preparationThe plant material (leaves from homozygous mutant and azygous plants, from eachT-DNA line) was prepared for ESEM by fixation (PBS, 1% glutaldehyde, 0.1% (v/v)Tween) and dehydration by an ethanol series, followed by critical point drying withcarbon dioxide. Samples were imaged using a Philips XL30 ESEM in low vacuummode (0.4 Torr).

This work was supported by the Biotechnology and BiologicalSciences Research Council (M.J.D., S.D., T.K., P.J.H.) and the FCTPortugal, SFRH/BD/8760/2002 (C.R., R.M.). We thank Steve Winderfor purified actin, and Christopher Staiger and Laurent Blanchoin forpersonal communications concerning the biochemistry of ArabidopsisCAP1.

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