my body is a cage: mechanisms and modulation of plant cell growth

Tansley review

Mybody is a cage:mechanisms andmodulationof plant cell growth

Author for correspondence:Keiko Sugimoto

Tel: +81 45 503 9575Email: [email protected]

Received: 26 May 2013

Accepted: 1 August 2013

Luke Braidwood, Christian Breuer and Keiko Sugimoto

RIKEN Center for Sustainable Resource Science, 1-7-22 Suehiro, Tsurumi, Yokohama, Kanagawa 230-0045, Japan

Contents

Summary 1

I. Introduction 1

II. Modes of growth 2

III. Biomechanics and modelling of plant cell growth 2

IV. Plant cell wall structure 3

V. Cell wall modification proteins 4

VI. Direction and duration of cell growth 6

VII. Regulation of plant cell growth 7

VIII. Conclusions 11

Acknowledgements 12

References 12

New Phytologist (2013)doi: 10.1111/nph.12473

Keywords: Arabidopsis, cell growth, cell wall,diffuse growth, plant cell, signal integration,tip growth, wall structure.

Summary

Thewall surrounding plant cells provides protection fromabiotic and biotic stresses, and support

through the action of turgor pressure. However, the presence of this strong elastic wall also

prevents cell movement and resists cell growth. This growth can be likened to extending a house

from the inside, using extremely high pressures to push out the walls. Plants must increase cell

volume inorder to explore their environment, acquire nutrients and reproduce. Cellwallmaterial

must stretch and flow in a controlledmanner and, concomitantly, new cell wall material must be

deposited at the correct rate and site to prevent wall and cell rupture. In this review,we examine

biomechanics, cell wall structure and growth regulatory networks to provide a ‘big picture’ of

plant cell growth.

I. Introduction

Plants use plastic growth to construct cells that vary enormously insize and shape (Fig. 1). Cell volumes can vary 10 000-fold within aspecies, increasing in size from meristemoids to xylem vessels(Cosgrove, 2005). Many plant cells are cylindrical or tubular, butcells may also be spherical, for example epidermal bladder cells inMesembryanthemum crystallinum, or even stellate, as in theaerenchyma of Cyperus javanicus. Complex plant cell shapes areproduced by tight regulation of growth, as in the interdigitation ofArabidopsis thaliana leaf epidermal cells, andmay bemaintained bystructural reinforcement after the cessation of growth, as in xylemvessels. How plant cells are shaped has been discussed elsewhere(Martin et al., 2001) and, in this review, we focus on the

mechanisms of cell growth and how these mechanisms areregulated. Understanding growth at the cellular level is vitalbecause plant shape is dictated by two factors: cell number and cellsize. Plant development includes numerous indeterminate pro-cesses, and responds drastically to environmental stimuli in order tomaximize reproductive potential within the local environment. Forexample,Cryptomeria japonica can be both a diminutive bonsai andthe ancient Jomon-sugi, which has a volume of c. 300 m3. Plant cellgrowth is produced and controlled at numerous different levels,which we will highlight in turn. Ultimately, plant cell growth is abiomechanical process governed by physical laws, which mathe-matics and modelling attempt to describe. The forces and materialproperties present in this process are determined by the osmolalityof plant cells and the composition of their walls. Furthermore, these

� 2013 The Authors

New Phytologist� 2013 New Phytologist TrustNew Phytologist (2013) 1

www.newphytologist.com

Review

structural and biochemical features of plant cells are controlled bycomplex regulatory networks, which integrate diverse develop-mental and environmental inputs. A full understanding of plant cellgrowth requires a broad perspective of these different hierarchicallevels, and the scientific fields that describe them.

II. Modes of growth

Plant cell growth is predominantly anisotropic; the rate anddirection of growth vary across the cell. This allows the plant toform cell shapes other than spheres, which are produced byisotropic growth. Anisotropic growth includes diffuse and tipgrowth. Plant cells grow mainly by diffuse growth, in which

expansion is dispersed over the cell surface, for example root cortexcells elongating in the longitudinal direction as they leave the rootmeristem. Some specialized cells produce structures by tip growth,such as root hairs and pollen tubes, in which expansion occurs at asingle site. Diffuse growth is unique to plant cells, whereas someanimal and fungal cells also undergo tip growth. There arenumerous important differences between these growth forms.Wallmaterial is deposited perpendicular to the direction of maximalgrowth in diffuse growth, whereas deposition is parallel withmaximal growth in tip growing cells. Diffuse growth is equivalentto stretching a wall lengthwise whilst reinforcing it to preventrupture; tip growth is like trying to burrow through the wallwithout breaking its surface. In addition to this mechanicaldistinction, the cytological properties of each mode of growthdiffer.

III. Biomechanics and modelling of plant cell growth

Plant cell growth occurs through the controlled expansion of thecell wall, which results from the interplay between turgor pressureand cell wall elasticity and extensibility (Ray et al., 1972). Turgorpressure is produced by the influx of water from the extracellularspace, as a result of the lower osmotic potential of the cytoplasm.Turgor pressures in the range 0.3–1.2 MPa (similar to that within acar tyre) have been measured in A. thaliana cells (Forouzesh et al.,2013).Within a turgid cell at equilibrium, this outwards pressure iscounterbalanced by stress within the cell wall, and produces a waterpotential difference of zero. Wall stress acts to oppose changes inwall shape, and therefore increases in cell volume. The negligiblecompressibility of water at physiologically relevant pressuresensures that any increase in cell volume (i.e. growth) requireslower water potential within the cell than outside it. The Lockhartequation (Lockhart, 1965) was the first to describe diffuse growthby linking turgor pressure andwall extensibility to volume increase:

dV

dt¼ UðP � Y Þ

(dVdt , rate of volume increase; Φ, irreversible wall extensibility; P,pressure; Y, yield threshold).

This models the cell wall as a viscoelastic substance thatundergoes plastic deformation when turgor pressure exceeds theyield threshold. The yield threshold is the point above whichpressure produces growth, and irreversible wall extensibilitygoverns how much growth this excess (i.e. above yield threshold)pressure produces. The yield threshold describes howmuch energythe wall components can store elastically before being permanentlydeformed or moved relative to each other. Irreversible wallextensibility describes the stiffness of the cell wall. Both of thesevalues are determined by the composition of the cell wall and theinteractions between the cell wall components, although theserelationships are complex and not currently well understood.

If a turgid cell is at equilibrium (dVdt = 0, P > 0), negative waterpotential must be generated for growth to begin. This can occur,mathematically, by either increasing the concentration of soluteswithin the cell or reducing stress within the wall and thereby

Fig. 1 Cell-type-specificdifferentiationprogrammes lead toawidevarietyofcell sizes and cell shapes in Arabidopsis thaliana. Representation of fivedifferent types of epidermal cell of the root, the hypocotyl, the rosette leafand the flower. Cotyledons are not shown. Note: the root hair image derivesfromanoptical confocal cross-sectionof theprimary root andalsodepicts thesubepidermal cell layers of cortex and endodermis. All other images areaccording to scanning electron micrographs (SEM). Stomatal guard cells ofthe adaxial leaf epidermis are highlighted in green. Bars, 25 lm.

New Phytologist (2013) � 2013 The Authors

New Phytologist� 2013 New Phytologist Trustwww.newphytologist.com

Review Tansley reviewNewPhytologist2

decreasing turgor pressure (Ray et al., 1972). The Lockhartequation has been modified repeatedly, for example to includefactors such as elastic wall extensibility and transpiration rate(reviewed in Geitmann & Ortega, 2009). Recent approaches havealso modelled Φ as variable through space and time (Pietruszka,2011). Although the Lockhart equation is simplistic, it stillencapsulates the key concepts thought to govern cell growth. Itremains a point of debate whether osmolality or wall stress is alteredfirst, to remove a cell from equilibrium and begin its growth. Putsimply, is plant cell growth ‘wall first’ or ‘water first’? The classicLockhart model has been challenged by loss-of-stability (LOS)theory (Wei & Lintilhac, 2003). LOS theory attributes growth tostructural failure of the wall as a result of increasing turgor ratherthan activemodification of wall properties (Wei&Lintilhac, 2007;Schopfer, 2008; for an opposing view). Aweakness of LOS theory isthat it cannot predict wall behaviour after the initiation of growth(Geitmann & Ortega, 2009).

The principles behind the modelling of diffuse growth have alsobeen adapted to try and recreate the dynamics of tip growth, withalterations to account for the unique geometric and biologicalfeatures of tip growth. Recent models include the hydrodynamicmodel and a calcium-cell-wall model. The hydrodynamic modelsuggests that water flux within pollen tubes can produce pressuregradients, and that this increased pressure at the tip drives growth(Zonia & Munnik, 2011). This model has been challenged onphysical grounds, and because a pressure gradient is yet to berecorded within plant cells (Winship et al., 2011). An alternativecalcium-cell-wall model has also been proposed, and is based onstretch-activated calcium channels in the pollen protoplast. As thecell wall thins with growth, wall stress increases (a general physicalprinciple). Above a certain level of stress, this opens the calciumchannels, stimulating vesicle fusion and wall material deposition atthe tip (Kroeger et al., 2011). The evidence for models (of bothdiffuse and tip growth) that rely on wall modification to initiategrowth is currently stronger than that for ‘water first’ models, suchas LOS, and the hydrodynamic model. It remains unknownwhether cells ever actively increase turgor pressure in order to grow,and ‘… in not a single example of cell growth has an active increasein turgor pressure been documented’ (Winship et al., 2011;Kierzkowski et al., 2012; Kutschera & Niklas, 2013).

Regardless of whether modification of wall properties or turgorpressure initiates growth, it is clear that wall modification isrequired to direct growth within a system governed by hydrostaticpressure.Turgor pressure is a scalar physical force and, although it ispossible to exert active control over its magnitude, through theactive transport of ions and solutes, its direction cannot becontrolled. Consequently, turgor pressure would produce a sphereif the cell wall had a homogeneous capacity for expansion. Plantscells must therefore spatially regulate construction and/or alter-ation of their walls to produce non-spherical cells. Counter-intuitively, although plant cells require turgor pressure for growth,in the widespread phenomenon of anisotropic growth, plant cellsare working actively against the largest pressure forces to producegrowth that takes the cell shape further away from spherical. In agrowing cylindrical cell, the hoop stress within thewall is double thelongitudinal axial stress, but most expansion occurs in the

longitudinal direction. Stress patterns formore complex cell shapesdo not seem to have been formally investigated. Unless notablehydrodynamic pressure can be produced by the directional flow ofwater through plant cells, it is universal that turgor pressure is arequisite for growth, but when and where it occurs is controlled bywall modification.

IV. Plant cell wall structure

We briefly summarize the structure of the plant cell wall beforediscussing how its modification and composition can controlgrowth. The plant cell wall is a complex and heterogeneous matrixof polysaccharides, glycoproteins, solutes and enzymes, whichvaries in composition within and between individual cells, tissuesand species. We focus on the major wall polysaccharides, but thereare numerous other wall components, such as glycoproteins andphenolics. Many excellent and recent reviews on the plant cell wallcover general structure (Cosgrove, 2005; Cosgrove& Jarvis, 2012)and specific components (Bunzel, 2010; Lamport et al., 2011).Primary cell walls are thin, lack lignification and are able to undergogrowth, so are referred to in this review.

1. Cellulose

Cellulose, linear b-1,4-glucan, is the key structural component ofcell walls. Crystalline cellulose microfibrils are inelastic andproduced by multimeric complexes of cellulose synthases (CESAs)at the plasma membrane, which contain up to 36 CESA proteins.CESA proteins catalyse the extension of b-1,4-glucan polymerchains, which then spontaneously cocrystallize into microfibrilswith other chains produced by the same complex (Somerville,2006). In A. thaliana, CESA complexes require at least threedifferent CESA subunits. The primary cell wall is produced byCESA1, CESA3 and, typically, CESA6 (Persson et al., 2007).

2. Pectins

The remaining matrix polysaccharides are synthesized in Golgistacks and released into the wall by exocytosis. Pectins are anextremely diverse group of complex high-molecular-weight poly-saccharides that are soluble in aqueous and acidic conditions.Structural groups of pectins include homogalacturonan (HG),rhamnogalacturonan I and II, and xylogalacturonan. Thesecomponents form complex hydrated gels, cross-linked by covalentand ionic bonds between different pectins and other wallpolysaccharides (Marcus et al., 2008; Caffall & Mohnen, 2009).Pectins contribute strength and flexibility to the wall, separatemicrofibrils and link adjacent cells in themiddle lamellae (Caffall&Mohnen, 2009).

3. Hemicelluloses

Hemicelluloses are typically defined as polysaccharides that are notcellulose or pectins, which primarily increase wall strength by cross-linking cellulose microfibrils (reviewed in Scheller and Ulvskov,2010). Xylans, xyloglucans (XGs), glucomannans and mannans


New Phytologist� 2013 New Phytologist TrustNew Phytologist (2013)


NewPhytologist Tansley review Review 3

are hemicelluloses thought to be present in all land plants. XG isthe primary hemicellulose in dicotyledonous plant cell walls, andinteracts with pectins and cellulose microfibrils, with a smallproportion entrappedwithinmicrofibrils (Dick-P�erez et al., 2011).

The structure of the cell wall has been conceptualized previouslyas a ‘tethered network’ of cellulose cross-linked by XG, which ispresent within a relatively separate pectinmatrix (Cosgrove, 2005).However, recent three-dimensional solid state nuclear magneticresonance (NMR) of A. thaliana cell walls suggests that pectin is anintrinsic part of the load-bearing network of the wall. Pectins werefound to interact extensively with both XG and cellulose, whereasXG–cellulose interactions were less common than previouslythought (Dick-P�erez et al., 2011). In addition, the use of variousb-1,4-endo-glucanases demonstrated that XG digestion alone wasnot sufficient to produce polymer creep (Park&Cosgrove, 2012b),which would be expected if the tethered network model representsthe load-bearing structure of the cell wall. The new cell wallstructure suggested by these data is that of a single complexpolysaccharide network featuring interactions between each ofcellulose, XG and pectin (Fig. 2; Dick-P�erez et al., 2011). Thesedata leave open the question of whether microfibrils are linkedtogether by XG, pectin or both, whether alone or in combination(all possibilities shown in Fig. 2), and further work should aim todetermine the network topography.

V. Cell wall modification proteins

Plants can control wall structure both through alteration of the wallmaterials deposited outside the plasma membrane and by dynamicmodification of the composition and interaction of wall compo-nents after their deposition. Post-depositional modification of cellwall polysaccharides allows plant cells to alter the physicalproperties of their walls in response to novel stimuli. This processis conceptually similar to the post-translational modification ofproteins, which allows rapid modification of the structure,interactions and stability of proteins already present as a result of

translation. Plant cells release numerous classes of proteins, such asglycosyltransferases, that alter cell wall properties and cell growththrough the modification of different wall components (summa-rized in Table 1). We focus on two well-studied examples directlylinked to cell growth: expansins, the classic example of a wall-modifying protein, and pectinmethylesterases (PMEs), which havebeen shown recently to play an important role in the growth ofpollen tubes, shoot apical meristem (SAM) primordia andhypocotyls (Tian et al., 2006; Pelletier et al., 2010; Peaucelleet al., 2011).

1. Expansins

Acid growth theory (Rayle, 1973) states that the influx of hydrogenions to the wall causes loosening, allowing increased expansionthrough polymer creep. Expansins, the proteins generally acceptedto be responsible for acid growth, were discovered in the early1990s. Heat-inactivated walls had pH-dependent expansionrescued by the addition of proteins extracted from cucumberseedlings, which could also induce extension in other species(McQueen-Mason et al., 1992). Another family of expansins waslater identified as a group of grass pollen allergens that loosen cellwalls (Cosgrove et al., 1997). These two families of expansins,named a-expansin or EXPA and b-expansin or EXPB, respectively,are involved in a wide variety of cell wall modification processes,including fruit ripening, abscission, and the penetration of the styleby pollen (Cosgrove, 2005). Expansins are present throughoutland plant phylogeny, their expression corresponds with growth(Cosgrove, 2000) and expansin antisense treatment reduces cell sizein vivo and cell wall extensibility in vitro (Cho & Cosgrove, 2002;Goh et al., 2012).

Expansins are small extracellular proteins with no apparent lyticactivity that enhance polymer creep, and thereby wall extensibility,by disrupting non-covalent interactions between wall polysaccha-rides, in particular between cellulose microfibrils. Expansinsweaken paper, which is a network of microfibrils with few other

Fig. 2 Primary cell wall structure andbiomechanics of cell growth. Cell wallstructure as hypothesized by Dick-P�erez et al.(2011), with interactions between each ofpectin, cellulose and xyloglucan (XG). A smallfraction of XG is entrappedwithinmicrofibrils.Hydrostatic pressure is equal betweenendomembrane compartments, and stressfrom the wall balances turgor, except in thedirection of growth (Szymanski & Cosgrove,2009). Axial stress is half that of hoop stress,but growth occurs in the axial direction as aresult of anisotropic wall construction andmodification. CESA, cellulose synthase.




polysaccharides, and enhance cellulase activity, which is limited bythe accessibility of glucan (Cosgrove, 2005). Targeted mutagenesisusing bacterial EXPANSIN-LIKE X1 (EXLX1) showed thatmutations that decreased hydrophobic cellulose binding reducedin vitro wall-loosening activity, whereas mutations that decreasedelectrostatic binding to pectins and hemicelluloses in fact increasedwall-loosening activity (Georgelis et al., 2011). These data raise theintriguing possibility that the modification of non-cellulosicpolysaccharides to alter their charge could modulate expansinactivity. These key residues correspond to similar residues in plantexpansins, but EXLX1 has lower wall-loosening activity and itsactivity is not pH dependent, suggesting possible differences inmechanism.

Even if the mechanism of expansin action only involvesmicrofibrillar interactions, other wall polysaccharides affectexpansin activity indirectly. For example, the A. thaliana XGsynthesis double mutant xylosyltransferase1 (xxt1)/xxt2, whichcontains no known XGs, has walls with altered biomechanicalproperties. Cell wall responses to a-expansin activity were reducedin xxt1/xxt2 mutants (Park & Cosgrove, 2012b). This decrease inactivity may be caused by XG absence if expansins act on themdirectly, or if XGs promote cellulose structures on whichexpansins act. However, pectin and arabinoxylan play a greaterstructural role in XG-deficient walls (Park & Cosgrove, 2012a),which may, in turn, be responsible for altered expansin activity. Itappears that functional divergence of the expansin superfamily hasoccurred, as the application of purified maize pollen b-expansinwas found to release glucuronoarabinoxylan and HG from the cellwalls of grasses, but not other plants, probably through a non-enzymatic mechanism (Tabuchi et al., 2011).

2. Pectin methylesterases

PMEs catalyse the removal of methyl esters from the (1?4) a-D-GalAbackbone ofHG, themost abundant cell wall pectin.HG isdeposited at the cell wall in a 70–80%methyl-esterified form. PMEaction produces a negatively charged carbon on the HG backbone,allowing the formation of calcium cross-links between chains of

unmethyl-esterified GalA residues. The egg-box model describesthis cross-linking (Liners et al., 1989): two negatively chargedpectin chains chelate positive calcium ions between them. Thismodification also increases HG susceptibility to lysis catalysed bypolygalacturonanases (PGases) and pectate lyases within the wall(Caffall & Mohnen, 2009). Calcium cross-linking increases thestiffness of cell walls, but pectin degradation increases wallextensibility. This seeming contradiction may be resolved by thepattern of demethyl-esterification. PMEs can remove methyl estersfrom the same HG backbone in a processive fashion, producing astring of negatively charged carbons, or randomly from multiplebackbones. It is thought that ‘blocks’ of negative charge are mostimportant for forming gels, whereas random methyl ester removalmay promote degradation by PGases without causing a largeincrease in wall stiffness (Pelloux et al., 2007). PME activity,appropriately, has been found to be responsible for both thepromotion and inhibition of cell wall growth.

Recent work has linked PME activity to cell growth in a numberof developmental processes. In the A. thaliana SAM, lateral organsare produced from lateral outgrowths called primordia. Subepi-dermal cells in lateral organ primordia are more elastic and growmore quickly than surrounding cells (Peaucelle et al., 2011). PMEactivity is likely to be at least partly responsible for this change inwall properties and cell growth rate. Antibody labelling showed thatmethyl-esterified HG dominated the centre of the SAM dome,whereas demethyl-esterified HG was present in incipient andemerging primordia, suggesting the involvement of PMEs inprimordium formation. Ethanol-inducible expression of theendogenous PME inhibitor PMEI3 prevented lateral organoutgrowth, and citrus PME loaded onto sepharose beads inducedprimordium initiation, showing that PME activity is necessary andsufficient for increased cell growth in the SAM during lateral organoutgrowth (Peaucelle et al., 2008). By contrast, tip growth in pollentubes uses PME activity to stiffen cell walls in the tube shank: highlymethyl-esterifiedHG is deposited at the growing tip, where thewallis softer (reviewed in Chebli et al., 2012). PME activity is restrictedby tip-localized PME inhibitors, such as AtPMEI2 (R€ockel et al.,2008).

Table 1 Cell wall-modifying enzymes

Enzyme group1 Example Targets Location References

b-Galactosidases AtBGAL10 Xyloglucans (XG) Cell wall Figueiredo et al. (2011); Sampedro et al. (2012)Expansins AtEXP7 Cellulose microfibrils, XG Cell wall Cosgrove (2000)Glycosyltransferases AtXEG113 Extensins, HRGPs2 Golgi Velasquez et al. (2011)Pectin methylesterases AtPME5 Pectins (e.g.

homogalacturonan)Cell wall Pelloux et al. (2007); Peaucelle et al. (2011)

Prolyl 4-hydroxylases AtP4H2 Extensins, HRGPs Golgi,endoplasmicreticulum

Velasquez et al. (2011)

Transglycosylases Unknown XG, hemicelluloses Cell wall Johnston et al. (2013); Mohler et al. (2013)UDP-D-Glucose 4-epimerase AtRHD1 XG, pectins Golgi R€osti et al. (2007)Xyloglucanendotransglucosylase/hydrolases

AtXTH18 XG Cell wall Van Sandt et al. (2007); Ekl€of & Brumer (2010)

1Enzymes required for the biosynthesis of wall components not known to be involved in wall modification are not included.2Hydroxyproline-rich glycoproteins.





There are a large number of putative PME isoforms in allsequenced plant genomes. For example, Populus trichocarpa has 89,A. thaliana has 66 and Oryza sativa has 35 (Pelloux et al., 2007).This diversity reflects their wide range of functions. PMEs areinvolved in seed germination, root tip elongation, lignin produc-tion (Pelloux et al., 2007) and pathogen responses (Lionetti et al.,2012). Isoforms differ in optimumpH, salt dependence and actionmechanism (Jolie et al., 2010). The charaphacean alga Characorollina utilizes a Ca+-pectatemechanism to control growth, and ithas been suggested that this mechanism is maintained in landplants, with the addition of the cellulose–expansin system tocomplement it (Boyer, 2009; Peaucelle et al., 2012). Recent studieshave shown PMEs, and other enzymes with pectin substrates, to beimportant in both starting and stopping plant cell growth(Peaucelle et al., 2012), and their ubiquity and diversity suggestthat they may provide an integral mechanism of cell growth.

VI. Direction and duration of cell growth

Cell growth must be tightly controlled to construct a complexmulticellular organism. Therefore, the control of the direction ofcell growth and the duration of this growth is essential.We examinehow diffuse growth and tip growth are guided, and discuss how thecell cycle may be involved in the control of the duration of cellgrowth.

1. Directing diffuse growth

Diffuse growth is essential for the growth of plant roots and shoots.Diffuse growth may occur over whole planes of the cell, as in thelongitudinal expansion of hypocotyl cells, or on a portion of theirsurface, as in the interdigitation of epidermal leaf pavement cells.Diffuse growth requires the deposition of wall material over a widearea of the cell surface, and the wall must be constructedanisotropically. This allows the wall to respond differently to thestress produced by turgor pressure depending on its direction,which controls the direction of growth.

The orientation of cellulose microfibril arrays is a majordeterminant of the direction of diffuse growth; they are typicallyperpendicular to the axis of maximum elongation. The alignmenthypothesis (Heath, 1974) suggested that cellulose deposition couldbe oriented by the cortical microtubule (MT) array. This wasconfirmed by observation of a fluorescent CESA6 construct(Paredez et al., 2006). CESA proteins are deposited preferentiallyat sites on the wall beside corticalMTs (Gutierrez et al., 2009), andMTs confer direction to CESA complex movement (Paredez et al.,2006). The association of complexes with individual MTs in cellstreated with the MT-destabilizing drug oryzalin (Paredez et al.,2006) suggests that complexes are tethered to individual MTs,rather than constrained passively between MT arrays, as suggestedpreviously (Giddings & Staehelin, 1991).

A crucial factor for CESA–MT association has been identifiedrecently. CELLULOSE SYNTHASE INTERACTIVE PRO-TEIN1/POM-POM2 (CSI1/POM2) is required for microfibrilsof normal length and orientation (Gu et al., 2010), binds MTsin vitro and is required for the association of CESA complexes and

MTs in vivo (Bringmann et al., 2012; Li S et al., 2012). Theguidance of CESA complexes by MTs provides a mechanism forplants to control the anisotropy of their walls. MT positioning wasfound to align with predicted principal stresses in the SAM, and torealign appropriately when stress patterns were altered throughlaser ablation (Hamant et al., 2008). This suggests that mechanicalstress at the cellular and tissue level regulates MT placement andmay therefore regulate microfibril arrangement in individual cells.

Ordered local cellulose deposition does not require an orderedcorticalMT array, suggesting the existence of a separatemechanismthat operates redundantly to produce transverse parallel arrays ofmicrofibrils. Arabidopsis thaliana root epidermal cell MTs deviatefrom a transverse orientation roughly coincident with a deceler-ation in growth rate, whereas microfibril orientation remainstransverse until long after cells cease elongation (Sugimoto et al.,2000). Furthermore, disruption of transverse cortical MTs using atemperature-sensitive mutant or oryzalin does not alter microfibrilorientation significantly (Sugimoto et al., 2003). These resultscould be explained if CESA complex movement can be guided bythe microfibril structure of the wall interior, that is, if CESAmovement parallel to the microfibril array is energetically favoured(Baskin, 2001). However, this kind of mechanism was challengedwith the observation that microfibril deposition can return to aparallel transverse array, without MTs, even after cell wallmicrofibril orientation is randomized (Himmelspach et al.,2003). The status of pectin within the wall may influence cellulosedeposition. Cobtorin disrupts the deposition of transverse cellulosemicrofibrils, but leaves parallel cortical MT arrays intact (Yonedaet al., 2007). Overexpression of a PME or a PGase can supress thisdisruption and return cellulose deposition to a direction parallel tocortical MTs (Yoneda et al., 2010).

2. Directing tip growth

Tip growth is a specialized form of growth that is spatially regulatedin the extreme. This growth form therefore provides a useful systemfor understanding how plant cells convert a scalar force, pressure,into growth, a vector. Land plants construct root hairs, pollen tubesand bryophyte rhizoids using tip growth. There are numerousdifferences between tip growth in different cell types andphylogenetic groups, but general features are as follows: an apicalregion enriched in vesicles; enrichment of Golgi stacks andmitochondria, for secretion; longitudinal arrangement of mictotu-bules and actin bundles along the tube shank, and cytoplasmicstreaming (Rounds & Bezanilla, 2013).

The first stage of tip growth is the specification of the growth site.In A. thaliana root hair cells, there is a local auxin gradient with thehighest concentration at the basal end, where the root hair forms;abolishing this gradient leads to hair formation at the apical or basalend of root hair cells (Fischer et al., 2006). The establishment ofpolarity is followedby the actionof aRhoGTPaseGDPdissociationinhibitor, which restricts ROOT HAIR DEFECTIVE2 (RHD2)to the growth site (Carol et al., 2005; Takeda et al., 2008). RHD2produces reactive oxygen species (ROS), which trigger Ca2+ influx,which, in turn, stimulate RHD2 to increase ROS production(Takeda et al., 2008). This positive feedbackmaymaintain polarity




in the growing tip. ROS production may also be involved in pollentube initiation (Speranza et al., 2012). The yield threshold of thewall at the growth site must be lowered to allow growth at this siterather than all over the cell. Wall pH drops at the site of hairinitiation, andblocking thepHdropwithbuffer reversiblyhalts hairinitiation and hair growth (Bibikova et al., 1998). This pH changemay increase expansin activity at the growth site, and two expansinsare expressed specifically in root hair cells immediately before theonset of root hair growth (Cho & Cosgrove, 2002).

Once a growing site has been specified and polarity established,cell wall material must be deposited at the site of growth. It is likelythat the growing tip of cells contains one or more specializedmembrane domains that regulate the targeted deposition ofvesicles, as both sterol and phosphoinositide signalling are requiredfor normal root hair initiation and growth (Ove�cka et al., 2010;Boss & Im, 2012). In pollen tubes, the direction of the growing tiplargely depends on a highly dynamic network of F-actin mediatingexo- and endocytosis for the deposition of plasma membrane andcell wall components and for the recycling of other components(Lee & Yang, 2008). The rate of deposition must be regulated toprevent wall rupture. As mentioned above, a recent modelpostulates that the stretch-activated calcium channels at pollentube tips regulate wall material deposition.

3. Duration of growth and endocycling

In numerous plant tissues, a rapid cell volume increase as a result ofdiffuse and tip growth is correlated with endocycling, also known asendoreduplication or endoreplication (Breuer et al., 2010; DeVeylder et al., 2011). Endocycling is DNA replication in theabsence of cytokinesis, leading to an increase in cell ploidy.Although various studies have revealed a positive correlationbetween cell size and ploidy level, such as in the A. thaliana leafepidermis (Melaragno et al., 1993), this concept is challenged byother plant studies, strongly suggesting the existence of ploidy-independent growth mechnisms (Beemster et al., 2002; Schnittgeret al., 2003). The trihelix protein GTL1 was found to repress cellgrowth during a late stage of trichome growth, through transcrip-tional repression of CELL CYCLE SWITCH PROTEIN 52 A1(CCS52A1), an anaphase-promoting complex/cyclosome (APC/C) activator (Breuer et al., 2012). This suggests that the inhibitionof endocycle progression is involved in halting cell growth intrichomes. Interestingly, the same study shows genetic evidencethat cell growth regulation by GTL1 also involves a ploidy-independent pathway, but the molecular mechanism still requiresfurther elucidation. In addition, a cell cycle regulator that promotesendocycling, KIP-RELATED PROTEIN5 (KRP5), has beenlinked to wall modification (J�egu et al., 2013). Further workshould aim to demonstrate more functional links betweenendocycle progression and cell growth.

VII. Regulation of plant cell growth

The astonishing ability of plants to shape themselves to theirenvironment requires complex regulatory systems that integrateendogenous and exogenous factors dynamically to control cell

growth appropriately. This sensitive regulation is necessary becauseplants are trapped within their local environment. Plant develop-ment is so plastic that it may be useful to view it as the response of acomplex network to endogenous and exogenous inputs of compa-rable importance, rather than the result of environmental pertur-bations to an underlying developmental programme. Recentgenetic studies have begun to uncover some of the regulatorsinvolved in these upstream signalling cascades, but how theyactually modify cell growth is far from clear. The network thatcontrols growth will probably do so by modulating genetranscription and altering enzymatic activity and protein stability.Potential targets for this network include cytoskeletal components,enzymes that synthesize and modify cell wall components, such asCESAs and PMEs, and enzymes that modify cell wall properties,such as expansins. In this section, we summarize our currentunderstanding on how diffuse growth and tip growth are regulatedby various environmental cues.

1. Regulation of diffuse growth

We examine inputs, signal integration and outputs in a growingA. thaliana hypocotyl, which serves as an excellent model system tostudy how various environmental stimuli modulate diffuse growth(Fig. 3), as hypocotyl growth is predominantly based on cellelongation, not proliferation (Gendreau et al., 1997). Understand-ing of this model system has advanced rapidly over recent years,especially in the search for signalling hubs at which diverse inputsconverge.

Inputs The shade avoidance response is a crucial feature ofhypocotyl growth, and ensures that energy is not expended oncotyledon expansion until they reach a favourable light environ-ment. Hypocotyls in favourable light conditions become de-etiolated, hypocotyl growth is inhibited and the cotyledons expand.In poor light conditions, rapid hypocotyl growth is promoted,cotyledons remain closed and an apical hook is present. Thisresponse is known as skotomorphogenesis or etiolation. Perceptionof the light environment occurs via multiple mechanisms, includ-ing phytochromes (Casal, 2013). A low red : far-red (R : FR) ratioindicates shade and produces low levels of active phytochromes, thefar-red-absorbing Pfr. Active Pfr proteins from the PhyB clade ofphytochromes are known to interact with PHYTOCHROMEINTERACTING FACTORs (PIFs), resulting in phosphorylationand degradation of PIFs (reviewed in Chen & Chory, 2011). PIFsare basic helix–loop–helix (bHLH) transcription factors thatpromote cell elongation by indirectly (stimulating auxin produc-tion) and directly modulating growth-related genes. PIFs have alarge overlap of target loci, and partial redundancy is common(Zhang et al., 2013). In high R : FR, such as in white light,phytochromes undergo a conformational change from the red-light-absorbing inactive Pr form into the active Pfr form, and repressPIF activity, resulting in decreased cell elongation and photomor-phogenesis.

The circadian clock also regulates hypocotyl growth. Proteinsinvolved in the maintenance of periodicity are known to form acomplex, involving EARLY FLOWERING3 (ELF3), ELF4 and





LUX ARRHYTHMO (LUX), which regulates the expression ofPIF4 and PIF5 in the early evening (Nusinow et al., 2011). Thehypocotyl growth rate varies through the day, peaking just beforedawn; normal variation requires both light receptors and afunctional circadian clock (Breton & Kay, 2007; Nozue et al.,2007). This system is known as the double coincidencemechanism,and is integrated with temperature and hormone signalling(Nomoto et al., 2012a,b). Metabolic status is also likely to regulategrowth as it determines howmuch growth the plant can afford andwhat type of growth will be beneficial. Recently, sucrose has beenfound to promote hypocotyl growth and auxin levels in a partiallyPIF-dependent fashion (Stewart et al., 2011; Lilley et al., 2012).

Moderately high temperatures (c. 29°C) also promote hypocotylgrowth (Gray et al., 1998). Recently, it has been discovered thattemperature changes can be sensed by a histone variant. H2A.Z-containing nucleosomes bindDNAmore tightly than typicalH2A-containing nucleosomes, but H2A.Z nucleosome occupancydeclines with increasing temperature (Kumar & Wigge, 2010).This increases the accessibility of DNA to transcription factors andthe transcriptional machinery at higher temperatures. This processprovides a direct functional link between temperature change and a

transcriptional response. Interestingly, the pif4mutation abolishesthe temperature-dependent hypocotyl growth response. PIF4expression is induced by high temperatures (Koini et al., 2009)and enhanced in arp6 mutants that lack H2A.Z incorporation(Kumar et al., 2012). This reflects direct or indirect control of PIF4expression by H2A.Z temperature sensing. PIF4 activity is alsoregulated by the temperature-dependent availability of target locilocated near H2A.Z nucleosomes (Kumar et al., 2012).

Integration We have seen that the activity of PIF4 is modulatedby the circadian clock, metabolic status, temperature and gibber-ellin (GA) levels in addition to light (reviewed in Proveniers & vanZanten, 2013). Recent research has suggested that PIF proteinsform a regulatory module with hormone response pathways,thereby allowing cell growth to respond to and balance an evenwider set of inputs.

DELLA proteins are nuclear-localized growth repressors(DELLA and GA signalling reviewed in Achard & Genschik,2009). DELLAs are known to bind and sequester PIFs, preventingthem from binding DNA. GA promotes growth by promotingDELLA degradation, and its levels are affected by environmental

Fig. 3 Extensive integration of a wide varietyof endogenous and exogenous inputs controlshypocotyl cell growth inArabidopsis thaliana.Conceptual diagram of factors modulatinghypocotyl cell growth, and key regulatorynodes for this modulation. ABA, abscisic acid;ACE1, ACTIVATOR FOR CELLELONGATION1; AIF, ATBS-INTERACTINGFACTORS; BR, brassinosteroid; BZR1,BRASSINOZALE-RESISTANT1; CK, cytokinin;COP1, CONSTITUTIVEPHOTOMORPHOGENIC1; CRY1,CRYPTOCHROME1; ERF1, ETHYLENERESPONSE FACTOR1; GA, gibberellin; HBI1,HOMOLOG OF BEE2 INTERACTINGWITHIBH1; HFR1, LONG HYPOCOTYL IN FAR-RED1; HY5, ELONGATED HYPOCOTYL5;IBH1, ILI1 BINDING BHLH1; PHY,phytochrome; PIF, PHYTOCHROMEINTEGRATING FACTOR; PIL5,PHYTOCHROME INTERACTING FACTOR3-LIKE5; PKL, PICKLE; PRE, PACLOBUTRAZOL-RESISTANT; SAUR, SMALL AUXIN UP RNA.




conditions, including temperature, light and salt stress (Achard &Genschik, 2009). In addition to binding PIFs, DELLAs also bind akey component of the brassinosteroid (BR) signalling pathway, thetranscription factor BRASSINOZALE-RESISTANT1 (BZR1).The DELLA protein REPRESSOR OF GA1 (RGA1) physicallyinteracts with BZR1 and thereby prevents its binding of transcrip-tional targets (Bai et al., 2012b). GA-stimulated hypocotyl elon-gation requires both PIFs and BR signalling, and this interaction issupported by the observation that 92% of genes differentiallyexpressed in both a BR-insensitive and GA-deficient mutant wereaffected in the same way (Bai et al., 2012b).

A multiple PIF mutant, pifq, was found to show reducedhypocotyl elongation in response to BR, and PIF4 overexpressionpartially rescued this phenotype. However, the null mutation ofpif4 showed a similar BR response to the wild-type, suggestingredundancy amongst PIFs in their interaction with BR signalling.BZR1 was found to interact with PIF4 in vitro and in vivo, andchromatin immunoprecipitation (ChIP) studies showed that targetgenes co-occupied the promoters of transcriptional targets in vivo.Chip-seq of BZR1 and PIF4 revealed a large overlap in targets, andthat 80% of common targets were also regulated by light, whereasRNA-seq showed that BZR1 and PIF4 regulated a large number oftarget genes interdependently (Oh et al., 2012). Together, thesedata strongly suggest a central DELLA–BZR1–PIF4 module thatintegrates a diverse range of environmental inputs to modulatehypocotyl cell elongation appropriately (Wang et al., 2012; Casal,2013). It will be fascinating to see how interaction affects the DNAbinding and regulatory activity of BZR1 and PIF4. A candidatemay beGATA2, which is bound directly by BZR1 and regulated byBR, but binding is much higher in dark-grown (when PIF4 levelsare highest) than in light-grown seedlings (Luo et al., 2010).A ChIP of PIF4 in wild-type and bzr1mutant plants would revealhow this interaction affects PIF4 DNA binding globally.

The power of signal integration on cell growth responses in thissystem can be demonstrated through the addition of ethylene. Thishormone suppresses hypocotyl elongation in darkness and pro-motes it in the light, demonstrating howone stimulusmay alter andeven reverse the effects of another stimulus on growth. Theethylene-responsive transcription factor, ETHYLENE-INSENSI-TIVE3 (EIN3), activates PIF3 expression directly, as well asinducing the expression of ETHYLENE RESPONSE FACTOR1(ERF1). PIF3 is required for ethylene-induced hypocotyl growth,whereas ERF1 represses it (Zhong et al., 2012). ERF1 ismore stablein the light than in the dark, andPIF3 ismore stable in the dark thanin the light (as a result of phytochromes). Zhong et al. (2012)predicted that the induction of the less stable protein will have alarger effect on growth. For instance, during the day, ERF1 levelsare high and PIF3 levels are low. The induction of both by ethylenehas a growth-promoting effect because of a large relative increasein PIF3 levels. Another possibility is that the activity of aninteraction factor for PIF3 and/or ERF1 varies depending on lightconditions.

Responses PIF4 plays a crucial role in the regulation of hypocotylcell elongation, partly as a result of its promotion of auxinbiosynthesis. PIF4 binds the promoters of several auxin

biosynthesis genes directly, including YUCCA8 (YUC8) (Sunet al., 2012). Genetic evidence supports this interaction, as YUC8expression is stimulated by temperature increase, but this increase isabsent in pif4 mutants. PIF7 also binds and activates YUC8 andYUC9 promoters directly (Li L et al., 2012).

The gain-of-function mutant indole-3-acetic acid19/massugu2(iaa19/msg2) shows reduced auxin sensitivity, and does not displayenhanced hypocotyl elongation in response to high temperature(Maharjan & Choe, 2011). Furthermore, the gain-of-functionshort hypocotyl2 shy2/iaa3mutant partially suppresses the hypocotylelongation caused by PIF4 overexpression (Sun et al., 2012). Thissuggests that auxin response is required for temperature-dependenthypocotyl elongation. PIF4 is required for temperature-dependentinduction of the SMALL AUXINUP RNA genes SAUR19–24, andSAUR19 overexpression can complement temperature-dependenthypocotyl elongation in pif4 mutants (Franklin et al., 2011). Inaddition to the indirect promotion of elongation through analteration in hormone levels, PIFs may also modulate growth bydirect alteration of the transcription of genes involved in cellgrowth. A gene ontology (GO) analysis of putative PIF-regulatedgenes obtained by combining ChIP-seq data from PIF4 withmicroarrays for multiple pifmutants shows significant enrichmentfor transcriptional regulation and regulation of cell size (Oh et al.,2012).

The PACLOBUTRAZOL-RESISTANT (PRE) family ofHLHtranscription factors is a key output of the DELLA–BZR1–PIFgene regulatory network (GRN). PRE1, PRE5 and PRE6 are directtargets of BZR1 and PIF4, and an artificial microRNA (amiRNA)multiple knockout of PRE1, PRE2, PRE5 and PRE6 results indwarfism, hypersensitivity to light and reduced sensitivity to BR,GA and high temperatures (Bai et al., 2012b; Oh et al., 2012).PREs are atypical HLH proteins that lack a known DNA bindingdomain, and promote cell elongation through antagonisticprotein–protein interactions with growth-repressing proteins, suchas ILI1 BINDING BHLH1 (IBH1) (Zhang et al., 2009). IBH1represses cell elongation by binding and inhibiting the action oftranscription factors that promote hypocotyl elongation, such asACTIVATOR FOR CELL ELONGATION1 (ACE1) andHOMOLOG OF BEE2 INTERACTING WITH IBH1 (HBI)(Bai et al., 2012a; Ikeda et al., 2012). There is further evidence thatPREs also bind and disrupt the activity of ATBS-INTERACTINGFACTORS (AIFs) and LONG HYPOCOTYL IN FAR-RED1(HFR1), which also repress cell elongation (Hyun & Lee, 2006;Wang et al., 2009).

CONSTITUTIVE PHOTOMORPHOGENIC1 (COP1) is aRING E3 ubiquitin ligase which represses photomorphogenesis.COP1 is essential for shade avoidance and is involved in a widevariety of light responses (reviewed in Lau & Deng, 2012). COP1activity is supressed by activated phytochromes. COP1 thereforehas high activity in the dark and poor light. Photomorphogenesis-promoting proteins, and phytochromes, are ubiquitinated byCOP1 (as part of a multi-subunit complex), which triggers theirdegradation by the proteasome (Jang et al., 2010). COP1 alsotargets light signalling transcription factors that promote photo-morphogenesis, including ELONGATED HYPOCOTYL5(HY5), HFR1 and HY5-HOMOLOG (HYH).





PICKLE (PKL/EPP1), an ATP-dependent chromatin remod-elling factor, represses photomorphogenesis and promotes hypo-cotyl cell elongation. PKL antagonizes the repressive activity ofHY5 by reducing H3K27me3 repressive marks at multiple loci in alight-dependent manner (Jing et al., 2013). PKL expression isrepressed by red, far-red and blue light by unknown mechanismsinvolving PhyB, PhyA and cryptochromes, respectively (Jing et al.,2013). pkl mutants have shorter hypocotyls and more expandedcotyledons than the wild-type. Single and double pkl and hy5mutants were found to be antagonistic; target genes repressed in pklwere upregulated in hy5, and the pkl hy5mutant had intermediateexpression levels and phenotype (Jing et al., 2013). Two target locimutants, exp2 and dwf4, also show short hypocotyls when grown inlight (Jing et al., 2013).

2. Regulation of tip growth

We examine both A. thaliana root hairs and pollen tubes asexamples of how tip growth is regulated by complex networks,which integrate multiple diverse inputs. Arabidopsis thaliana hastype III root hair patterning, in which hair and non-hair cells arepresent in separate cell files. The underlying cortex providespositional information, releasing an unknown signal that causesepidermis cells overlapping two cortex cells to become hair cells.Epidermal cells not present over cortex cell junctions become non-hair cells. The GRN that converts the positional cue intodifferentiated cell files has been studied extensively (Grebe, 2012;Ryu et al., 2013).

Hair-forming cells express ROOT HAIR DEFECTIVE6(RHD6) and RHD6-LIKE1 (RSL1), bHLH transcription factorsthat promote root hair outgrowth in a partially redundant manner.Root hair growth is almost abolished in rhd6 mutants. Patterninggenes, such as CAPRICE (CPC), WEREWOLF (WER),TRANSPARENT TESTA GLABRA (TTG) and GLABRA2 (GL2),are epistatic to RHD6 function, and CPC promotes RHD6expression, whereasWER,TTG andGL2 repress it (Menand et al.,2007). This suggests that the patterning network controls theexpression domain of RHD6. Interestingly, the two genes mostsimilar to RHD6 in the moss Physcomitrella patens are required forthe production of analogous rooting structures in the haploidgametophyte (Menand et al., 2007), demonstrating that regulatorynetworks controlling cell growth may be conserved and co-optedacross land plant phylogeny.The bHLHtranscription factor RSL4,a direct target of RHD6, is necessary and sufficient for root hairgrowth. Amicroarray analysis identified c. 80 genes up-regulated byRSL4, 20% of which were exclusively expressed in root hair cells.Five of these genes have a demonstrated role in root hair growth,including a phosphatidylinositol transporter and extensins(Yi et al., 2010; Velasquez et al., 2011). Extensins are structuralcell wall glycoproteins that self-assemble and may guide pectinpositioning (Lamport et al., 2011).

Root hair growth is altered by a wide variety of nutrients (Fig. 4),such as phosphate, iron, zinc and manganese (P�eret et al., 2011).Phosphate has a large effect on A. thaliana root hair development;root hair length is inversely correlated with phosphate concentra-tion (Ma et al., 2001). A recently discovered class of plant

hormones, strigolactones, promotes root hair growth in lowphosphate conditions. This strigolactone-mediated responseappears to be partially dependent on both auxin and ethylenesignalling (Kapulnik et al., 2011a,b). RSL4 integrates nutritionaland hormone responses as its transcription is activated by both lowphosphate availability and auxin, leading to the promotion of roothair elongation (Yi et al., 2010). This induction appears to beindependent of RHD6 and RSL1, suggesting that these environ-mental and developmental signals converse at the level of RSL4transcription (Masucci & Schiefelbein, 1996).

Ethylene promotes root hair elongation (Pitts et al., 1998; Cho& Cosgrove, 2002) in response to both potassium and irondeficiency (Romera & Alc�antara, 2004; Jung et al., 2009).Ethylene-stimulated root hair elongation requires auxin transportand response genes (Strader et al., 2010), suggesting that ethyleneacts in a common pathway with auxin. Cytokinin (CK) inhibitshair growth in potassium-limited plants, and CK levels are reducedin low potassium conditions (Nam et al., 2012). However, CK hasbeen reported to increase root hair length when applied exoge-nously (An et al., 2012).

CK and ethylene signalling is integrated via the action of ZINC-FINGER PROTEIN5 (ZFP5), which directly promotes CPCexpression and enhances root hair elongation (An et al., 2012).CPC overexpression alone has not been observed to increase roothair length (Wada et al., 1997), suggesting the requirement ofadditional factors to enhance root hair elongation. zfp5 mutantshave fewer and shorter root hairs as a result of a lower root hairgrowth rate. Exogenous application of an ethylene precursor and asynthetic CK could increase root hair length in wild-type but notzfp5 plants (An et al., 2012). CK treatment induces ZFP5transcription, whereas ethylene increases ZFP5 protein levelsthrough protein stabilization (An et al., 2012), demonstrating thatCK and ethylene act synergistically to promote ZFP5 activity.

Pollen germination is followed by the formation and mainte-nance of an apical growth domain with highly polarized growth.Pollen tube growthmust be tightly regulated for successful deliveryof the sperm cells for egg cell fertilization and, to do so, the malegametophyte must communicate with multiple female tissues. Ongermination at the stigma, pollen tube growth is successively guidedto the female gametophyte in a multi-step process. In A. thaliana,initial growth from the stigma to the style is mainly mediated bychemo-attractants (Higashiyama & Hamamura, 2008). Thereaf-ter, pollen tube growth is mechanically guided through thetransmitting tract to reach the ovary (Crawford et al., 2007),although a recent study has suggested that cysteine-rich peptides arealso involved (Chae et al., 2009). The last step of pollen tubegrowth involves passage through the septum, along the funiculusand towards the micropylar end of the ovule. Here, the path of thepollen tube is guided by chemo-attractants from the femalegametophyte, such as peptides and amino acids, which triggerreceptor-mediated signalling cascades, ion- and cyclic nucleotide-gated channels at the growth tip of the pollen tube (reviewed byTakeuchi &Higashiyama, 2011). Despite these recent advances inunderstanding the nature of signallingmolecules and the signallingresponses involved, it remains unclear how directional changes inchemo-attractant gradients cause the rearrangement of structural




components, vesicular flows and cytoskeleton dynamics to shift theapical growth domain and alter the direction of pollen tube growth.

VIII. Conclusions

Plant cells convert turgor pressure into growth using tightlyregulated deposition and modification of cell wall components.Complex regulatory networks integrate myriad endogenous andexogenous inputs to produce appropriate responses, which varyacross cell types and species. The understanding of these regulatorysystems has accelerated rapidly in the past decade, and numerouscell growth models have now been characterized in impressivedetail. The links between these systems and the process of growthremain unclear, but growth regulators are regularly linked to largelists of potential targets using high-throughput techniques. Ulti-mately, these growth regulators must modify cell wall properties tomodulate cell growth. To further our understanding, it is crucialthat more functional links are made between regulators of growthand effectors of growth.

Understanding the process of growth is difficult as a result of thecomplexity of plant cell walls, and the incredible ability of plants toadapt to environmental insults (includingmutations) through generedundancy and compensation mechanisms. It is practicallyludicrous that removing a polysaccharide that makes up aroundone-third of the wall by weight (Cosgrove & Jarvis, 2012) does notproduce a major phenotype. A range of new technologies andtechniques should help to overcome these difficulties. Redundancycan be combatted with chemical genetics, sensitized screens andtargeted knockdowns of related genes using amiRNAs (reviewed inSablok et al., 2011; Vidaurre & Bonetta, 2012), and lethality

circumvented with inducible constructs. Visualization techniquesfor wall polysaccharides are improving, with increasingly specificdyes and polysaccharide labelling using click chemistry nowavailable (reviewed in Gonneau et al., 2012; Wallace & Anderson,2012). In addition, microscopy is developing rapidly, with variableangle epifluorescence microscopy (VAEM) able to obtain high-quality images of cytoskeletal dynamics and vesicle deposition atthe plasma membrane (Konopka & Bednarek, 2008), and light-sheet-based fluorescence microscopy (LSFM), which will allowcellular growth to be tracked in three dimensions over long timeperiods at the tissue level (Maizel et al., 2011).

The combination of modelling, structural and mechanical cellwall studies, and genetic approaches, will be required to generate afull understanding of plant cell growth. Modelling approacheshave effectively utilized terms representing structures and pro-cesses suggested by experimental approaches (Kroeger et al., 2011;Dyson et al., 2012), illustrating that communication betweenthese disciplines can be extremely beneficial. In summary, we arebeginning to understand growth regulatory systems, or ‘why’plant cells grow, but ‘how’ plant cells grow remains somethingof a mystery, as challenging and exciting as it was to Lockhart,50 yr ago.

Acknowledgements

This work was supported by a grant from the Ministry ofEducation, Culture, Sports, Science and Technology (MEXT;No.22119010) to K.S.We thankmembers of the Sugimoto laboratoryfor helpful discussions, and Bart Ryman, Momoko Ikeuchi andSarah Ball for comments on the manuscript.

Fig. 4 Fate patterning and environmentalsignals regulate tip growth in Arabidopsis

thaliana root hair cells. Interactions betweenfate specification, the environment andgrowth regulation in root hair cells. The size ofthe protein/hormone icons corresponds totheir concentration relative to the other soilconditions, and the thickness of the arrowscorresponds to the strength of the interaction.CK, cytokinin; CPC, CAPRICE; GL, GLABRA;SL, strigolactone; RHD, ROOT HAIRDEFECTIVE; ROS, reactive oxygen species;RSL4, RHD6-LIKE; TTG, TRANSPARENTTESTA GLABRA; WER, WEREWOLF; ZFP5,ZINC-FINGER PROTEIN5.





References

Achard P, Genschik P. 2009. Releasing the brakes of plant growth: how GAs

shutdown DELLA proteins. Journal of Experimental Botany 60: 1085–1092.An L, Zhou Z, Sun L, Yan A, Xi W, Yu N, Cai W, Chen X, Yu H,

Schiefelbein J et al. 2012. A zinc finger protein gene ZFP5 integrates

phytohormone signaling to control root hair development in Arabidopsis.

Plant Journal 72: 474–490.Bai M-Y, Fan M, Oh E, Wang Z-Y. 2012a. A triple helix–loop–helix/basichelix–loop–helix cascade controls cell elongation downstream of multiple

hormonal and environmental signaling pathways in Arabidopsis. Plant Cell 24:4917–4929.

Bai M-Y, Shang J-X, Oh E, Fan M, Bai Y, Zentella R, Sun T, Wang Z-Y. 2012b.

Brassinosteroid, gibberellin and phytochrome impinge on a common

transcription module in Arabidopsis. Nature Cell Biology 14: 810–817.Baskin TI. 2001.On the alignment of cellulose microfibrils by cortical

microtubules: a review and a model. Protoplasma 215: 150–171.Beemster GTS, Vusser KD, Tavernier ED, Bock KD, Inz�e D. 2002. Variation

in growth rate between Arabidopsis ecotypes is correlated with cell

division and A-type cyclin-dependent kinase activity. Plant Physiology 129:854–864.

BibikovaTN, JacobT,Dahse I,Gilroy S. 1998.Localized changes in apoplastic and

cytoplasmic pH are associatedwith root hair development inArabidopsis thaliana.Development 125: 2925–2934.

Boss WF, Im YJ. 2012. Phosphoinositide signaling. Annual Review of Plant Biology63: 409–429.

Boyer JS. 2009.Evans review: cell wall biosynthesis and themolecularmechanismof

plant enlargement. Functional Plant Biology 36: 383–394.Breton G, Kay SA. 2007. Plant biology: time for growth. Nature 448: 265–266.Breuer C, Ishida T, Sugimoto K. 2010.Developmental control of endocycles and

cell growth in plants. Current Opinion in Plant Biology 13: 654–660.Breuer C, Morohashi K, Kawamura A, Takahashi N, Ishida T, Umeda M,

Grotewold E, Sugimoto K. 2012. Transcriptional repression of the APC/C

activator CCS52A1 promotes active termination of cell growth. EMBO Journal31: 4488–4501.

BringmannM,Li E, Sampathkumar A,KocabekT,HauserM-T, Persson S. 2012.

POM-POM2/CELLULOSE SYNTHASE INTERACTING1 is essential for thefunctional association of cellulose synthase and microtubules in Arabidopsis.

Plant Cell 24: 163–177.Bunzel M. 2010. Chemistry and occurrence of hydroxycinnamate oligomers.

Phytochemistry Reviews 9: 47–64.Caffall KH, Mohnen D. 2009. The structure, function, and biosynthesis of plant

cell wall pectic polysaccharides. Carbohydrate Research 344: 1879–1900.Carol RJ, Takeda S, Linstead P, DurrantMC, Kakesova H, Derbyshire P, Drea S,

Zarsky V, Dolan L. 2005. A RhoGDP dissociation inhibitor spatially regulates

growth in root hair cells. Nature 438: 1013–1016.Casal JJ. 2013. Photoreceptor signaling networks in plant responses to shade.

Annual Review of Plant Biology 64: 403–427.Chae K, Kieslich CA, Morikis D, Kim S-C, Lord EM. 2009. A gain-of-function

mutation of Arabidopsis Lipid Transfer Protein 5 disturbs pollen tube tip growth

and fertilization. Plant Cell 21: 3902–3914.Chebli Y, Kaneda M, Zerzour R, Geitmann A. 2012. The cell wall of the

Arabidopsis pollen tube – spatial distribution, recycling, and network formation

of polysaccharides. Plant Physiology 160: 1940–1955.Chen M, Chory J. 2011. Phytochrome signaling mechanisms and the control of

plant development. Trends in Cell Biology 21: 664–671.ChoH-T, Cosgrove DJ. 2002.Regulation of root hair initiation and expansin gene

expression in Arabidopsis. Plant Cell 14: 3237–3253.Cosgrove DJ. 2000. Loosening of plant cell walls by expansins. Nature 407:321–326.

Cosgrove DJ. 2005. Growth of the plant cell wall. Nature Reviews Molecular CellBiology 6: 850–861.

Cosgrove DJ, Bedinger P, Durachko DM. 1997.Group I allergens of grass pollen

as cell wall-loosening agents. Proceedings of the National Academy of Sciences, USA94: 6559–6564.

Cosgrove DJ, Jarvis MC. 2012. Comparative structure and biomechanics of plant

primary and secondary cell walls. Frontiers in Plant Science 3: 204.

Crawford BCW, Ditta G, Yanofsky MF. 2007. The NTT gene is required for

transmitting-tract development in carpels ofArabidopsis thaliana.Current Biology17: 1101–1108.

De Veylder L, Larkin JC, Schnittger A. 2011.Molecular control and function

of endoreplication in development and physiology. Trends in Plant Science 16:624–634.

Dick-P�erez M, Zhang Y, Hayes J, Salazar A, Zabotina OA, Hong M. 2011.

Structure and interactions of plant cell-wall polysaccharides by two- and

three-dimensional magic-angle-spinning solid-state NMR. Biochemistry 50:989–1000.

Dyson RJ, Band LR, Jensen OE. 2012. A model of crosslink kinetics in the

expanding plant cell wall: yield stress and enzyme action. Journal of TheoreticalBiology 307: 125–136.

Ekl€of JM, Brumer H. 2010.The XTH gene family: an update on enzyme structure,

function, and phylogeny in xyloglucan remodeling. Plant Physiology 153: 456–466.

Figueiredo SA, Lashermes P, Arag~ao FJL. 2011.Molecular characterization and

functional analysis of the b-galactosidase gene during Coffea arabica (L.) fruitdevelopment. Journal of Experimental Botany 62: 2691–2703.

FischerU, IkedaY, LjungK, SerralboO, SinghM,HeidstraR, PalmeK, ScheresB,

Grebe M. 2006. Vectorial information for Arabidopsis planar polarity is

mediated by combined AUX1, EIN2, and GNOM activity. Current Biology: CB16: 2143.

Forouzesh E, Goel A, Mackenzie SA, Turner JA. 2013. In vivo extraction of

Arabidopsis cell turgor pressure using nanoindentation in conjunction with finite

element modeling. Plant Journal 73: 509–520.Franklin KA, Lee SH, Patel D, Kumar SV, Spartz AK, Gu C, Ye S, Yu P, Breen G,

Cohen JD et al. 2011. PHYTOCHROME-INTERACTING FACTOR 4

(PIF4) regulates auxin biosynthesis at high temperature. Proceedings of theNational Academy of Sciences, USA 108: 20231–20235.

Geitmann A, Ortega JKE. 2009.Mechanics and modeling of plant cell growth.

Trends in Plant Science 14: 467–478.Gendreau E, Traas J, Desnos T, Grandjean O, Caboche M, Hofte H. 1997.

Cellular basis of hypocotyl growth in Arabidopsis thaliana. Plant Physiology 114:295–305.

Georgelis N, Tabuchi A, Nikolaidis N, Cosgrove DJ. 2011. Structure–functionanalysis of the bacterial expansin EXLX1. Journal of Biological Chemistry 286:16814–16823.

Giddings TH, Staehelin LA. 1991.Microtubule-mediated control of microfibril

deposition: a re-examination of the hypothesis. In: Lloyd CW, ed. Thecytoskeletal basis of plant growth and form. London, UK: Academic Press,

85–99.Goh H-H, Sloan J, Dorca-Fornell C, Fleming A. 2012. Inducible repression of

multiple expansin genes leads to growth suppression during leaf development.

Plant Physiology 159: 1759–1770.Gonneau M, Hofte H, Vernhettes S. 2012. Fluorescent tags to explore cell wall

structure and dynamics. Frontiers in Plant Science 3: 145.GrayWM, €Ostin A, SandbergG, RomanoCP, EstelleM. 1998.High temperature

promotes auxin-mediated hypocotyl elongation in Arabidopsis. Proceedings of theNational Academy of Sciences, USA 95: 7197–7202.

Grebe M. 2012. The patterning of epidermal hairs in Arabidopsis – updated.Current Opinion in Plant Biology 15: 31–37.

Gu Y, Kaplinsky N, BringmannM, Cobb A, Carroll A, Sampathkumar A, Baskin

TI, Persson S, Somerville CR. 2010. Identification of a cellulose

synthase-associated protein required for cellulose biosynthesis. Proceedings of theNational Academy of Sciences, USA 107: 12866–12871.

Gutierrez R, Lindeboom JJ, Paredez AR, Emons AMC, Ehrhardt DW. 2009.

Arabidopsis cortical microtubules position cellulose synthase delivery to the

plasmamembrane and interact with cellulose synthase trafficking compartments.

Nature Cell Biology 11: 797–806.HamantO,HeislerMG, JonssonH, Krupinski P, UyttewaalM, Bokov P, Corson

F, Sahlin P, Boudaoud A, Meyerowitz EM et al. 2008. Developmental

patterning by mechanical signals in Arabidopsis. Science 322: 1650–1655.Heath IB. 1974. A unified hypothesis for the role of membrane bound enzyme

complexes and microtubules in plant cell wall synthesis. Journal of TheoreticalBiology 48: 445–449.




Higashiyama T, Hamamura Y. 2008.Gametophytic pollen tube guidance. SexualPlant Reproduction 21: 17–26.

Himmelspach R, Williamson RE, Wasteneys GO. 2003. Cellulose microfibril

alignment recovers from DCB-induced disruption despite microtubule

disorganization. Plant Journal 36: 565–575.Hyun Y, Lee I. 2006. KIDARI, encoding a non-DNA binding bHLH protein,

represses light signal transduction in Arabidopsis thaliana. PlantMolecular Biology61: 283–296.

Ikeda M, Fujiwara S, Mitsuda N, Ohme-Takagi M. 2012. A triantagonistic basic

helix–loop–helix system regulates cell elongation in Arabidopsis. Plant Cell 24:4483–4497.

Jang I-C, Henriques R, Seo HS, Nagatani A, Chua N-H. 2010. Arabidopsis

PHYTOCHROME INTERACTING FACTOR proteins promote

phytochrome B polyubiquitination by COP1 E3 ligase in the nucleus. Plant Cell22: 2370–2383.

J�eguT,LatrasseD,DelarueM,MazubertC,BourgeM,HudikE,Blanchet S, Soler

M-N, Charon C, De Veylder L et al. 2013.Multiple functions of Kip-related

proteins connect endoreduplication and cell elongation. Plant Physiology 161:1694–1705.

Jing Y, ZhangD,Wang X, TangW,WangW,Huai J, Xu G, ChenD, Li Y, Lin R.

2013. Arabidopsis chromatin remodeling factor PICKLE interacts with

transcription factor HY5 to regulate hypocotyl cell elongation. Plant Cell 25:242–256.

Johnston SL, Prakash R, Chen NJ, Kumagai MH, Turano HM, Cooney JM,

Atkinson RG, Paull RE, Cheetamun R, Bacic A et al. 2013. An enzyme activity

capable of endotransglycosylation of heteroxylan polysaccharides is present in

plant primary cell walls. Planta 237: 173–187.Jolie RP, Duvetter T, Van Loey AM, Hendrickx ME. 2010. Pectin

methylesterase and its proteinaceous inhibitor: a review. Carbohydrate Research345: 2583–2595.

Jung J-Y, ShinR, SchachtmanDP. 2009.Ethylenemediates response and tolerance

to potassium deprivation in Arabidopsis. Plant Cell 21: 607–621.Kapulnik Y, Delaux P-M, Resnick N,Mayzlish-Gati E,Wininger S, Bhattacharya

C, S�ejalon-Delmas N, Combier J-P, B�ecard G, Belausov E et al. 2011a.Strigolactones affect lateral root formation and root-hair elongation in

Arabidopsis. Planta 233: 209–216.Kapulnik Y, Resnick N, Mayzlish-Gati E, Kaplan Y, Wininger S, Hershenhorn J,

Koltai H. 2011b. Strigolactones interact with ethylene and auxin in regulating

root-hair elongation in Arabidopsis. Journal of Experimental Botany 62:2915–2924.

Kierzkowski D, Nakayama N, Routier-Kierzkowska A-L, Weber A, Bayer E,

Schorderet M, Reinhardt D, Kuhlemeier C, Smith RS. 2012. Elastic domains

regulate growth and organogenesis in the plant shoot apical meristem. ScienceSignaling 335: 1096.

KoiniMA,Alvey L, AllenT, TilleyCA,HarberdNP,WhitelamGC, FranklinKA.

2009.High temperature-mediated adaptations in plant architecture require the

bHLH transcription factor PIF4. Current Biology 19: 408–413.Konopka CA, Bednarek SY. 2008. Variable-angle epifluorescence microscopy: a

new way to look at protein dynamics in the plant cell cortex. Plant Journal 53:186–196.

Kroeger JH, Zerzour R, Geitmann A. 2011.Regulator or driving force? The role of

turgor pressure in oscillatory plant cell growth. PLoS ONE 6: e18549.

Kumar SV, Lucyshyn D, Jaeger KE, Al�os E, Alvey E, Harberd NP, Wigge PA.

2012. Transcription factor PIF4 controls the thermosensory activation of

flowering. Nature 484: 242–245.Kumar SV, Wigge PA. 2010.H2A.Z-containing nucleosomes mediate the

thermosensory response in Arabidopsis. Cell 140: 136–147.Kutschera U, Niklas KJ. 2013. Cell division and turgor-driven stem elongation in

juvenile plants: a synthesis. Plant Science 207: 45–56.Lamport DTA, Kieliszewski MJ, Chen Y, Cannon MC. 2011. Role of the

extensin superfamily in primary cell wall architecture. Plant Physiology 156:11–19.

Lau OS, Deng XW. 2012. The photomorphogenic repressors COP1 and DET1:

20 years later. Trends in Plant Science 17: 584–593.Lee YJ, Yang Z. 2008.Tip growth: signaling in the apical dome.Current Opinion inPlant Biology 11: 662–671.

Li L, Ljung K, Breton G, Schmitz RJ, Pruneda-Paz J, Cowing-Zitron C, Cole BJ,

Ivans LJ, PedmaleUV, JungH-S et al.2012.Linking photoreceptor excitation tochanges in plant architecture. Genes & Development 26: 785–790.

Li S, Lei L, Somerville CR, Gu Y. 2012. Cellulose synthase interactive protein 1

(CSI1) links microtubules and cellulose synthase complexes. Proceedings of theNational Academy of Sciences, USA 109: 185–190.

Lilley JLS, Gee CW, Sairanen I, Ljung K, Nemhauser JL. 2012. An endogenous

carbon-sensing pathway triggers increased auxin flux and hypocotyl elongation.

Plant Physiology 160: 2261–2270.Liners F, Letesson J-J, Didembourg C, Cutsem PV. 1989.Monoclonal antibodies

against pectin recognition of a conformation induced by calcium.Plant Physiology91: 1419–1424.

Lionetti V, Cervone F, Bellincampi D. 2012.Methyl esterification of pectin plays a

role during plant–pathogen interactions and affects plant resistance to diseases.

Journal of Plant Physiology 169: 1623–1630.Lockhart JA. 1965. An analysis of irreversible plant cell elongation. Journal ofTheoretical Biology 8: 264–275.

Luo X-M, Lin W-H, Zhu S, Zhu J-Y, Sun Y, Fan X-Y, Cheng M, Hao Y, Oh E,

Tian M et al. 2010. Integration of light- and brassinosteroid-signaling pathwaysby a GATA transcription factor in Arabidopsis.Developmental Cell 19: 872–883.

MaZ,BielenbergDG,BrownKM,Lynch JP. 2001.Regulation of root hair density

by phosphorus availability in Arabidopsis thaliana. Plant, Cell & Environment 24:459–467.

Maharjan PM, Choe S. 2011.High temperature stimulates DWARF4 (DWF4)expression to increase hypocotyl elongation in Arabidopsis. Journal of PlantBiology 54: 425–429.

Maizel A, von Wangenheim D, Federici F, Haseloff J, Stelzer EHK. 2011.

High-resolution live imaging of plant growth in near physiological bright

conditions using light sheet fluorescence microscopy. Plant Journal 68: 377–385.Marcus SE, Verhertbruggen Y, Herv�e C, Ordaz-Ortiz JJ, Farkas V, Pedersen HL,

Willats WG, Knox JP. 2008. Pectic homogalacturonan masks abundant sets of

xyloglucan epitopes in plant cell walls. BMC Plant Biology 8: 60.Martin C, Bhatt K, Baumann K. 2001. Shaping in plant cells. Current Opinion inPlant Biology 4: 540–549.

Masucci JD, Schiefelbein JW. 1996.Hormones act downstream of TTG and GL2to promote root hair outgrowthduring epidermis development in theArabidopsis

root. Plant Cell 8: 1505–1517.McQueen-Mason S, Durachko DM, Cosgrove DJ. 1992. Two endogenous

proteins that induce cell wall extension in plants. Plant Cell 4: 1425–1433.Melaragno JE, Mehrotra B, Coleman AW. 1993. Relationship between

endopolyploidy and cell size in epidermal tissue of Arabidopsis. Plant Cell 5:1661–1668.

Menand B, Yi K, Jouannic S, Hoffmann L, Ryan E, Linstead P, Schaefer DG,

Dolan L. 2007. An ancient mechanism controls the development of cells with a

rooting function in land plants. Science 316: 1477–1480.Mohler KE, Simmons TJ, Fry SC. 2013.Mixed-linkage glucan:xyloglucan

endotransglucosylase (MXE) re-models hemicelluloses in Equisetum shoots but

not in barley shoots or Equisetum callus. New Phytologist 197: 111–122.Nam Y-J, Tran L-SP, Kojima M, Sakakibara H, Nishiyama R, Shin R. 2012.

Regulatory roles of cytokinins and cytokinin signaling in response to potassium

deficiency in Arabidopsis. PLoS ONE 7: e47797.

Nomoto Y, Kubozono S, Miyachi M, Yamashino T, Nakamichi N, Mizuno T.

2012a. A circadian clock- and PIF4-mediated double coincidence mechanism is

implicated in the thermosensitive photoperiodic control of plant architectures in

Arabidopsis thaliana. Plant and Cell Physiology 53: 1965–1973.Nomoto Y, Kubozono S, Yamashino T, Nakamichi N, Mizuno T. 2012b.

Circadian clock- and PIF4-controlled plant growth: a coincidence mechanism

directly integrates a hormone signaling network into the photoperiodic control

of plant architectures in Arabidopsis thaliana. Plant and Cell Physiology 53:1950–1964.

NozueK,CovingtonMF,DuekPD, Lorrain S, FankhauserC,Harmer SL,Maloof

JN. 2007. Rhythmic growth explained by coincidence between internal and

external cues. Nature 448: 358–361.Nusinow DA, Helfer A, Hamilton EE, King JJ, Imaizumi T, Schultz TF, Farr�e

EM, Kay SA. 2011. The ELF4–ELF3–LUX complex links the circadian clock to

diurnal control of hypocotyl growth. Nature 475: 398–402.





Oh E, Zhu J-Y, Wang Z-Y. 2012. Interaction between BZR1 and PIF4 integrates

brassinosteroid and environmental responses. Nature Cell Biology 14: 802–809.Ove�cka M, Berson T, Beck M, Derksen J, �Samaj J, Balu�ska F, Lichtscheidl IK.2010. Structural sterols are involved in both the initiation and tip growth of root

hairs in Arabidopsis thaliana. Plant Cell 22: 2999–3019.Paredez AR, Somerville CR, Ehrhardt DW. 2006. Visualization of cellulose

synthase demonstrates functional association with microtubules. Science 312:1491–1495.

Park YB, Cosgrove DJ. 2012a. A revised architecture of primary cell walls based on

biomechanical changes induced by substrate-specific endoglucanases. PlantPhysiology 158: 1933–1943.

Park YB, CosgroveDJ. 2012b.Changes in cell wall biomechanical properties in the

xyloglucan-deficient xxt1/xxt2mutant of Arabidopsis. Plant Physiology 158:465–475.

Peaucelle A, Braybrook S, Hofte H. 2012.Cell wall mechanics and growth control

in plants: the role of pectins revisited. Frontiers in Plant Science 3: 121.PeaucelleA, LouvetR, Johansen JN,H€ofteH,Laufs P, Pelloux J,MouilleG. 2008.

Arabidopsis phyllotaxis is controlledby themethyl-esterification status of cell-wall

pectins. Current Biology 18: 1943–1948.Peaucelle A, Louvet R, Johansen JN, Salsac F, Morin H, Fournet F, Belcram K,

Gillet F, H€ofte H, Laufs P et al. 2011. The transcription factor BELLRINGER

modulates phyllotaxis by regulating the expression of a pectin methylesterase in

Arabidopsis. Development 138: 4733–4741.Pelletier S, VanOrden J,Wolf S, Vissenberg K,Delacourt J, Ndong YA, Pelloux J,

Bischoff V, Urbain A, Mouille G et al. 2010. A role for pectin

de-methylesterification in a developmentally regulated growth acceleration in

dark-grown Arabidopsis hypocotyls. New Phytologist 188: 726–739.Pelloux J, Rust�erucci C, Mellerowicz EJ. 2007. New insights into pectin

methylesterase structure and function. Trends in Plant Science 12: 267–277.P�eret B, Cl�ement M, Nussaume L, Desnos T. 2011. Root developmental

adaptation to phosphate starvation: better safe than sorry. Trends in Plant Science16: 442–450.

Persson S, Paredez A,Carroll A, PalsdottirH,DoblinM, Poindexter P,KhitrovN,

AuerM, Somerville CR. 2007.Genetic evidence for three unique components in

primary cell-wall cellulose synthase complexes in Arabidopsis. Proceedings of theNational Academy of Sciences, USA 104: 15566–15571.

PietruszkaM. 2011. Solutions for a local equation of anisotropic plant cell growth:

an analytical study of expansin activity. Journal of the Royal Society, Interface 8:975–987.

Pitts RJ, Cernac A, Estelle M. 1998. Auxin and ethylene promote root hair

elongation in Arabidopsis. Plant Journal 16: 553–560.Proveniers MCG, van Zanten M. 2013.High temperature acclimation through

PIF4 signaling. Trends in Plant Science 18: 59–64.Ray PM, Green PB, Cleland R. 1972. Role of turgor in plant cell growth. Nature239: 163–164.

RayleDL. 1973.Auxin-induced hydrogen-ion secretion inAvena coleoptiles and itsimplications. Planta 114: 63–73.

R€ockel N,Wolf S, Kost B, Rausch T, Greiner S. 2008. Elaborate spatial patterning

of cell-wall PME and PMEI at the pollen tube tip involves PMEI endocytosis, and

reflects the distribution of esterified and de-esterified pectins. Plant Journal 53:133–143.

Romera FJ, Alc�antara E. 2004. Ethylene involvement in the regulation of

Fe-deficiency stress responses by Strategy I plants. Functional Plant Biology 31:315–328.

R€osti J, Barton CJ, Albrecht S, Dupree P, Pauly M, Findlay K, Roberts K, Seifert

GJ. 2007. UDP-glucose 4-epimerase isoforms UGE2 and UGE4 cooperate in

providing UDP-galactose for cell wall biosynthesis and growth of Arabidopsisthaliana. Plant Cell 19: 1565–1579.

Rounds CM, Bezanilla M. 2013. Growth mechanisms in tip-growing plant cells.

Annual Review of Plant Biology 64: 243–265.Ryu KH, Zheng X, Huang L, Schiefelbein J. 2013. Computational modeling of

epidermal cell fate determination systems. Current Opinion in Plant Biology 16:5–10.

Sablok G, P�erez-Quintero �AL, Hassan M, Tatarinova TV, L�opez C. 2011.

Artificial microRNAs (amiRNAs) engineering – on how microRNA-based

silencing methods have affected current plant silencing research. Biochemical andBiophysical Research Communications 406: 315–319.

Sampedro J, GianzoC, IglesiasN,Guiti�an E, RevillaG, Zarra I. 2012.AtBGAL10

is the main xyloglucan b-galactosidase in Arabidopsis, and its absence results

in unusual xyloglucan subunits and growth defects. Plant Physiology 158:1146–1157.

Scheller HV, Ulvskov P. 2010.Hemicelluloses. Annual Review of Plant Biology 61:263–289.

Schnittger A, Weinl C, Bouyer D, Sch€obinger U, H€ulskamp M. 2003.

Misexpression of the cyclin-dependent kinase inhibitor ICK1/KRP1 insingle-celled Arabidopsis trichomes reduces endoreduplication and cell size and

induces cell death. Plant Cell 15: 303–315.Schopfer P. 2008. Is the loss of stability theory a realistic concept for stress

relaxation-mediated cell wall expansion during plant growth? Plant Physiology147: 935–938.

Somerville C. 2006.Cellulose synthesis in higher plants. Annual Review of Cell andDevelopmental Biology 22: 53–78.

Speranza A, Crinelli R, Scoccianti V, Geitmann A. 2012. Reactive oxygen

species are involved in pollen tube initiation in kiwifruit. Plant Biology 14:

64–76.Stewart JL, Maloof JN, Nemhauser JL. 2011. PIF genes mediate the effect of

sucrose on seedling growth dynamics. PLoS ONE 6: e19894.

Strader LC, Chen GL, Bartel B. 2010. Ethylene directs auxin to control root cell

expansion. Plant Journal 64: 874–884.SugimotoK,HimmelspachR,WilliamsonRE,WasteneysGO. 2003.Mutation or

drug-dependent microtubule disruption causes radial swelling without altering

parallel cellulose microfibril deposition in Arabidopsis root cells. Plant Cell 15:1414–1429.

Sugimoto K, Williamson RE, Wasteneys GO. 2000. New techniques enable

comparative analysis of microtubule orientation, wall texture, and growth rate in

intact roots of Arabidopsis. Plant Physiology 124: 1493–1506.Sun J, Qi L, Li Y, Chu J, Li C. 2012. PIF4-mediated activation of YUCCA8expression integrates temperature into the auxin pathway in regulating

Arabidopsis hypocotyl growth. PLoS Genetics 8: e1002594.Szymanski DB, Cosgrove DJ. 2009.Dynamic coordination of cytoskeletal and cell

wall systems during plant cell morphogenesis. Current Biology 19: R800–R811.Tabuchi A, Li L-C, Cosgrove DJ. 2011.Matrix solubilization and cell wall

weakening by b-expansin (group-1 allergen) frommaize pollen. Plant Journal 68:546–559.

Takeda S, Gapper C, Kaya H, Bell E, Kuchitsu K, Dolan L. 2008. Local positive

feedback regulation determines cell shape in root hair cells. Science 319:1241–1244.

Takeuchi H, Higashiyama T. 2011. Attraction of tip-growing pollen tubes by the

female gametophyte. Current Opinion in Plant Biology 14: 614–621.Tian G-W, Chen M-H, Zaltsman A, Citovsky V. 2006. Pollen-specific pectin

methylesterase involved in pollen tube growth. Developmental Biology 294:83–91.

Velasquez SM, Ricardi MM,Dorosz JG, Fernandez PV, Nadra AD, Pol-Fachin L,

Egelund J, Gille S, Harholt J, Ciancia M et al. 2011.O-glycosylated cell wall

proteins are essential in root hair growth. Science 332: 1401–1403.Vidaurre D, Bonetta D. 2012. Accelerating forward genetics for cell wall

deconstruction. Frontiers in Plant Science 3: 119.Wada T, Tachibana T, Shimura Y, Okada K. 1997. Epidermal cell

differentiation in Arabidopsis determined by a Myb homolog, CPC. Science277: 1113–1116.

Wallace IS, Anderson CT. 2012. Small molecule probes for plant cell wall

polysaccharide imaging. Frontiers in Plant Science 3: 89.Wang Z-Y, Bai M-Y, Oh E, Zhu J-Y. 2012. Brassinosteroid signaling network and

regulation of photomorphogenesis. Annual Review of Genetics 46: 701–724.Wang H, Zhu Y, Fujioka S, Asami T, Li J, Li J. 2009. Regulation of Arabidopsisbrassinosteroid signaling by atypical basic helix-loop-helix proteins.Plant Cell 21:3781–3791.

Wei C, Lintilhac PM. 2003. Loss of stability – a new model for stress relaxation in

plant cell walls. Journal of Theoretical Biology 224: 305–312.Wei C, Lintilhac PM. 2007. Loss of stability: a new look at the physics of cell wall

behaviour during plant cell growth. Plant Physiology 145: 763–772.Winship LJ, Obermeyer G, Geitmann A, Hepler PK. 2011. Pollen tubes and the

physical world. Trends in Plant Science 16: 353–355.




Yi K, Menand B, Bell E, Dolan L. 2010. A basic helix–loop–helix transcriptionfactor controls cell growth and size in root hairs. Nature Genetics 42: 264–267.

Yoneda A, Higaki T, Kutsuna N, Kondo Y, Osada H, Hasezawa S, Matsui M.

2007.Chemical genetic screening identifies a novel inhibitor of parallel alignment

of cortical microtubules and cellulose microfibrils. Plant and Cell Physiology 48:1393–1403.

YonedaA, ItoT,HigakiT,KutsunaN, SaitoT, IshimizuT,OsadaH,HasezawaS,

Matsui M, Demura T. 2010. Cobtorin target analysis reveals that pectin

functions in the deposition of cellulose microfibrils in parallel with cortical

microtubules. Plant Journal 64: 657–667.Zhang L-Y, BaiM-Y,Wu J, Zhu J-Y,WangH, Zhang Z,WangW, Sun Y, Zhao J,

Sun X et al. 2009. Antagonistic HLH/bHLH transcription factors mediate

brassinosteroid regulation of cell elongation and plant development in rice and

Arabidopsis. Plant Cell 21: 3767–3780.Zhang Y, Mayba O, Pfeiffer A, Shi H, Tepperman JM, Speed TP, Quail

PH. 2013. A quartet of PIF bHLH factors provides a transcriptionally

centered signaling hub that regulates seedling morphogenesis through

differential expression-patterning of shared target genes in Arabidopsis. PLoSGenetics 9: e1003244.

Zhong S, ShiH, XueC,Wang L, Xi Y, Li J, Quail PH,Deng XW,GuoH. 2012.A

molecular framework of light-controlled phytohormone action in Arabidopsis.

Current Biology 22: 1530–1535.Zonia L, Munnik T. 2011. Understanding pollen tube growth: the hydrodynamic

model versus the cell wall model. Trends in Plant Science 16: 347–352.





my body is a cage: mechanisms and modulation of plant cell growth

Documents