diffuse growth of plant cell walls1[open]update on plant cell walls diffuse growth of plant cell...

Update on Plant Cell Walls

Diffuse Growth of Plant Cell Walls1[OPEN]

Daniel J. Cosgrove2

Department of Biology, Penn State University, University Park, Pennsylvania 16802

ORCID ID: 0000-0002-4020-5786 (D.J.C.).

The primary wall of a growing cell is a versatile,subtle, and dynamic structure, with unique propertiesand functions in the life of the plant (Burton et al., 2010).When a cell grows, its wall stretches irreversibly as thecell enlarges in volume. Cells can start and stop thisprocess quickly, in less than a minute in some cases,revealing that the molecular processes underlying ir-reversible wall expansion are dynamically controlled.Such dynamic behavior may be mediated, at least inpart, by changes in wall pH (Hager, 2003; Barbez et al.,2017), which modulates the wall-loosening action ofexpansins (Cosgrove, 2015) and potentially other wall-modifying agents. Wall pH in turn is dynamicallymodulated by plasma membrane H+-ATPase activity(Haruta et al., 2014, 2015) and other processes in thewall. Because growing cell walls are thin and in closephysical contact with plasma membranes, wall pH canbe rapidly modulated (Bibikova et al., 1998;Monshausen et al., 2007; Barbez et al., 2017). As a resultof the pH-dependent activity of expansins, the growingcell wall behaves like a “smart material”—one whoseproperties (extensibility in this case) reversibly andrapidly change with environment (e.g. pH). Slowerchanges in wall structure that influence the wall’sability to expand also occur as part of the natural courseof cell development, e.g. as cells are displaced throughthe elongation zone of a stem (Phyo et al., 2017), or inresponse to external perturbations, e.g. Sahaf andSharon (2016). These slower changes may includechanges inmechanics, such aswall stiffening, and in thedensity or accessibility of sites where expansins or otherproteins can loosen the wall.

The wall itself is synthesized in a team effort: mobilecellulose synthesis complexes (Paredez et al., 2006; Liet al., 2016b) produce long, thin, strong, stiff cellulosemicrofibrils at the cell surface, while matrix polysac-charides and glycoproteins are deposited to the cellsurface via the secretory apparatus (Zhu et al., 2015;

Kim and Brandizzi, 2016). The cytoskeleton guides thewall synthesis machinery to supply wall components toappropriate locations on the cell surface (Szymanskiand Staiger, 2017), where the components assemble to

1 Work on expansins is supported by the U.S. Department of En-ergy (grant no. DE-FG2-84ER13179) andwork on cell wall structure issupported as part of the Center for LignoCellulose Structure andFormation, an Energy Frontier Research Center funded by the U.S.Department of Energy, Office of Science, Basic Energy Sciences(award no. DE-SC0001090).

2 Address correspondence to [email protected]. wrote the article.[OPEN] Articles can be viewed without a subscription.www.plantphysiol.org/cgi/doi/10.1104/pp.17.01541

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form an organized, mechanically strong structure thatcan withstand the in-plane tensile forces generated bythe outward push of cell turgor pressure yet is able toexpand in a controlled manner. The structural re-quirements for orderly expansion of the cell wall are notwell defined at this time. Moreover, except with thepossible exception of tip-growing cells (Dumais et al.,2006; Rojas et al., 2011), synthesis, secretion, and wallassembly are only distantly coupled to the wall exten-sion process itself. For instance, cellulose synthesis incarbon-limited Arabidopsis (Arabidopsis thaliana) hy-pocotyls was temporally distinct from cell expansion(Ivakov et al., 2017); likewise, wall deposition did notkeep up with cell expansion in dark-grown hypocotyls,resulting in substantial wall thinning (Refrégier et al.,2004). On the other hand, gradients in wall thickness inthe growing trichome of Arabidopsis preciselymatchedpredictions for mechanical stability of the wall, imply-ing good coordination between local wall depositionand expansion (Yanagisawa et al., 2015). We still havemuch to learn about how plant cells build a stable yetextensible wall.When the normal molecular assembly of the cell wall

is disturbed, for instance, by mutations that affect syn-thesis of cellulose (Fagard et al., 2000), xyloglucan(Cavalier et al., 2008), or pectic polysaccharides (Mouilleet al., 2007), cell expansion may be disrupted in unpre-dictable ways. Genetic results suggest that errors in wallassembly trigger surface receptor-like kinases such asFERONIA that may act as sensors of cell wall integrity(CWI; Humphrey et al., 2007; Höfte, 2015) and mecha-nosensation (Hamant and Haswell, 2017). The ensuingresponses, which include production of reactive oxygenspecies and inactivation of the plasma membrane H+

-ATPase, likely give rise to some of the complex phe-notypes that originate from rather simple modificationsof wall polysaccharides (Voxeur and Höfte, 2016). Forexample, growth defects stemming from mutation of acellulose synthase gene in Arabidopsis were partiallysuppressed by mutation of THESEUS1 (Hématy et al.,2007), another member of the same receptor kinasefamily as FERONIA (Cheung and Wu, 2011; Li et al.,2016a). Evidently, CWI responses compound and con-found the direct effects of cell wall defects. Defects inpectin metabolism appear particularly prone to triggerCWI responses that activate the brassinosteroid path-way, leading to diverse growth phenotypes (Wolf et al.,2012, 2014). On the other hand, FERONIA and its ex-tracellular peptide ligand (“rapid alkalinization factor”)are also required for normal root growth and auxin re-sponses (Haruta et al., 2014; Shih et al., 2014; Velasquezet al., 2016; Barbez et al., 2017). Cell expansion thus ap-pears to be intimately linked to these wall sensor path-ways in ways we are only beginning to fathom.This Update focuses on the growing cell wall, in

particular, the structural, mechanical, and physico-chemical processes underlying irreversible wall en-largement during diffuse cell growth. Diffuse growthrefers to surface expansion occurring on entire facets ofcell walls, for instance, the side walls of elongating cells

in the body of a growing root or stem. Diffuse growthmay occur with or without a directional bias, whichdepends partly on wall structure and partly on patternsof mechanical stress in the wall (Baskin and Jensen,2013). Its intensity may vary along a cell wall surfaceand on different cell wall facets. For instance, side wallsof a hypocotyl cell may elongate rapidly, whereas itsend walls may not enlarge much at all (Peaucelle et al.,2015). In the jigsaw-puzzle-like pavement cells of theArabidopsis leaf epidermis, a complex pattern of localwall surface expansion occurs in the periclinal (outerepidermal) wall as well as in the anticlinal (side) walls(Szymanski, 2014; Armour et al., 2015). These complexexpansion patterns have been linked to cytoskeletaldynamics within the cell and to spatial patterns oftensile stress (Szymanski and Cosgrove, 2009; Zhanget al., 2013; Sampathkumar et al., 2014a).

Diffuse growth is the dominant pattern for most cellsin the plant body and is traditionally contrastedwith tipgrowth, for instance, in pollen tubes and root hairs,where surface expansion is localized to limited regionsof the hemispherical tip (Campàs et al., 2012; SanatiNezhad and Geitmann, 2013; Velasquez et al., 2016).Despite differences in spatial patterning, tip growthmay involve some of the same cell wall processes asoccur in diffuse growth, but in a more intense, spatiallylocalized manner. This point is supported by the stun-ted elongation of root hairs in plants with genetic le-sions in expansin genes specifically expressed in roothairs (Yu et al., 2011). Although expansins have beencharacterized largely in the context of diffuse growth,they also are necessary for root hair growth. Leaf tri-chomes in Arabidopsis are fiber-like cells with a conicalshape that arises from highly anisotropic diffusegrowth; its tip-biased gradient in surface expansion hasbeen related to spatial gradients in cell diameter, wallstress, and wall thickness (Yanagisawa et al., 2015).

In this Update, I begin with a review of the biophys-ical basis of cell wall growth, followed by our changingconcepts of the role and interactions of cell wall com-ponents that compose the cell wall, and end with recentinsights from atomic force microscopy (AFM), showingthe details of microfibril organization and motionsduring wall enlargement.

WALL STRESS RELAXATION DRIVESCELL GROWTH

What happens when a cell grows? Cell volume in-creases as a result of water uptake, and wall surfacearea enlarges irreversibly by local separation of cell wallcomponents (Cosgrove, 2016b). The phrase “turgor-driven growth” (or its variants) is often used to de-scribe this process, but this phrasing can be misleadingif it is taken to imply that growth is simply amechanicalstretching of an inherently pliant cell wall, likestretching a piece of putty. To understand this pointbetter, it is useful to distinguish different patterns of cellwall stretching in response to an applied tensile force

Plant Physiol. Vol. 176, 2018 17

Wall Structure, Mechanics, and Growth

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(Fig. 1). Elastic deformation is by definition reversibleand instantaneous upon applied force. Small wall de-formations are typically elastic, whereas after large,rapid, transient deformations, many cell walls do notfully return to their initial size. The irreversible part is aplastic deformation, which occurs when the wall isstretched beyond a yield point. When force is applied toa wall in a sustained manner, the stretch (“strain” inengineering terms) is partly elastic and partly plastic orpartly viscoelastic and partly viscoplastic, dependingon the time scales involved (Boudaoud, 2010; Moulia,2013). These latter two terms refer to time-dependentdeformations. Without sustained cell wall loosening, aconstant force applied to a wall typically results in atime-dependent strain that approaches a nearly steadyvalue in a few minutes, depending on the sample andits strain history (Hohl and Schopfer, 1992). Such de-formation is a result of the polymeric nature of plant cellwalls, but the exact structural basis for thesemechanicalproperties is largely unknown and needs further theo-retical development.

These purely mechanical responses of walls differ inan essential way from the sustainedwall expansion thatoccurs during cell growth (Cosgrove, 2016b; Zhanget al., 2017), which depends on continuous loosening byexpansins or other wall-loosening agents (Cosgrove,2016a). Wall loosening results in wall stress relaxationthat drives cell growth. The significance of wall stressrelaxation for plant growth was first recognized by Rayet al. (1972) and later solidified by detailed theory andexperimental results (Cosgrove, 1993a, 1993b; Ortega,2017). As a result of the complex interactions of cell wallcomponents, there are various distinctive ways inwhich cell walls may become mechanically softer(meaning more easily deformed by mechanical force),but they do not necessarily result in an increase in wallrelaxation and growth, and they therefore do not

qualify as wall loosening processes (see below andTable I). For instance, with rare exceptions (Yuan et al.,2001), lytic enzymes may soften walls, but they do notstimulate cell growth or cause long-term cell wall ex-tension (creep; Ruesink, 1969; Cosgrove and Durachko,1994; Fleming et al., 1997). On the other hand,a-expansins cause stress relaxation and prolonged en-largement of cell walls, but they lack wall lytic activityand they do not soften the wall, as measured by tensiletests (see figure 8 in Yuan et al. [2001]). These are re-markable facts that seem counterintuitive to expecta-tions based on conventional models of cell walls(Carpita and Gibeaut, 1993). To illustrate the point inanother way: Xyloglucan-deficient hypocotyl wallsfrom the xxt1,xxt2 mutant of Arabidopsis are morecompliant (more easily stretched) in tensile tests com-pared with wild-type walls, yet xxt1,xxt2 hypocotylsgrowmore slowly than the wild type (Xiao et al., 2016).Moreover, despite their greater mechanical compliance,the xxt1,xxt2 walls extend more slowly in creep tests,exhibit less stress relaxation, and are less responsive toa-expansins compared with wild-type walls (Park andCosgrove, 2012a). Another example illustrates the flipside of the coin: Low temperature strongly reduced cellwall expansion in Chara cells, but wall elasticity washardly affected (Proseus et al., 2000). Many other ex-amples have been documented where mechanical testsdo not reliably report on the growth properties of thecell wall (Cosgrove, 2016b).

Despite this “inconvenient truth” [apologies to AlGore (2006)], wall elasticity is frequently taken to besynonymous with cell wall growth properties. Some-times this is a matter of convenience—elasticity is rel-atively straightforward to incorporate into simulationsof growth (Fayant et al., 2010; Huang et al., 2015)—andnumerous methods have been devised in recentyears to measure elasticity of tissues and cells with

Figure 1. Schematic comparisons of dif-ferent strain patterns of cell walls (top)stretched with a uniaxial force (bottom). A,Purely elastic (reversible) strain. Cell wallsoften display a retarded elasticity becausewall polymer motions are not instanta-neous. B, A combination of elastic andplastic strain. Beyond the yield point strainis partly elastic and partly plastic. C, Vis-coelastic and viscoplastic strains. Polymermotions require time to reach equilibrium.D,Wall loosening results in sustained time-dependent extension (creep).

18 Plant Physiol. Vol. 176, 2018

Cosgrove



mechanical devices (e.g. Routier-Kierzkowska et al.,2012; Nezhad et al., 2013; Sanati Nezhad et al., 2013;Beauzamy et al., 2015b; Vogler et al., 2015; Mosca et al.,2017). To be fair, there are indeed cases where elasticityroughly correlates with cell growth, as discussed be-low. Such correlation may indicate a change in wallstructure that actually contributes to altered growth byamplifying wall stress relaxation (see Cosgrove[2016b]), or may be entirely coincidental, resulting fromstructural changes independent of wall extensibility.Elastic changes may also be a consequence of the al-tered growth. For instance, it has long been known thatauxin treatment results in increased wall compliancesinmany cases (Heyn, 1932; Edelmann andKohler, 1995;Braybrook, 2017). However, the onset of auxin-inducedgrowth is fast and precedes the gradual change in wallmechanics (Cleland, 1984), which may reflect longer-term changes in wall structure induced by auxin orfaster growth itself.Another factor in the confusion between elasticity

and extensibility may be the fuzzy definition of termssuch as wall loosening, softening, and weakening (Ta-ble I). We do not have an established vocabulary todistinguish the many facets of wall properties. As dis-cussed here, wall loosening refers to a shift or cut of aload-bearing part of the wall, relaxing tensile stress inthe whole wall and simultaneously reducing cell tur-gor, which is the Newtonian counterbalance to wallstress. The reduction in turgor enables passive wateruptake by osmosis, which elastically stretches the cellwall, restoring turgor. Such loosening-dependent pol-ymer movement might be called a chemorheologicalflow (Dumais, 2013; Moulia, 2013; Cosgrove, 2016b).However, this term usually implies a chemical changein covalent bondingwithin the cell wall, which does notappear to be necessary for cell wall creep, i.e.a-expansin facilitates cell wall creep without evidenceof hydrolysis or other covalent modification of wallpolymers (Cosgrove, 2015, 2016a). I use wall “soften-ing” to denote a process that changes wall stiffness,without the implication that the wall can grow morequickly. I use wall “weakening” to refer to cases wherethe wall breaking strength is reduced. Thus, xxt1,xxt2hypocotyls are weaker than the wild type because theirbreaking strength is reduced (Cavalier et al., 2008).Despite being softer and weaker, xxt1,xxt2 hypocotyls

are less extensible in assays of growth and cell wallcreep. The key conclusion is that elasticity reports onwall structure, not the dynamic relaxation processesthat determinewall extensibility and that drive cell wallgrowth. With the definitions used here, a stretchyrubber band has significant elasticity but no extensi-bility.

CELL WALL MODELS NEED FURTHER REFINEMENTAND TESTING

This physical framework, in which wall stress relax-ation initiates cell growth and couples water uptakewith wall expansion, leaves numerous molecular de-tails unresolved. What is the molecular nature of wallloosening? Which wall components of the wall are thetargets of wall loosening, and which components limitthe ensuing polymer motions (e.g. spreading of cellu-lose microfibrils)? Are there multiple ways to inducewall relaxation and loosen the wall? What are the mo-lecular bases of wall elasticity and plasticity?

The answers to these questions require an accuratemodel of cell wall structure and a deep understandingof the molecular basis of wall enlargement. Notwith-standing textbook models, this remains a major unmetchallenge. For many years, xyloglucan was imagined tofunction as a load-bearing tether linking well-separatedcellulose microfibrils (Carpita and Gibeaut, 1993;Albersheim et al., 2011), with pectins functioning as acompliant, hydrated, gel-like matrix between the mi-crofibrils. This concept entailed predictions of wallmechanics that were largely untested. Recent results,however, call for reevaluation of the roles of cellulose,xyloglucan, and pectins in wall structure and growth(Cosgrove, 2016b). For instance, extensive xylogluca-nase treatment of isolated cell walls from cucumber(Cucumis sativus) and Arabidopsis did not induce cellwall extension (creep) or increase mechanical compli-ances (stress/strain behavior in tensile tests), eventhough at least half of the xyloglucan was removed(Park and Cosgrove, 2012b). On the other hand, lowconcentrations of bifunctional endoglucanases (able tocut both xyloglucan and cellulose) induced cell wallcreep and mechanical softening (increase in tensilecompliances). The extent of wall hydrolysis needed toinduce these biomechanical actions was very small.

Table I. Brief explanation of terms related to wall mechanics and growth properties, as used here

Term Meaning

Extensibility General term for the ability of the cell wall to grow; in other contexts, this is the coefficient relating growth rate to turgorpressure (Cosgrove, 1993b); not elasticity and not a purely mechanical property, as it depends on wall loosening

Loosening Molecular process causing wall stress relaxation, resulting in water uptake and cell growth; it confers irreversibility to wallstrains; expansins are well-recognized as wall-loosening agents

Softening A process that makes the wall more deformable to mechanical forceWeakening A process that reduces the force or energy need to break wallsElasticity A measure of how readily the cell wall changes shape in response to a transient mechanical forcePlasticity A measure of the irreversible component of wall deformation in response to a transient mechanical forceCompliance The slope for strain/stress curves; it is the reciprocal of modulus or stiffness





Enigmatically, the combination of a xyloglucan-specificendoglucanase with a cellulose-specific endoglucanasedid not mimic the bifunctional enzymes in their actionon cell wall creep. To explain this enigma, we proposedthat the bifunctional enzymes hydrolyze relatively in-accessible load-bearing junctions between cellulosemicrofibrils where xyloglucan and noncrystalline cel-lulose are entwined (Fig. 2). Digestion of these nexuspoints, dubbed “biomechanical hotspots,” releasessome of the load-bearing junctions between the micro-fibrils, enabling stress relaxation and irreversible mi-crofibril movements. Limited accessibility and kineticsaccount for the failure of combined xyloglucan-specificand cellulose-specific enzymes to cause cell wall creep.

The origin of these proposed junctions is uncertain.One possibility is that they form spontaneously byphysical entrapment and self-assembly as cellulose andxyloglucan are deposited to the cell surface. Coordi-nation of vesicle secretion with cellulose synthesismight facilitate their formation. Another possibility isthat hotspots are formed enzymatically. Recentreports have identified enzymes (xyloglucan endo-transglucosylase/hydrolase) that can cut noncrystal-line cellulose and ligate xyloglucan onto the celluloseend (Hrmova et al., 2007; Simmons et al., 2015;Shinohara et al., 2017). The specific activity of theseenzymes in performing such hetero-transglucosylations

is very low, so the biological significance of such reac-tions is uncertain. Further work is needed to uncover theorigin of biomechanical hotspots.

The hotspot concept gains support from a study thatused a novel solid-state NMR strategy to characterizethe target of expansin binding in complex walls fromArabidopsis seedlings (Wang et al., 2013). With use of asensitivity-boosting technique called dipolar nuclearpolarization to enhance 1H-13C cross-polarization, nu-clear magnetic spins were selectively filtered through15N,13C-labeled expansin protein to 13C in wall com-ponents within close proximity to expansin. The re-sults showed that expansin binds to cellulose with aslightly different chemical shift than the bulk of thecellulose, indicating cellulose chains that are packedtogether differently than most of the cellulose. More-over, there was evidence of xyloglucan in close prox-imity to expansin. These NMR characteristics resemblethose that might be expected for the biomechanicaljunctions targeted by wall-loosening bifunctionalendoglucanases (Park and Cosgrove, 2012b). Addi-tional NMR characterization found that the expansin-binding sites are on the surface of cellulose microfibrils,yet are relatively distant from water, consistent withlimited accessibility (Wang et al., 2016b).

The concept of wall structure emerging from theseresults emphasizes the importance of direct connec-tivity between cellulose microfibrils, as opposed toprevious concepts where microfibrils were depictedas well separated by matrix polymers and connectedonly via tethers. As described below, AFM reveals theorganization of cellulose microfibrils to be a networkof laterally bundled microfibrils rather than a collec-tion of well-spaced microfibrils connected only bymatrix, as traditionally depicted in textbook modelsof primary cell walls.

Another cellulose-related topic emerging from recentwork is the potential significance of the cross-sectionalshape of cellulose microfibrils, which influences theproportion of hydrophilic and hydrophobic faces on themicrofibril (Newman et al., 2013; Cosgrove, 2014;Wangand Hong, 2016). This issue is important for cell wallmodels because of the potential for these two faces tointeract differently with matrix components. The hy-drophilic faces of cellulose microfibrils are populatedby hydroxyl groups extending from the sides of the Glcresidues, whereas the hydrophobic faces are those thatexpose the Glc rings with their nonpolar 2CH groups.X-ray crystallography studies show that expansins bindthe hydrophobic face of cellulose chains (Georgeliset al., 2012). Computational studies indicate that xylo-glucan likewise preferentially binds to the hydrophobicface of cellulose (Zhao et al., 2014). This computationalresult is supported by a recent field emission scanningelectron microscopy (FESEM)-based study showingthat xyloglucan indeed covers the hydrophobic faces ofcellulose microfibrils in onion (Allium cepa) cell walls(Zheng et al., 2017b). Complementing these results,recent experimental and computational studies indicatethat substituted xylans in secondary cell wallsmay bind

Figure 2. Conceptual depiction of structural features of primary cellwalls. Cellulose microfibrils are represented as thick rods with hydro-phobic (blue) and hydrophilic faces (orange). Xyloglucan (green) isfound in solvated, coiled conformations and in extended conformationsbound to the hydrophobic faces of cellulose, based on Zheng et al.(2017b). It is also depicted as entrapped between microfibrils. Pectins(yellow) are represented as coiled structures that fill the space betweenmicrofibrils and bind to the hydrophilic surfaces (based on solid-stateNMR results). Microfibrils are bundled by direct contacts and at junc-tions where cellulose and xyloglucan intertwine. The red arrows pointto cellulose-xyloglucan-cellulose junctions that are sites of wall loos-ening by bifunctional endoglucanases. This depiction is a synthesisbased on the most recent results from AFM, FESEM, solid-state NMR,and mechanics.


Cosgrove



the hydrophilic faces of cellulose (Simmons et al., 2016;Grantham et al., 2017; Pereira et al., 2017). Primary cellwalls generally contain only small amounts of xylan,e.g. Zablackis et al. (1995), with the notable exception ofcell walls from grass species (Carpita, 1996). Never-theless, a recent study characterized a xylan in primarywalls of Arabidopsis (Mortimer et al., 2015) that mightselectively bind to the hydrophilic faces of microfibrils.Because b-expansin was recently shown to target xy-lans in grass cell walls (Wang et al., 2016a), it seemsplausible that a- and b-expansins may loosen the con-nections between different faces of cellulosemicrofibrilsin the wall.Direct cellulose-cellulose contacts may be prevalent

in primary walls, but whether such interactions aremediated via their hydrophilic or hydrophobic faces isuncertain and indeed may differ for primary and sec-ondary cell walls. Some studies propose that microfi-brils aggregate via contacts of their hydrophilic faces(Ding et al., 2012; Oehme et al., 2015), whereas an AFManalysis of microfibril patterns in onion cell walls sug-gested that microfibril bundling occurs via the hydro-phobic faces of microfibrils (Zhang et al., 2016). InArabidopsis hypocotyls of the xyloglucan-deficientxxt1,xxt2 mutant (Xiao et al., 2016), cellulose microfi-brils were more aligned and closely packed than in thewild type, an indication that xyloglucan may promotedispersion of microfibrils within a lamella. It is alsopossible that these changes in cellulose organization area consequence of CWI responses to wall defects.

PECTINS

As the idea of a direct cellulose network within pri-mary cell walls has gained support, pectins have alsoattracted new attention as potential modifiers of wallenlargement. The case has been most cogently arguedfor pollen tubes (Parre and Geitmann, 2005; Rojaset al., 2011; Sanati Nezhad et al., 2014), where pectinsdominate wall structure, and for the giant-celled algaChara corallina (Boyer, 2016). In Arabidopsis cell walls,results from multidimensional solid-state NMR indicateextensive noncovalent cellulose-pectin interactions (Wanget al., 2012, 2015). This is surprising because such inter-actions are not observed in binding studies in vitro(Zykwinska et al., 2008a, 2008b). A recent study of cellwall properties along the axial growth gradient of theArabidopsis stem gives additional clues (Phyo et al.,2017). Pectins in the apical (faster-growing and softer)region of the stem are more mobile, more hydrated, moreesterified, and more branched compared with pectins inthe lower (more slowly growing and stiffer) region of thestem. How these correlated structural changes in pectinproperties influence pectin-cellulose interactions, cell wallmechanics, and growth needs further testing. In contrast,a very different concept of pectinwas proposed in a studythat characterized a proteoglycan with covalently linkedpectin and xylan domains (Tan et al., 2013). This structureis reminiscent of the early macromolecular model of

primary cell walls by Keegstra et al. (1973). The potentialrole of such proteoglycans in wall structure, mechanics,and growth remains to be evaluated.

In another vein, a series of elegant experiments withclear but perplexing results implicate localized de-esterification of homogalacturonan as a signature eventin the auxin-induced patterning of the shoot apicalmeristem of Arabidopsis, resulting in elastically softerregions of the meristem surface, as measured bymicroindentation techniques, where leaf primordiaemerge (Peaucelle et al., 2008, 2011; Braybrook andPeaucelle, 2013). Experiments with the auxin-transportpin1mutant and genes encoding pectin methyl esteraseinhibitor proteins suggest that auxin patterning of theshoot apical meristem requires pectin de-esterification.

The correlation of de-esterified pectin with softermeristem regions is perplexing because, in the broadercontext of cell wall properties, pectin de-esterification iscommonly thought to result in stiffer, not softer walls,as a result of increased ability for calcium-mediatedcross-linking of homogalacturonan. For instance, in thehemispherical tip of growing pollen tubes, de-esterifiedhomogalacturonan is associated with stiffer wallsand cessation of wall expansion (Geitmann and Parre,2004; Sanati Nezhad et al., 2014). Likewise, pectinde-esterification is associated with the decline in thegrowth rate and increased wall stiffness along theapical-to-basal gradient of growing stems (Goldberget al., 1986; Phyo et al., 2017). Moreover, wall looseningby expansin is hindered in the basal regions of growingstems where the extent of de-esterified pectin is high(Cosgrove, 1996), and this hindrance may be partiallyreversed by removal of pectins and calcium (Zhao et al.,2008). Hocq et al. (2017) have questioned the concept ofwall stiffening by calcium cross-linking of pectins, yetolder results with calcium chelators yielded a nuancedconclusion: that calcium cross-links are indeed load-bearing but are not broken during acid-induced(expansin-mediated) wall loosening (Virk and Cleland,1990). Likewise, imaging with AFM shows that additionand removal of calcium reversibly stiffens pectins on thesurface of onion epidermal cell walls (Zhang et al., 2016),yet parallel experiments show little effect of calcium onwall extension in vitro. Thus, more research is needed toclarify the perplexing observations about pectin esteri-fication, calcium cross-linking, wall softening, auxin re-sponses, and cell growth in the meristem. Are CWIresponses complicating this story?

The perplexing results about pectin esterificationreported for the shoot apical meristem raise the fol-lowing question: Is cell wall enlargement in the meri-stem regulated in the sameway as in subapical zones ofrapid cell enlargement, or do walls in the meristemfollow a different set of rules? Indeed, we know ratherlittle about cell wall structure and wall extensibility inmeristematic regions, other than the limited informa-tion inferred from immunohistochemistry (Yang et al.,2016) and osmomechanical probing of cell elasticity(Kierzkowski et al., 2012; Nakayama et al., 2012;Routier-Kierzkowska and Smith, 2013). In a recent





multifaceted study of the swollen shoot apical meri-stems from the clavata3-2 mutant of Arabidopsis, wallcomposition was found to consist of approximately30% cellulose, approximately 26% pectin, and 15%xyloglucan (Yang et al., 2016). This is unremarkable, asit is similar to the wall composition of whole Arabi-dopsis seedlings (White et al., 2014). Older studies alsobear on the question of cell enlargement mechanisms inmeristems: Local application of a-expansin proteinto the surface of the shoot apical meristem resultedin an outgrowth resembling early stages of a leafprimordium (Fleming et al., 1999), and more pro-nounced outgrowth resulted from transient induc-tion of an a-expansin on the flanks of the meristem(Pien et al., 2001), indicating cell walls in the meri-stem are sensitive to the loosening action ofa-expansin. Moreover, a-expansin is endogenouslyexpressed at the site of incipient leaf primordia be-fore primordium outgrowth (Reinhardt et al., 1998),evidence that modulation of cell wall enlargement issimilar to that of other plant tissues. Allowing for thehigh frequency of dividing cells in meristems and theunique wall synthesis machinery involved in cellplate formation during cell division (Gu et al., 2016),current results indicate meristem walls are similarin composition and growth mechanisms as docu-mented in rapidly elongating cells that emergefrom meristems. Thus, the enigmatic function ofpectin de-esterification in the meristem remains anopen question.

INSIGHTS FROM AFM OF EPIDERMAL CELL WALLS

Plant developmental biologists are probably mostfamiliar with AFM from studies that have used thedevice to indent the surface of growing tissues toevaluate local stiffness (e.g. Peaucelle et al., 2011; Milaniet al., 2014; Sampathkumar et al., 2014a). This is acomplex topic beyond the scope of this review, butreaders are referred to reviews that assess the variedapproaches and interpretations of these stiffness mea-surements (Milani et al., 2013; Mosca et al., 2017). AFM-based stiffness maps of the shoot apical meristem havebeen compared with maps of cell shape, cell division,auxin flow, gene expression, and cytoskeletal patterns(Nakayama et al., 2012; Robinson et al., 2013; de Reuilleet al., 2014; Sassi et al., 2014). These studies contribute tosophisticated models of meristem morphogenesis andphyllotaxis in which wall stress, mechanics, enlarge-ment, and the cytoskeleton play interacting roles(Nakayama et al., 2012; Kierzkowski et al., 2013;Sampathkumar et al., 2014b), concepts rooted in thepioneering efforts of Paul Green to understand the bi-ophysics of meristem dynamics (Green et al., 1996).

AFM can also be used to image cell wall surfaces atsufficiently high resolution to detect individual cellu-lose microfibrils (approximately 3 nm diameter; Fig. 3,A and B), revealing cellulose organization in unprec-edented detail (Zhang et al., 2014). Note that fluorescence

microscopy, even super resolution versions, lacks theresolution needed to see individual microfibrils in thecomplex and dense fibrillar network of cell walls.

Figure 3. AFM images showing the arrangement of cellulose microfi-brils and matrix on the inner surface of the periclinal wall of onionepidermis. A, Large-scale (2 3 2 mm) peak force error map showingmicrofibril bundling and cross-lamellate organization of microfibrils.Image adapted from Zhang et al. (2014). B, Close-up of boxed region inA. Single microfibrils are seen to bundle into groups of two of more.Some microfibril details are obscured by relatively stiff matrix. C, Two-color merged image based on a height map (red), which highlightsmicrofibrils, and a modulus map (green), which indicates stiffness (re-sistance to indentation). The area outlined in white contains regions ofstiff matrix (bright green) closely associated with microfibrils. Othermatrix regions are dark, indicating they are soft. D, Two-color mergedimage based on a height map (red, predominantly microfibrils) and adeformation map (green, predominantly matrix). Regions of soft matrixbetween microfibrils are obvious in this image. Images adapted fromZhang et al. (2016). E, Modulus map of onion wall in a relaxed state(small axial force); the predominance of blue color indicates low re-sistance to indentation. F, Modulus map of the same wall region uponapplication of axial force, stretching the wall in the direction indicatedby the arrow. The predominance of red shows that microfibrils havebeen pulled taut by the axial force, indicating they are load bearing, asare matrix components, though to a lesser extent. Images adapted fromZhang et al. (2017).


Cosgrove



Transmission electron microscopy generally gives lim-ited information about microfibril organization in cellwalls, with the exception of the replica/shadowingmethod (McCann et al., 1990), which has been super-seded by FESEM. FESEM has excellent resolving powerfor microfibril detection (Fujita and Wasteneys, 2014;Zheng et al., 2017a), but to attain high resolution, thewall must be dehydrated, which means wall polymersmay become distorted or shift position as the water isremoved. In contrast, AFM can be carried out underwater, allowing imaging of walls in a near-native state.The surface topology can be measured with nm reso-lution and simultaneously probed mechanically tomeasure an indentation modulus (resistance to surfacedeformation).We used AFM to characterize cellulose microfibril

organization of the outer (periclinal) wall of onionscales (the fleshy leaf of the bulb). Epidermal peels fromonion scales have been used in numerous studies toconnect tissue-level mechanics, growth, and net cellu-lose orientation (Wilson et al., 2000; Hepworth andBruce, 2004; Suslov et al., 2009; Beauzamy et al.,2015a). In previous studies, whole epidermal layerswere prepared with intact (living) cells. We devel-oped an alternative procedure to prepare epidermalstrips in which the outer (periclinal) wall tears awayfrom the rest of the cell. This exposes the inner (mostrecently synthesized) surface for imaging by AFMand provides a simpler material for mechanicalstudies (a sheet of outer epidermal walls rather thana layer of intact turgid cells with complex architec-ture).Like the outer epidermal walls in many plant organs,

including Arabidopsis hypocotyls (Crowell et al., 2011),the epidermal wall in onion is thick (2 or moremm, depending on which scale is used) and has a cross-lamellate construction. In AFM images, individual mi-crofibrils in the surface lamella are seen to form a networkwith single microfibrils merging into and out of bundledregions where two or more microfibrils are laterallyaligned and in close contact (Fig. 3, A and B; Zhang et al.,2016). Microfibrils are arranged roughly in a commondirection in each lamella, andmicrofibril orientation shiftsabruptly by 30° to 90° between adjacent lamellae, pro-ducing a wall comprised of many, highly anisotropic la-mellae in a wide range of orientations. The result of thiscross-lamellate structure is that the whole wall has muchweaker net structural anisotropy than the individual la-mellae that make up the wall. By combining height mapswith modulus maps and deformation maps, we can vi-sualize microfibrils in the context of rigid and soft regionsof thematrix (Fig. 3, C andD).Moreover, by stretching thewall, we can detect microfibril motions and detect whichcomponents become more resistant to indentation, anindication that they bear some of the tensile force (Fig. 3, Eand F).This cross-lamellate construction has important im-

plications for the patterns of microfibril separationduring cell growth. It is commonly accepted that cellwalls expand preferentially in the direction at right

angle to the net direction of cellulose alignment (Baskin,2005; Suslov et al., 2009). Multicellular tissues presentadditional structural complications beyond the scope ofthis review (Crowell et al., 2010; Baskin and Jensen,2013). For a cell wall with net transverse orientation, it is

Figure 4. Predicted patterns of microfibril movement in a cross-lamellatecell wall stretched uniaxially. Shown is cellulose alignment in single lamellaebefore (left) and after (right) stretching in the vertical direction. Axial elon-gation is accompanied by transverse shrinkage during elastic uniaxial ex-tensions. A, In lamellae with cellulose oriented transverse to the direction ofstretch, the axial distance between microfibrils will increase and cellulosemicrofibrils will bend or kink in the transverse direction. B, For lamellaewithcellulose oriented at approximately 45°, microfibril angle will shift in theaxial direction and distance between microfibrils will be reduced. C, Forlamellae with cellulose oriented in the same direction as the axial stretch,microfibrils will become straighter, more closely packed, and will undergoside-by-side sliding. Figure is adapted from Zhang et al. (2016).





easy to imagine that the distance between microfibrilsincreases as the cell elongates axially. This is com-monly illustrated by analogy with the way a woundspring elongates (like a Slinky toy). But what happensin a cross-lamellate wall where cellulose is organizedin lamellae that are aligned in many orientations, ax-ially, transversely, and at a spectrum of angles be-tween these two orthogonal directions? Specifically,how do microfibrils move in lamellae where cellulosemicrofibrils are aligned parallel to the direction ofmaximal growth? In considering this question, I as-sume that microfibrils are too stiff to stretch appre-ciably and too strong to break, and also assume thatadjacent lamellae do not slip past each other. Withthese assumptions, it seems that microfibrils in theseaxially aligned lamellae must have a mechanism ofside-by-side gliding or axial shearing (Fig. 4). If axialshearing of this type requires more force than lateralseparation of microfibrils in transversely aligned la-mellae, then axial shearing between microfibrilsaligned in the direction of cell growth may limit therate of cell wall enlargement. This reasoning puts thefocus of attention on themolecular nature of the lateralassociations between microfibrils and the patterns ofmicrofibril motions during wall expansion. We knowrather little about this aspect of primary cell wallgrowth.

These inferences about microfibril movements gainsupport from a recent study in which the nanoscalemovements of cellulose microfibrils were directlymonitored by AFM (Zhang et al., 2017). The cell wallwas extended in a well-defined series of extensionsthat included elastic and plastic strains imposed byaxial force as well as time-dependent creep inducedby treatment with an endoglucanase with wall-loosening activity. For lamellae in which the micro-fibrils were oriented at 30° to 60° off the axis ofapplied tensile force, mechanical stretching resultedin passive reorientation of microfibrils in the direc-tion of stretch, as predicted by multinet growthmodels (Preston, 1982). In addition, examples of axialshearing and lateral separation were observed, pro-viding direct evidence for the microfibril movementsinferred above from general considerations. Whencell wall creep was induced by application of awall-loosening endoglucanase, remarkably differentpatterns of microfibril movements were observedcompared with those during elastic and plastic de-formations. This difference was attributed to changesin microfibril connectivity, i.e. selective looseningat hotspots by endoglucanase action. Thus, thestretching of a wound spring does not seem an aptanalogy for how microfibrils move during cell wallgrowth.

This AFM study documents an example wheremicrofibril movements motivated by applied forcediffered from those that occurred when wall expan-sion was induced by wall loosening. The microfibrilsin differently aligned lamellae displayed the largediversity of microfibril movements that are required

for enlargement of cross-lamellate walls. It is possiblethat different patterns of microfibril movement in-volve different matrix components, e.g. xyloglucans,xylans, pectins, or even water alone, and are medi-ated by different wall-loosening agents.

PERSPECTIVE

As new tools and new approaches have been appliedto investigate cell growth, it has become evident thatour conventional model of the growing cell wall fallsshort. Xyloglucan seems to have rather different func-tional roles than those hypothesized in conventionaldepictions of the growing cell wall of the past 40 years.New evidence for the role of pectins in wall structureand morphogenesis are tantalizing, yet raise newquestions and the biological responses evoked by CWIsensors complicate the interpretation of cell wall mu-tants. Finally, the new appreciation of the complexity ofcellulose organization in the growing wall presentsopportunities for rethinking the molecular control ofdiffuse growth. Experimental systems that enablestudies of both pectin and cellulose networks areneeded for future integration of these emerging ideas.Received October 25, 2017; accepted November 13, 2017; published November14, 2017.


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http://dx.doi.org/10.1111/tpj.13778

http://dx.doi.org/10.1111/tpj.13778


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