1. Driving growth: turgor pressure, the Lockhart equation and the cell wall
In plants (as in bacteria and fungi), osmotic water uptake causes the cells to become turgid. The internal pressure, or turgor pressure, makes the cells swell and it is only the presence of a stiff exoskeleton, the cell wall, that prevents them from bursting. In growing plant cells, cell wall synthesis and remodeling causes the external matrix to yield to the turgor pressure, thus allowing the cell to grow. Turgor pressure is thus often considered as the motor of growth.
Pfeffer was the first to measure osmotic pressure indirectly (Pfeffer, 1877). In the 1960s, Paul Green developed the first direct method to measure turgor pressure in cells, which would become the precursor of modern pressure probes (Green & Stanton, 1967; Green, 1968): an oil-filled microcapillary, which is connected to a pressure sensor and a moveable plunger, is introduced into a cell and the pressure necessary to maintain the liquid in the cell is measured as a quantification of internal turgor pressure (Husken et al., 1978; Cosgrove & Durachko, 1986). Based on these measurements, it has been estimated that turgor pressure in plant cells can build up to 10 bar. This value is extremely high, considering for example that 0.03 bar corresponds to a high blood pressure, and explains why plants are able to grow and even push their way through rocks or concrete.
Mathematically, the concept of growth driven by turgor was qualitatively formulated in 1877, and quantitatively summarized in 1965 by the Lockhart model (Pfeffer, 1877; Lockhart, 1965; Ortega, 1989; Tomos et al., 1989; Moulia & Fournier, 2009), which equates the expansion rate of a cell (dV/Vdt) to m(P − Y), where V is the volume of the cell, t is time, m is the extensibility of the cell wall, P is the turgor pressure, and Y is the minimal threshold of P below which the cell will not grow. This equation stresses the importance of the rheological properties of the cell wall in growth control. In the following we will, therefore, review its main features.
2. Defining the fundamental growth parameters at the cellular level: the role of the cell wall
Figure 1. Anisotropy is controlled by the cortical microtubules. (a) Three-dimensional reconstructions of GFP-LTI6b meristems 72 h after a treatment with oryzalin: GFP-LTI6b meristems are imaged using confocal microscopy, and a projection of the cell surface is generated using the merryproj software (INRA-INRIA, France) (de Reuille et al., 2005). Note the formation of spherical primordia (i.e. a loss of anisotropy) and the presence of differential growth rates between the center of the meristem (lower growth) and the primordia (faster growth). Bar, 20 μm. (b) Close-up from (a). The shape of individual cell can be tracked and analysed. In the presence of oryzalin, walls tend to meet at 120° and cells tend to become hexagonal. Owing to the differential growth rates between the different domains of the shoot apex, each cell, being glued to its neighbors, displays some degree of anisotropic growth, even in the absence of microtubules. (c) Turgor pressure is the motor of growth. If the microtubules are oriented in a transverse orientation, the deposition of cellulose microfibrils should follow the same pattern and prevent wall expansion in a direction parallel to the microfibrils (i.e. promote a direction of growth in the axis of the cylinder created by the spirals of cellulose). If the orientation of the microtubules is more random, growth becomes isotropic. (d) A model: the cortical microtubules (CMT) located on the cytoplasmic side of the plasma membrane (PM) guide the movement of the rosette (CSC, cellulose synthesis complex) which synthesizes the cellulose microfibrils (CMF) in the cell wall, in a direction parallel to that of the CMT. (Reprinted from Emons et al. (2007) with permission from Elsevier.)
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During growth, cellulose is deposited outside the plasma membrane, thus preventing the wall from becoming thinner and weaker. Because of their high tensile strength, cellulose microfibrils are the main determinants of cell wall stiffness. In addition, being structurally filamentous and often aligned in parallel arrays, cellulose microfibrils are also the main determinants of cell wall anisotropy: cells hardly grow in the direction parallel to the spirals of cellulose (Lloyd & Chan, 2004; Baskin, 2005; Cosgrove, 2005; Marga et al., 2005; Kutschera & Niklas, 2007; see next sections).
Between the microfibrils, two classes of polysaccharides, pectins and hemicelluloses, largely constitute the so-called cell wall ‘matrix’ together with structural proteins. Contrary to cellulose microfibrils, hemicelluloses are branched and this structural feature, together with their ability to bind cellulose, is thought to link the cellulose microfibrils into a cellulose–hemicellulose network (Somerville et al., 2004; Cosgrove, 2005). Other models envision hemicelluloses as spacers that prevent microfibrils aggregation (Thompson, 2005). Pectins control the stiffness of the cell wall, allowing or preventing the sliding of microfibrils (Willats et al., 2001; Cosgrove, 2005). More generally, through its noncovalent interactions with cellulose microfibrils, the matrix contributes to the mechanical properties and the dynamics of the cell wall.
While matrix polysaccharides are synthesized in the Golgi apparatus before being integrated into the cell wall, cellulose microfibrils are synthesized by cellulose synthases (CESA) located at the plasma membrane. Contrary to matrix polysaccharides which can diffuse into the cell wall, the newly synthesized microfibrils remain at the inner wall (Ray, 1967). The CESA proteins assemble into hexameric complexes forming particle ‘rosettes’ at the plasma membrane (as observed in the electron microscope after cryofracture). In Arabidopsis, the synthesis of the primary cell wall requires CESA1, CESA3 and CESA6 (Arioli et al., 1998; Fagard et al., 2000). Each CESA produces (1,4)-β-d-glucan chains that spontaneously bundle into a 4 nm wide microfibril. Each microfibril is long enough to run along the cell circumference several times (Cosgrove, 2005).
The turgor driven cell wall expansion is irreversible, and involves a slow reorganization of the microfibrils and matrix (also called creep). Schematically, the matrix loosens allowing the microfibrils to slide more easily and causing the wall to yield to forces generated by the internal pressure. Turgor pressure thus provides the mechanical energy for this polymer motion, while the loosening of the linkages between microfibrils controls the direction and rate of deformation. Several classes of molecules, namely expansin, pectin methylesterase, xyloglucan endotransglycolase/hydrolase, endo-(1,4)-β-d-glucanase and reactive oxygen species have been implicated in wall loosening. In the following paragraphs, we will discuss this topic more in detail.
3. Wall loosening and growth rate
Expansins are pH-dependent wall loosening proteins. When exposed to expansin or when expansin genes are overexpressed, the cell growth rate increases (Fleming et al., 1997; Pien et al., 2001). Conversely, reduction of expansin expression decreases cell growth (Cho & Cosgrove, 2000; Choi et al., 2003). Last, the expansin expression pattern matches and predicts the growth pattern (Cho & Kende, 1997; Reinhardt et al., 1998). Mechanistically, expansins do not hydrolyse wall polymers but are believed to weaken hydrogen bonds between wall polysaccharides, thus relaxing wall stresses and allowing growth (McQueen-Mason & Cosgrove, 1994, 1995; Cosgrove, 2005).
The role of pectin methyl esterase (PME) in cell growth has recently received increased attention. Pectin is thought to represent one-third of all primary wall macromolecules (Willats et al., 2001). A defining aspect of pectins is that they are rich in galacturonic acid. When pectins are deposited in the cell wall, most (70–80%) of it is methylesterified (Ridley et al., 2001). The PMEs remove the methyl-ester groups resulting in stretches of acidic residues that can bind calcium and cross-link with other pectic chains. In this scenario, consistent with data obtained in pollen tube and hypocotyls, PMEs render the cell wall stiffer, leading to reduced growth (Jiang et al., 2005; Derbyshire et al., 2007; Rockel et al., 2008). By contrast, in the absence of calcium, de-methyl-esterification by PMEs is supposed to render the cell wall more fluid. De-methyl-esterification is also a prerequisite for pectin degradation by pectate lyase, which could further loosen the cell wall by opening an access to expansins and other hydrolytic enzymes in the cell wall (Cosgrove, 1999). The PMEs thus act as enzymes that make the cell wall able to become more or less stiff, depending on the action and presence of other effectors. To add a layer of complexity, PMEs belong to a large multigenic family and each isoform seems to have differences in their activities (Willats et al., 2001). Recent data are consistent with a loosening role of PMEs during the process of organ initiation in the shoot apical meristem. Notably, primordia position at the periphery of the shoot meristem correlates with the presence of de-methyl-esterified pectins, and primordium formation is inhibited when pectin de-methyl-esterification is impaired (Peaucelle et al., 2008).
Xyloglucans are the major hemicelluloses of primary cell walls (10–20% of the cell wall dry weight; Cavalier et al., 2008). Xyloglucans are the target of hydrolytic enzymes, namely the xylogucan endotransglucosylase/hydrolase (XTH). These have been implicated in both cell wall loosening and cell wall strengthening (Antosiewicz et al., 1997; Takeda et al., 2002; Cosgrove, 2005). Recently, using a reverse genetics approach, Cavalier and coworkers (Cavalier et al., 2008) obtained Arabidopsis lines with no detectable xylogucans. Results from micromechanical stress tests on these lines revealed a significant decrease in stiffness consistent with the contribution of xyloglucans to the mechanical properties of the primary cell wall. Surprisingly, the absence of detectable xyloglucans did not significantly alter the plant phenotype, thus challenging our understanding of the role of xyloglucans in the cell wall and more generally in growth and morphogenesis (Cavalier et al., 2008).
Xyloglucans and cellulose are also putative targets of endo-(1,4)-β-d-glucanases, also called cellulases. Three of them, including KORRIGAN, are membrane-bound and have rather been involved in cellulose formation (Nicol et al., 1998). Overexpression of PopCel1, a secreted endo-(1,4)-β-d-glucanases from poplar, in Arabidopsis increased wall extensibility and growth. Conversely, the reduction of endoglucanase expression resulted in reduced growth (Ohmiya et al., 1995, 2000; Tsabary et al., 2003). Although these enzymes appear to be important for wall loosening, their role in development is still poorly documented (Cosgrove, 2005).
In the past 10 yr, reactive oxygen species (ROS) have emerged as major wall loosening agents (Foreman et al., 2003; Liszkay et al., 2003; Schopfer & Liszkay, 2006). Contrary to the proteins presented so far, ROS do not display substrate specificity and can cleave any wall polysaccharides. Their effect on wall extension, directly by polysaccharide cleavage or indirectly through activation of signaling pathways, has been demonstrated in vitro and observed in vivo in actively growing tissues (Schopfer & Liszkay, 2006). Reactive oxygen species are produced by the NAD(P)H oxidase and cell wall peroxidase (Foreman et al., 2003). Interestingly, production of ROS, and their role in cell elongation, is auxin dependent (Schopfer, 2001; Schopfer et al., 2002) and might be subject to spatial regulation.
4. Cell wall anisotropy and the microtubular cytoskeleton
Turgor pressure is nondirectional, but the presence of aligned cellulose microfibrils in the cell wall translates turgor into anisotropic growth. Anisotropy is defined as the ratio of the two principal rates of growth (strain rates). Unless otherwise stated in the text, anisotropy will refer to the mechanical properties of the cell wall. Although the many determinants of cell wall loosening are still far from being completely characterized, the control of cell wall anisotropy is relatively well established and, in the most widely accepted concept, relies mainly on a single factor: the cortical microtubular cytoskeleton (Fig. 1). The morphogenetic role of the other major cytoskeletal element, the actin network, seems to be rather unspecific and/or indirect in most cells, although actin is crucial for the morphogenesis and development of cells undergoing tip growth (i.e. pollen tubes and root hairs).
Using the green alga Nitella, Paul Green first showed that colchicine disrupts cellulose alignment in the cell wall leading to isotropic (spherical) growth (Green, 1962). The major target of this drug turned out to be the cortical microtubules, which are attached to the plasma membrane in highly ordered arrays seemingly parallel to the cellulose microfibrils. Almost since their discovery, plant microtubule orientation has therefore been associated with the orientation of the cellulose microfibrils (Ledbetter & Porter, 1963; Emons et al., 2007). After the complex of cellulose synthesis, the CESA complex (also called the rosette), was located at the plasma membrane, it was hypothesized that cortical microtubules guide the movement of the rosette, thus explaining the colinearity between microtubules and microfibrils in growing cells (Heath, 1974; Mueller & Brown, 1980; Kimura et al., 1999; Fig. 1c,d). In this scenario, the polymerization of the stiff cellulose microfibrils would provide the force necessary for the movement of the rosettes, driving them forward. Using fluorescently labeled microtubules and cellulose synthase, Paredez and coworkers observed this movement in vivo and demonstrated that cortical microtubules guide the deposition of cellulose microfibrils in the cell wall (Paredez et al., 2006). In accordance with this scenario is the observation that microtubules are highly aligned perpendicular to the axis of cell expansion in highly anisotropic root cells. By contrast, cortical microtubules display rotary movements in very young hypocotyl cells and in the meristematic dome, consistent with the observation that growth is rather isotropic in those tissues (Chan et al., 2007; Hamant et al., 2008). It must be noted, however, that in certain cases, this colinearity is not observed, suggesting that the deposition of oriented cellulose microfibrils involves other levels of control (Himmelspach et al., 2003). Recently, an additional function of the cortical microtubules in the delivery of CESA complexes to the plasma membrane has been identified, showing a more complex role of the cortical microtubules in organizing cellulose synthesis at the cell cortex (Crowell et al., 2009; Gutierrez et al., 2009)
From a morphogenetic point of view, this provides a mechanism linking microtubules to the control of anisotropy. Indeed, in the footsteps of Paul Green, we observed a shift to isotropic growth in meristems treated with the microtubule depolymerizing agent oryzalin (Grandjean et al., 2004; Hamant et al., 2008; Fig. 1a,b) Interestingly, the geometry of meristematic cells after oryzalin treatment resembled that of soap bubbles in a two-dimensional (2D) froth: angles between adjacent walls in vertices tend towards 120° as in foams. More strikingly, the curvature of the walls was concave in cells having more than six walls and convex in cells having less than six walls as observed in 2D froths (Corson et al., 2009). Beyond the attractive homology between these two systems, this further demonstrates the crucial role of microtubules in anisotropy, as 2D froths are isotropic by essence.
Consecutively, the importance of the microtubules in controlling morphogenesis raises the question of the identity of the factors that control the organization of the microtubules. The cell wall itself influences microtubule organization: in a screen for mutants that are hypersensitive to oryzalin, Paredez an coworkers (Paredez et al., 2008) identified PRC1 and KOR, two proteins involved in cellulose synthesis, as regulators of microtubule organization. In a more extreme situation, when the cell wall is entirely removed during protoplasting, microtubules are unable to remain aligned, again confirming that the cell wall is directly or indirectly required for microtubule organization. The exact nature of the cell wall–microtubule dialog is still far from being completely elucidated.
A major aspect of cortical microtubules functions relates to their dynamics. Changes in the main microtubule orientation within a cell, seem to occur through the synthesis of microtubules in a new direction, which will then affect the orientation of the whole array by depolymerization and/or by reorientation of pre-existing microtubules (Lloyd & Chan, 2004). Katanin, a protein that cleaves the microtubules, appears to play an important role in this reorganization process. Mutations in the Arabidopsis katanin strongly affects anisotropy and result in compact, dwarf phenotypes, associated with reduction of cell length and increased cell width. This defect in the cytoskeleton also leads to problems in the cell wall which is weakened, owing to a decrease in the production of cellulose and hemicelluloses (Bichet et al., 2001; Burk et al., 2001). In katanin mutants, microtubules do not manage to organize in parallel arrays, and seem to remain connected to their nucleating site longer than in the wild-type (Burk et al., 2001). Using field emission scanning electron microscopy, a slight but significant alteration of microfibrils alignment could be detected in the innermost layer of the cell wall in the katanin mutant allele fra2, consistent with the role of microtubule organization in controlling cellulose deposition (Burk & Ye, 2002). Conversely, inducible overexpression of katanin in Arabidopsis favored the organization of microtubules into highly organized arrays. Longer induction led to complete microtubule depolymerization (Stoppin-Mellet et al., 2006). Based on these data, it was proposed that katanin activity helps to free the microtubules from their nucleating site by severing, thus affecting the frequency of their encounters at the cortex and allowing them to organize into arrays. Several other microtubule regulators, such as TONNEAU, CLASP1, MAP65 and MOR1, have been shown to control the organization of cortical microtubules by directly or indirectly modulating their dynamics (for a review, see Ehrhardt, 2008; Wasteneys & Ambrose, 2009).