Cell division and turgor-driven stem elongation in juvenile plants: A synthesis

Plant Science 207 (2013) 45–56

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Review

Cell division and turgor-driven stem elongation in juvenile plants: A synthesis Ulrich Kutschera a,∗ , Karl J. Niklas b a b

Institute of Biology, University of Kassel, Heinrich-Plett-Str. 40, D-34132 Kassel, Germany Department of Plant Biology, Cornell University, Ithaca, NY 14853, USA

a r t i c l e

i n f o

Article history: Received 26 October 2012 Received in revised form 16 January 2013 Accepted 8 February 2013 Available online 20 February 2013 Keywords: Auxin action Cell division Cell elongation Sucrose metabolism Turgor regulation Wall extensibility

a b s t r a c t The growth of hypocotyls and epicotyls has been attributed to the turgor-driven enlargement of cells, a process that is under the control of phytohormones such as auxin. However, the experiments presented here and elsewhere using developing sunflower (Helianthus annuus L.) seedlings raised either in darkness (skotomorphogenesis) or in white light (WL) (photomorphogenesis) indicate that auxin-mediated segment elongation ceases after 1 day, whereas hypocotyl growth continues in the intact system. Based on these results and data from the literature, we propose that hypocotyl growth consists of three interrelated processes: (1) cell division in the apical meristematic regions; (2) turgor-driven cell elongation along the stem; and (3) cell maturation in the basal region of the organ. We document that the closed apical hook (or the corresponding region after opening in WL) is the location where cell division occurs, and suggest that the epidermis and the outer cortex plays an important role in a “pacemaker system” for cell division. Results from the literature support the hypothesis that pectin metabolism in the expansionlimiting epidermal cell wall(s) is involved in wall-loosening and -stiffening. During hypocotyl growth in darkness and WL, turgor pressure is largely maintained, i.e., in H. annuus no hydrostatic pressureregulated growth occurs. These data do not support the “loss of stability theory” of cell expansion. Finally, we document that turgor maintenance during organ elongation is caused by sucrose catabolism via vacuolar acid invertases, resulting in the generation of hexoses (osmoregulation). Based on these data, we present an integrative model of axial elongation in developing seedlings of dicotyledonous plants and discuss open questions. © 2013 Elsevier Ireland Ltd. All rights reserved.

Contents Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Segment elongation versus organ growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Wall extension, auxin action, and pectin metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Meristematic activity in the outer and inner tissues . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Regulation of the plant cell cycle . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cell division and organ growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Turgor pressure and the loss of stability theory . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Cell enlargement and wall plasticity . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Sucrose metabolism and turgor regulation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Conclusions and outlook . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

45 46 47 49 50 50 52 53 53 54 55 55

Introduction In most animals, body size and organ number are predetermined developmentally and, baring extensive trauma, largely

∗ Corresponding author. Tel.: +49 561 804 4467. E-mail address: [email protected] (U. Kutschera). 0168-9452/$ – see front matter © 2013 Elsevier Ireland Ltd. All rights reserved. http://dx.doi.org/10.1016/j.plantsci.2013.02.004

insensitive to external environmental conditions. This mode of size increase, which is called determinate growth, contrasts sharply with the majority of land plants (embryophytes), which continue to add new cells, tissues, and organs throughout their lifetime as a result of continued meristematic activity [1,2]. This mode of indeterminate growth has been extensively studied at the morphological, physiological, and molecular level for a variety of

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Fig. 1. Developing sunflower plants as model organisms for the analysis of organ growth. Photograph of a 3-day-old etiolated seedling (Helianthus annuus L.) that was raised in moist vermiculite at 100% rel. humidity (25 ◦ C, darkness) (A). Light micrographs of the epidermal cells in the hook (B) and basal region (C) of the hypocotyl. Transverse section of the sub-apical region of the hypocotyl (see arrow in A) that shows the vacuolated cells of the outer three tissue layers, inclusive of the thickened peripheral walls (D). Co = cortex, Ep = epidermis, Ho = apical hook, Hy = hypocotyl.

reasons, not the least of which is that it affords an opportunity to experimentally manipulate a reiterative developmental system. Curiously, however, despite extensive investigations, controversy continues regarding the phenomenology of plant organ growth, particularly in terms of the relative importance of cell division and expansion as mechanisms of irreversible volume increase in developing roots, stems and leaves. The German botanist Julius Sachs (1832–1896) distinguished, at the level of the individual organ, among three separate stages of plant growth: (1) cell division in the meristem, (2) subsequent cell enlargement (elongation or expansion), and (3) cell maturation attended by the stiffening of the cell wall [3]. Despite this insight, some workers have reduced the entire process of organ growth to the level cell enlargement [e.g., Ref. [4]], a mode of thinking that originated with the experimental work of the Dutch/American plant physiologist Anton N.J. Heyn (1906–1992), who studied the growth of excised coleoptile- or stem-segments incubated in an aerated solution of auxin (indole-3-acetic acid, IAA). Based on his studies, Heyn formulated the “plasticity theory”, which states that turgor-dependent cell- and organ-elongation is determined by the plastic extensibility of cell walls [5–7]. In turn, this perspective, in which organ growth is controlled by cell wall extension, lead to the development of the “Lockhart-concept” of turgor-driven cell growth in size [8–10], which has been challenged recently by Wei and Lintilhac [11–13] who conclude that turgor pressure must reach a critical value at which point the cell wall looses stability and enlarges – a concept called the “loss of stability theory”. Alternatively, some workers have argued that plant organs can grow by cell division without subsequent elongation [14], a perspective challenged based on physical principles by Paul Green (1931–1998) and others who argue that “plant growth by cell division” is impossible [15–17]. More recent work using the model plant Arabidopsis thaliana (L.) Heynh. has yielded conflicting results concerning this “cell division versus enlargement” controversy (see Ref. [18], and references cited therein). In two recent papers, we analysed the changes in metabolic activity (cell respiration) during the development of sunflower (Helianthus annuus L.) seedlings [19,20] and showed that the apical hypocotyl hook has a much higher rate of oxygen uptake per gram fresh mass than the region where cell elongation takes place. This finding motivated us to explore in greater detail the relationships between hypocotyl hook-associated cell division and turgor-driven elongation in a well known model plant system, particularly with

reference to conflicting theories concerning cell wall loosening and cell enlargement (Fig. 1). In this review article, which is based in part on new data, we show that cell division and turgor-driven organ expansion are inter-related processes, that must be integrated, to more comprehensively model the process of plant development. Segment elongation versus organ growth As noted, Heyn [5,6] inaugurated a series of extremely influential experimental studies on the biophysical basis of IAA-mediated organ (surface) elongation [4,21,22]. Using excised, auxin-depleted sections cut from etiolated coleoptiles or stems (Figs. 1A and 2A), he showed that the growth response in the presence of IAA is attributable to turgor-dependent cell anisotropic expansion. As a consequence of his studies, Heyn [6], who published his last paper on this topic fifty years after his first report appeared in print [7], proposed the “wall plasticity concept”, which posits that the rate of organ elongation is determined by the plastic extensibility of the cell wall, but not by wall elasticity, or changes in turgor pressure, a conceptualization that has been questioned recently (see Turgor Pressure and the Loss of Stability Theory section). This view of plant growth raised many questions, as for example “How is the direction of growth determined?”, and “Why do some cells expand anisotropically (elongate) under the equi-directional force of turgor pressure?”. Although much remains to be learned in this context, a number of studies have shown that, in developing maize seedlings, microtubule (MT)-depolymerizing drugs, such as colchicine, exert a dramatic effect on the shape of the turgid organ [4]. Instead of anisotropic enlargement, the cells expand isotropically in all directions, and, as a result, organs become nearly spherical. Cytological analysis reveals that in colchicine-treated seedlings, the cellulose–microfibrils on the inner wall layer are randomly oriented, and not perpendicularly as in the axial organs of the controls. These and other data indicate that microtubule orientation influences the direction of microfibrile deposition, and hence cell- and organ shape [4]. A detailed discussion of the microtubule– microfibrile-relationship is beyond the scope of this article. It has long been known that IAA-induced segment elongation in vitro, which is promoted by sucrose [22], ceases after 24 h, even in the presence of the disaccharide. As illustrated in Fig. 2A, 10 mm long segments cut from the growing region of the stem and incubated for 24 h in sucrose elongate by 30% and 70% in the absence or presence of IAA, respectively. However, in the intact system, the

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Fig. 2. Auxin-mediated elongation of excised hypocotyl segments versus organ growth in 4-day-old etiolated sunflower seedlings. Sections, 1 cm in length, were excised and incubated for 24 h on sucrose solution (50 mM) ± auxin (indole-3-acetic acid, IAA; hormone concentration: 10 ␮mol l−1 , darkness, 25 ◦ C) (A). Growth of the corresponding sub-apical region over 48 h in darkness is indicated in the seedling depicted on the right side (B). Note that, in the intact plant (in situ), organ growth continues, whereas segment elongation (in vitro) ceases after 24 h.

marked 10 mm region of the sunflower stem continues to elongate over the subsequent 48 h by 250% (Fig. 2A and B). This long-term growth in situ is due to cell division in the apical hook, which has the highest metabolic activity in the axial organ [19,20]. The data summarized in Fig. 3A and B show that stem elongation and increases in cell number are positively correlated in H. annuus. The hypocotyl of 2-day-old and 6-day-old etiolated seedlings (length ca. 8 mm and ca. 85 mm, respectively) consists of approximately 0.6 and 3.0 million cells, respectively. Upon irradiation with white light (WL), hypocotyl elongation and cell division are reduced in parallel. These data show that cell division and enlargement via turgor-driven wall expansion are equally important. In the next section, we focus on the mechanism(s) of cell expansion, and thereafter discuss cytokinesis in the context of plant meristems. Wall extension, auxin action, and pectin metabolism The chemical composition of the plant cell wall changes as a function of cell differentiation. It also differs among cell-types and even among otherwise comparable cells isolated from closely related species. Nevertheless, the cell walls of the chlorobionta (i.e., the green and charophycean algae, and the land plants) [1,2] is a fibre-reinforced composite material that consists of cellulose cross-linked microfibrils held in tension within a matrix composed of hemicellulose, pectins, structural proteins, and, in some cells, lignin. The growth of a cell with this kind of wall involves a balance between the extensibility of the wall and the internal mechanical force exerted on it by turgor pressure. By analogy, the growth of plant stems involves a balance between the extensibility of epidermal cell walls and the mechanical hydrostatic forces exerted on them by the thin-walled, extensible cells these walls surround (i.e., the “epidermis-in-control-theory” of organ growth, see Kutschera

Fig. 3. Time-course changes in hypocotyl length (organ growth in situ) during development of sunflower seedlings that were either grown in darkness (D) or irradiated with continuous white light (WL, photon fluence 100 ␮mol m−2 s−1 ) (A). The changes in the number of cells per hypocotyl are shown in the lower graph (B). Arrows denote the onset of WL irradiation. Source: Adapted from Ref. [46].

and Niklas [23]). Although an increase in turgor pressure can cause a cell or an organ to expand temporarily, permanent (plastic) cell or organ growth requires a change in the mechanical properties of the cell wall or the epidermis (specifically, a reduction in the cell wall yield stress), and an influx of water that causes an increase in total volume. Growth slows and may ultimately stop with a cessation of cell division and a subsequent return to the previous mechanical properties of the wall or the epidermis. The physiological mechanisms responsible for cellular expansion, and hence stem or coleoptile elongation, resulting from a reduction in the yield stress of the growth-controlling cell wall(s), are comparatively well known [24,26,27]. They include auxin (IAA), a plasma membrane-bound auxin binding protein (ABP–IAA), and (V-type) ATPases that can collectively acidify the endomembrane system or the cell wall, thereby loosening it via the promotion of Golgi-mediated secretion/incorporation of cell wall materials [22,28]. More recent studies of the stems of dicotyledonous plants also highlight the role of methylesterases and Ca2+ in affecting the mechanical properties of pectins and altering the degree of cell wall hydration. These two “cell expansion subsystems” can be rendered as a logic circuit, which shows how the components interact (Fig. 4). The first of these two subsystems involves rapid and delayed responses to the formation of the ABP–IAA conjugate, which serves

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Fig. 4. Schematic for a logic circuit for IAA-mediated cell wall loosening mediated by two subsystems, one involving an auxin (IAA) binding protein (AMP) signal transduction (subsystem assembly I) and another involving pectin methylesterase (PME) effects on galacturonan rich pectins (subsystem assembly II). Each subsystem is diagrammed as a signal-actuated (S-A) system with an actuator/suppressors, an assembly process, and an output signal (demarked by thin rectangles). Each subsystem also requires one or more feedback elements and comparators, which are largely currently unknown for the subsystems diagrammed in this logic circuit. See Ref. [27] for a detailed treatment of logic circuit terminology. PM = plasma membrane; ER = endoplasmic reticulum.

as the subsystem’s actuator switch. The ABP binding of IAA results in a rapid hyper-polarization of the cell membrane, actives electrogenic ATPases, that acidifies the apoplast, which may contribute to the reduction in the cell wall yield stress, presumably as a result of increasing the distance between neighbouring cellulose microfibrils. Alternatively, the activation of apoplastic polysaccharidases that loosen the growth-limiting cell wall(s) has been proposed for IAA-(and fusicoccin)-mediated coleoptile elongation [24]. More recently, a rapid IAA-mediated activation of Golgi-secretion in the epidermal cell layer of grass coleoptiles has been implicated as a mechanism of hormone-mediated wall loosening [22,28]. It is important to note that the use of the word “actuator” in the context of this or any logic circuit diagram referencing a physiological system is used to denote the subsystem responsible for a “signal transduction” event that stimulates subsequent referenced events in the logic circuit. This usage contrasts with the logic circuit notation “leads to”, which references downstream events about which one or more details are uncertain. For example, auxin-mediated processes at the level of gene expression that occur hours after application of the hormone are not yet elucidated in detail [29]. A 3- to 8-fold enhancement in translatable mRNA is associated with accumulations in ribosomes and rRNA-molecules, respectively. Within 5 h after IAA treatment, about 40 mRNAs are up- or down-regulated. The functions of these gene products during long-term growth are also largely unknown, but it is likely that some of them may be involved in the process of Golgi-mediated cell wall biosynthesis [29] (see Fig. 5B). In contrast to the walls of monocots, such as grass coleoptiles, the cell walls of dicots, such as sunflower (Figs. 1 and 2),

are rich in pectic substances (up to 30% per g wall mass). Recent data obtained from experiments with the growing hypocotyls of dicots [30–32] and the logic circuit for cell wall assembly indicate that the mechanical properties of pectins may provide a specific “dicot-mechanism” capable of altering the mechanical properties of the growth-controlling cell wall(s) (Fig. 4). Pectins form a functionally and structurally diverse class of galacturonic acid-rich polysaccharides that can undergo significant modifications in their physicochemical properties. Recent attention has focused on homogalacturonan demethylesterification catalysed by the ubiquitous enzyme pectin methylesterase (PME) as an important component in the control of cell wall hydration, expansion, and growth [33,34]. Reconstructions of systems composed of cellulose and pectin, using cellulose-producing bacteria, document that pectin enhances the extensibility of the composite, even after the removal of these “soft” wall polymers [34]. Several studies have shown that cross-linking of pectin sub-domains by boron is necessary for wall biogenesis and organ growth [35,36]. Perhaps more important is the fact that dynamic modifications of homogalacturonan (HG), a component of the pectins within the matrix of the wall, may play important roles during cell growth (Fig. 5A and B). Specifically, HG is polymerized in the cytoplasmic Golgi apparatus by glycosyl transferases. After substitution with methyl groups at the C6 position, the material is deposited into the walls in a methyl-esterified state [36] (Figs. 4 and 5). Once outside the cell, pectin methylesterase (PME) can remove methyl units, which releases free carboxylic acid groups, methanol (which can be consumed by some epiphytic bacteria [37]), and protons. Depending on pH, ion availability, and other physiological factors, PME1 activity

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Fig. 5. Scheme depicting the biosynthesis of cellulose microfibrils by apposition and the Golgi-mediated secretion of wall matrix components in the growth-controlling outer epidermal wall. A transmission electron micrograph (A) shows the ultrastructure of the cytoplasm and the outer epidermal wall, covered by the cuticle, of the peripheral hypocotyl cell in a 4-day-old etiolated sunflower seedling. Cellulose deposition by apposition of microfibrils and delivery of matrix components are depicted (B). G = glucose, Cu = cuticle, Cw = cell wall (outer epidermis), Cy = cytoplasm, Ma = matrix, Mc = mitochondrion, Mi = microfibril, MT = microtubule, Ve = vesicle; Ca2+ = calcium ions, B = boron (cell wall-associated).

also results in a variety of different methyl esterification configurations. For example, in hypocotyls and pollen tubes, a low percentage of pectin methyl esterification is correlated with a decline in cell wall extensibility and the cessation of cell enlargement [36]. In addition, Ca2+ -mediated cross-linking of dimethyl-esterified pectin may contribute to cell wall stiffening by reducing the ability of cellulose microfibrils to shear in the cell wall matrix [32,33]. Alternatively, pectin dimethyl esterification can also provide a mechanism for wall loosening by facilitating the degradation of homogalacturonan via polygalacturonase. What can be said with some certainty is that dimethyl esterification expedites cell wall hydration and decreases cell wall pH, both of which can significantly alter the mechanical properties of cell walls. The physiological versatility of PME-mediated dimethyl esterification appears to be ancient among the chlorobionta, since HG methyl esterification epitopes are reported to exist in unicellular charophycean algae (e.g., Penium margaritaceum) [38,39]. However, since the walls of grass coleoptiles contain low amounts of pectic substances, the wall-loosening model discussed here applies primarily to the growing organs of dicot plants, such as sunflower hypocotyls (Figs. 1 and 2), mung beans, and the model organism A. thaliana [32,33,40]. Finally, the logic circuit diagram for cell wall growth shows that sustained turgor maintenance via osmoregulation and cell wall loosening are required for continued cell expansion (Fig. 4). In developing sunflower hypocotyls, these processes have been analysed in detail and are discussed in Cell Enlargement and Wall Plasticity section. However, the feedback loop and comparator elements that are involved in the cessation of the cell expansion machinery are poorly known. Auxin degradation, the re-orientation of cellulose microfibrils, the biosynthesis of additional cell wall binding polymers, the apposition of secondary cellulose layers, and the reduced secretion of wall-loosening agents into the maturing walls are among the candidates for these not yet elucidated circuit components. An additional process that may be involved is extensin cross-linking. Extensin, a hydroxypoline-rich cell wall protein, can establish networks via intra-molecular tyrosine cross-links. Extensin molecules, which are amphiphilic, bind to pectins through electrostatic interactions. This causes the dehydration of the “soft”

matrix fraction of the walls. Together with lignification, this “extensification” of the wall matrix may be part of the processes leading to the decrease in cell wall extensibility and the cessation of organ growth [41]. Meristematic activity in the outer and inner tissues According to the “epidermal-growth-control theory”, the thickened, helicoidal outer walls of an organ regulate the rate of expansion of the entire system [23,42–44]. However, the question as to the tissue-specific localization of cell division activity has not yet been discussed in this context. In order to localize the meristematic activity of the cells along the hypocotyl of developing sunflower seedlings, an immunocytochemical method was employed. To quantify how many cells are in DNA-synthesis (S)-phase (as a percent per tissue), segments were cut from different regions of the hypocotyl and analysed [45,46]. The results of these studies are summarized in Fig. 6. The data show that a high percentage of nuclei in S-phase, which indicates an active state of cell division within the tissue, is largely restricted to the closed apical hook, or the region below the cotyledons. In the sub-apical (elongating) region, where the cells elongate by turgor-driven anisotropic expansion of the walls (Figs. 1 and 2), cell division activity is close to zero, whereas in the basal region above the onset of the root, no nuclei in S-phase are found. These results are in accord with our recent finding that the hook displays the highest rate of oxygen uptake per g fresh mass per minute (metabolic activity) [19,20]. The data summarized in Fig. 6 further show that meristematic activity (i.e., the percentage of nuclei in S-phase) manifests a developmental pattern that is consistent with the overall growth of the hypocotyl (see Fig. 3A). In young, actively expanding organs, meristematic activity is high. However, during the cessation of growth at day 6 or under white light (WL), cell division slows down. The only exception to this generalization occurs during the opening of the hook (which occurs in 4-day-old seedlings, irradiated for 24 h with continuous WL, Fig. 6B), which displays a higher meristematic activity compared to all other stages of development (skotomorphogenesis in darkness and photomorphogenesis in WL, respectively) [19,20].

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Fig. 6. Cell division activity in developing sunflower hypocotyls. Changes in meristematic activity, determined as percentage of nuclei in S-phase, in seedlings that were either grown for 2, 4 and 6 days in darkness (2D, 4D, 6D) or for 3 days in darkness and for 1 or 3 days in continuous white light (1 WL, 3 WL; photon influence see Fig. 3). The dark bars denote the tissues of the stem where cell division activity occurs (hook, apical part). Ep = epidermis, Pi = pith.

are the manner in which their functionalities are regulated by plant hormones such as IAA, cytokinins (CytoKs), and abscisic acid (ABA). The available information indicates that these complexes participate in at least three critical cell cycle control points: (1) the negative regulation of retinoblastoma-related (RBR) protein (by the phosphorylation of S and T residues) that in turn binds to and either actives or suppresses the activity of a variety of E2F/DP transcription factors participating in the G1-to-S transition; (2) the regulation of pre-RC functionality during the initiation of DNA replication, and (3) the control of the G2-to-M transition, e.g., CDKB1 (an E2F target gene product) is maximally expressed in the G2-to-M transition and is necessary for the balance between cell division and, if present, the endoreplication cycle [49]. As noted, hormones are also required for the control of cellular proliferation and differentiation. In plants, the roles of IAA and CytoKs are particularly well understood, while the roles of ABA and other hormones, such as brassinosteroids and ethylene, are increasingly well documented [50]. Recent research continues to reveal the central role played by IAA. For example, IAA signal transduction activates the AXR1 gene whose product is structurally required for the SCFSKP2 complex as well as involved in the proteolysis of E2F transcription factors, such as E2Fc. AXR1 is also implicated in the degradation of targeted proteins. CytoKs are involved in the induction of cell division and help to regulate the G1-to-S and the G2-to-M transitions. In conjunction, IAA and CytoKs are known to activate the PROPORZ1 (PRZ1) gene responsible for the production of the transcriptional adapter protein PRZ1, which appears to regulate the expression of cell division genes and mediates the hormonal signal for cellular proliferation (as evidenced by the ability of lovastatin, a CytoK biosynthesis inhibitor, to block cells in mitosis, which can be alleviated by the exogenous addition of CytoKs) [51,52].

Source: Adapted from Ref. [45].

Cell division and organ growth

Finally, the results shown in Fig. 6 indicate that meristematic activity is much higher in the epidermis (plus outer cortex, see Fig. 1D) than in the centre of the stem (i.e., the pith). Hence, the growth-controlling outer cell layer(s) not only determine(s) the rate of organ elongation, but may also serve as part of a “pacemaker system” for cell division. It should be noted that, in sunflower hypocotyls, the nuclei do not show endopolyploidy, i.e., they remain diploid throughout the organ’s entire development (day 0–6 after sowing) [46].

A critical feature in plant development is the orientation and location of the future cell wall during cell division, which in turn results in the formation of different types of tissue construction [1,52–54]. In our previous reports on organ development in sunflower hypocotyls with respect to metabolic scaling [19,20], we showed that cell division activity is high in the hook of juvenile seedlings (Fig. 1A), and undetectable at the base of the stem (see Fig. 1B and C). Here we discuss general aspects of this process as illustrated in Fig. 8A, which shows meristematic cells in the hook of 2-day-old sunflower seedlings (see Fig. 6A). Among the charophycean algae and the embryophytes, the location of the future cell wall is prefigured by the appearance of the preprophase band (PPB) and later by the phragmoplast (Fig. 8B). The mechanisms underlying the orientation and location of these cytological features have been investigated for many decades but are not well understood. In the 19th century, it was noted that cell plate formation occurs at right angles to existing walls so that two derivative cells of equal size are created as a result of every cell division (Sachs’s rule [3,59]). Early in the 20th century, it was reported that the application of pressure to a dividing cell forced the mitotic figure into the position in which the longitudinal axis was oriented at right angles to the applied pressure such that the future cell wall was oriented parallel to this direction [55,56]. Likewise, Steward et al. [57] noted that cells in free suspension have highly irregular and unpredictable planes of division, perhaps because they are not restricted peripherally as they would when cells normally grow within the plant body. Among certain colonial cyanobacteria, flagellates, and pollen sporocyctes wherein cell divisions are simultaneous, the planes of successive division tend to be at right angles to one another such

Regulation of the plant cell cycle The results presented in the previous sections show that meristematic activity is much higher in juvenile organs. However, our immuno-cytochemical data (Fig. 6) do not reveal how the cell cycle is regulated in meristematic tissues. Phylogenetic and molecular analyses indicate that the basic machinery and control of the cell cycle is highly conserved among fungi, animals, and plants, although phylogenetically specific cellcycle genes and hormones are known for each of these major eukaryotic clades. For example, as in fungi and animals, cyclindependent kinases (CDKs) are major components in plant cell cycle transitions [47]. Among the complex CDK Arabidopsis family, which consists of 12 members, CDKA is homologous to yeast Cdc2 and necessary for the G1-to-S and the G2-to-M transitions (Fig. 7). CDKB proteins, such as CDKB1, which are specific to plants, are influenced by auxin (IAA) and are expressed from S to M and in the G2-to-M transition during the cell cycle. They also play a role in the development of stomata [48]. Nevertheless, the roles of individual plant CDK-cyclin complexes are not known completely nor

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Fig. 7. Schematic for a logic circuit for the cell cycle diagrammed as a signal-actuated (S-A) system with an actuator/suppressors, a subsystem assembly process, and an output signal (each demarked by thin rectangles). The subsystem requires one or more feedback elements and comparators, which are largely currently unknown or poorly understood. See text for additional details. ABA = abscisic acid, CDKs = cyclin-dependent kinases, CDKB1 = an E2F target gene product, CytoKs = cytokinins, RBR = retinoblastoma-related protein.

that regular patterns of two, four, eight, etc. form, all in one plane [58]. A complementary geometric view of this process is known as Errera’s rule and has recently been explored as a modelling approach [59]. That biomechanically induced mechanical stresses may be involved in cell wall orientation is consistent with many observations [60,61]. The simplest plant cells are those that constitute parenchyma, which have thin primary walls that are

hydrostatically inflated. The turgor pressure exerted against the walls of these cells is more or less uniform, i.e., the stresses resulting from turgor pressure within cell walls are uniform, both within each cell, and among neighbouring cells. However, at the vertices created by adjoining cells, opposing tensile stresses are resolved into additional stresses acting in the radial direction on the angle of each vertex according to its angle size. In theory, the tensile stresses in walls at 180◦ should be equal and opposite, and

Fig. 8. Cell division in the meristematic region of a developing stem. Paradermal view of the Helianthus hypocotyl, showing dividing cells in the apical hook of a 2-day old etiolated seedling (A) and schematics of the establishment of the future cell wall during embryophyte cell separation (B). The preprophase band (PP-band) is established around the nucleus before mitosis and prefigures the location of the phragmoplast, which facilitates the deposition the future cell wall (upper series of diagrams). In turn, it is hypothesized that the location of the nucleus (and thus the PP-band) is determined by microtubules (MTs) (see Fig. 5B) achieving equilibrium lengths minimizing the distance between their attachments to the cell wall and the nucleus (lower series of diagrams).

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thus this angle experiences no additional radial stress from the resolution of the opposing tensile stresses in the two intersecting walls. However, these tensile stresses are resolved into progressively larger radial stresses as the angle of a vertex decreases, reaching their maxima as the angle approaches 0◦ . Because these additional radial stresses are correlated directly to the size of the angle, stresses reach mechanical equilibrium at equiangular vertices [62]. Consequently, the observation that the vertices in the region of isodiametrical expansion can act as cellular pivots for wall rotation between successive divisions (so as to coincide with cellular mechanical equilibria) provides some evidence for the biomechanical regulation of cell shape [54,62]. This biomechanical scenario is vastly different from that operating in an elongating cell (e.g., xylem fibre), wherein existing walls rotate around their vertices to align either perpendicular or parallel to the longitudinal axis and future cell walls are generally oriented perpendicular to the growth axis. Here, the principal stress trajectories likely resolve the global stress patterns into orthogonal components and are thus likely to be oriented parallel and perpendicular to the growth axis. In this condition, cell walls may be oriented so as to minimize shear stresses [62].More recent research on the orientation of the future cell wall has focused on highly detailed analysis of the charophycean alga Coleochaete orbicularis and the dicot Zinnia elegans. Digital analyses and computer simulations of these patterns indicate that multiple competing division planes exist [59]. However, if significant differences in the surface areas of competing division planes are established, the smaller area configuration will be, on average, achieved. However, if small differences exist among competing planes of division, the probability of achieving the division plane with the minimal area is inversely proportional to the difference in length of the division plane (Fig. 8B). Thus, in cells differing significantly in length, the shortest transverse division planes are achieved, whereas competing planes of cell division with similar surface areas are achieved with near equi-probability in polygonal cells [62]. The mechanism responsible for the selection of the future cell wall plane is conjectural. However, it is speculated that a microtubule (MT) length-dependent force-sensing system permits the MT cytoskeleton to reposition the nucleus into an equilibrium position [59,63]. According to this scenario, the nucleus during interphase is positioned off centre and MTs radiating from it, outward to the cortex, re-centre the nucleus based on differences in the tensile forces generated among MTs differing in length. Shorter as opposed to longer MTs would be favoured collectively to achieve the equilibrium configuration, which would automatically coincide with the minimal area plane that concurrently triggers the formation of the PPB (Fig. 8B). It should be noted that the proposed mode of MT-dependent cell division [59] has not yet been analysed in the meristematic region of sunflower hypocotyls (Fig. 8A), i.e., more work is required to further corroborate this cell division concept. At any rate, G0-phase cells that have divided elongate by vacuolation, water uptake, and wall expansion. This process has been analysed in detail in developing sunflower hypocotyls and is discussed in the next section.

Turgor pressure and the loss of stability theory Heyn [5,6] was the first to hypothesize that IAA reduces the ability of the cell wall to resist internal hydrostatic pressure (cell turgor), which places the cell wall under tension [24,25]. This model of surface growth was elaborated and extended by Lockhart [9,10] for a cylindrical cell by means of a simple equation: dV = m(P − Y ), dt

where the growth rate (i.e., volume increase, dV/dt, unit: m3 s−1 ) is a function of the yielding coefficient of the wall, also known as extensibility (m, unit: m3 s−1 MPa−1 ), turgor pressure (P, unit: MPa), and the yield threshold required for irreversible cell expansion (Y, unit: MPa) [64,65]. This “single cell-equation” was later extended by Lockhart [10] to treat the mechanics of axial organ growth by drawing a physical analogy between a Chara or Nitella (Characeae) cell wall and the epidermis of a stem – an analogy that serves as the basis of the “epidermal-growth-control-theory” of organ elongation [23,42–44]. This classical Heyn–Lockhart-model has been challenged recently by Wei and Lintilhac [11–13], who argue that the “viscoelastic/creep-concept” of turgor-driven wall expansion must be replaced by their “loss-of-stability” theory, which is largely predicated on experiments with growing Chara cells. This theory posits that turgor pressure must rise to a critical point “determined by material properties and cell geometry, followed by a loss of stability that manifests itself as wall extension and growth” [11]. Wei and Lintilhac go on to say that “As water enters a cell, turgor pressure increases; once turgor . . . reaches its critical value, the wall loses stability, with wall stress relaxation and cell enlargement resulting” [11]. This alternative model of cell enlargement has been criticized by Schopfer [66] on the basis that the Wei–Lintilhac-model fails “to appreciate that plant cells behave as hydraulic systems modelled by osmometers, the mechanical properties of which are governed by osmotic water relations rather than by physical mechanics of closed pressure vessels.” This criticism is valid since the material properties of a closed pressurized vessel do not change as a function of internal pressure, nor are they enzymatically regulated as they are in a cell wall. However, the “loss-of-stability theory” proposed by Wei and Lintilhac is conceptually not at great odds with the classical model in that both posit that cell expansion occurs, once a critical internal pressure is reached (i.e., a pressure that exceeds the yield strength of the cell wall). The primary difference between the perspectives offered by Schopfer and the Wei–Lintilhac-model is that the latter posits cell wall material “failure”, whereas the former proposes a hormone (or light)-mediated “controlled relaxation and deformation” of the extensible polymeric structure of the cell wall [4,66]. Indeed, we think that the salient difference between the classical Heyn–Lockhart-model and the Wei–Lintilhac-concept lies in how the mechanical response of the cell wall to the hydrostatic pressure exerted on it by the protoplast is conceptualized, which is not a trivial issue. The classical model requires that the cell wall responds comparatively slowly and incrementally as cellulose microfibrils are allowed to shear in a matrix that has become more fluid-like (see Fig. 5B). On the other hand, the Wei–Lintilhac-concept essentially posits that the cell wall suddenly mechanically “fails” at a critical turgor pressure. Therefore, we think that these two contrasting views do not differ in terms of the fundamental mechanism responsible for cell wall expansion. However, they differ critically regarding how the cell wall mechanically responds to turgor pressure, which arguably depends on cell wall chemistry (which can differ even among related species) [39,53]. Only detailed studies will show whether the Wei–Lintilhac-model is appropriate for Chara internodal cells. However, even if true, this finding would not necessarily invalidate the classical Heyn–Lockhart-theory, since this concept may apply to other model systems. Likewise, it is important to note that the mechanical behaviour of an isolated Chara internodal cell (or any other single plant cell) cannot be extrapolated to predict the behaviour of an organ composed of numerous adhering cells by virtue of middle lamellae, since adjoining cell walls can mechanically restrain the deformations of neighbouring cells. While a physical analogy can be drawn between the elongation of a Chara internodal cell and the growth of a hypocotyl, the two systems

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differ in many important physical as well as biological respects [1,2]. Finally, it should be noted that the “Wei–Lintilhac vs. Heyn–Lockhart”-controversy concerning the isotropic surface growth of axial plant cells (or multicellular organs) discussed above has a counterpart in a related area of plant research. The growth of pollen tubes occurs at the tip of the organ and is likewise a turgordependent process. According to the “hydrodynamic model” of pollen tube (tip) growth, the increase in cell volume is driven by localized pressure differences [67]. This view, which corresponds to the “loss of stability (Wei–Lintilhac)”-concept [11–13], is at odds with the finding that turgor pressure does not change during tip elongation [68]. These and other data strongly support the “cell wall-yielding”-theory of pollen tube expansion [68], a concept that corresponds to the “plasticity (Heyn–Lockhart)”-model of stem elongation [5–10]. Finally, it should be noted that the use of the word “plasticity” in this and other contexts is a conceptual redaction of the phrase “a reduction in the cell wall yield stress with accompanying cell wall plastic deformation resulting from turgor pressure” (see Refs. [25,64] for a discussion of this topic).

Cell enlargement and wall plasticity Numerous reports have been published indicating that, during stem elongation, turgor pressure does not change very much [4,5,8–10]. Here we illustrate this biophysical principle based on results obtained on populations of developing sunflower seedlings. The “loss-of-stability” phenomenology predicted by the Wei–Lintilhac-model [11–13] is inconsistent with our data regarding changes in cell length (elongation growth), turgor pressure, and wall plasticity summarized in Fig. 9A–C. All three parameters were measured in the sub-apical (elongating) region of sunflower hypocotyls, as described in Ref. [25]. In juvenile stems (2 days after sowing, see Fig. 6A), a cell turgor pressure of ca. 0.58 MPa was measured. Over the subsequent 2 days, turgor pressure declined and reached a constant value of ca. 0.5 MPa during the linear phase of cell elongation (and organ growth, see Fig. 3A). Cessation of stem elongation in darkness at day 6 after sowing was accompanied by a large drop in turgor pressure, which reached a constant value of ca. 0.3 MPa on days 6 and 7 after sowing, corresponding to the yield threshold for growth in the Lockhart equation (Fig. 9B). Irradiation of the seedlings with continuous white light (WL) causes a large decrease in cell elongation (Fig. 9A), but has no effect on turgor pressure, with the exception of day 6, when turgor is maintained in irradiated seedlings (Fig. 9B). During stem elongation in darkness, wall plasticity displays a significant increase, and declines on day 6. Upon irradiation with WL, wall plasticity is reduced and reaches values that are about half of those observed in the dark controls (Fig. 9C). These data show that hypocotyl elongation is not regulated by an increase in cell turgor (P), but is modulated via changes in wall plasticity. These changes in extensibility are regulated by the behaviour of the epidermis, which is mechanically reinforced by the thickened walls of sub-epidermal cells (see Fig. 1D). The inner tissues of the sunflower hypocotyl (pith, vascular tissues and inner cortex) are composed of thinwalled, turgid cells that remain highly extensible throughout organ development [23,25,69]. Our data are likewise incompatible with a recent report by Kierzkowski et al. [70], who analysed the growth of tomato vegetative shoot apices and conclude that “cells must be able to deform elastically in order to grow”. In developing sunflower hypocotyls, wall elasticity remains largely constant [71], whereas plasticity of the peripheral wall(s) displays changes that correspond to the

Fig. 9. Relationships between the changes in cell elongation (A), turgor pressure (B), and wall plasticity (C) in the sub-apical region of sunflower hypocotyls. The seedlings were grown in darkness or white light (WL) as described in the legend of Fig. 3. The arrows denote onset of WL-irradiation. Y = yield threshold for growth. Source: Adapted from Ref. [25], supplemented by unpublished results.

pertinent growth rates of the cells and the entire organs (Fig. 9A and C). Therefore, the “elasticity-concept” of Kierzkowski et al. [70] does not apply to all growing axial organs, such as sunflower hypocotyls. Sucrose metabolism and turgor regulation In our previous study on auxin-induced growth of excised hypocotyl sections incubated in water (±IAA) (see Fig. 2A), we showed that cell enlargement is negatively correlated with a decline in turgor pressure [23]. Hence, in the absence of absorbable solutes, such as sucrose, a dilution of the vacuolar contents by water uptake results in a corresponding drop in hydrostatic cell pressure. Accordingly, if sections cut from hypocotyls or coleoptiles are incubated in the presence of sucrose (50 mM), turgor is maintained over a longer time period and growth continues for up to 24 h [22]. Our data obtained on intact sunflower stems document that, between days 3 and 5 after sowing, turgor pressure is maintained at a value of about 5.2–5.0 MPa, whereas the length of epidermal cells increases from ca. 70 to 145 ␮m (Fig. 9A and B). Hence, a ca. 100%

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Fig. 10. An integrative model of the cellular basis of stem elongation in juvenile plants, such as sunflower seedlings (A). Cell reproduction (1) is restricted to the apical (meristematic) region of the stem. Isodiametric cells that leave the cell cycle pass into G0-phase and elongate (2). When they have reached their final size, cell maturation occurs and the walls thicken (3). Scheme depicting processes (1) and (2) at the organ level (B), with respect to osmotic adjustment and turgor maintenance via the accumulation of soluble sugars (glucose, fructose) within the enlarging vacuoles. G1 = gap 1; S = DNA-synthesis; G2 = gap 2; M = mitosis; C = cytokinesis; Cw = cell wall; Suc = sucrose; V = vacuole. Source: Adapted from Ref. [25].

increase in cell volume (which should, in the absence of absorbable solutes, result in a 50% drop in cell turgor) is accompanied by the maintenance in hydrostatic pressure. A number of studies have shown that sucrose catabolism in the sub-apical (elongating) part of the hypocotyl (Fig. 9A) is positively correlated with the maintenance of hexose levels and hence turgor pressure [reviewed in Ref. [25]]. Specifically, the catalytic activity of the vacuolar acid invertase (which catalyses the hydrolysis of sucrose into glucose and fructose) was found to be associated with the rate of cell growth. It should be noted that, in addition to potassium ions, glucose and fructose are the major osmotica in the vacuoles of sunflower hypocotyls. Based on these and other data, it was concluded that turgor maintenance in dark-grown sunflower hypocotyls (and during de-etiolation of the stem in WL) is caused by a corresponding rate of hydrolysis of imported sucrose via the specific catalytic activity of a vacuolar acid invertase. Moreover, it was postulated that sucrose is produced via the catabolism of lipid reserves stored in the oil-rich cotyledons of the sunflower seedlings [25]. However, this “sucrose metabolism-turgor maintenance” model of osmoregulation in H. annuus rests on the assumption that the non-reducing disaccharide is in fact transported as the major carbohydrate from the cotyledons into the expanding hypocotyl cells via the phloem (Figs. 1A and 9A). The finding that sucrose is the major sugar carried in the sieve tubes of crop species such as sunflower plants [25] has recently been challenged. In many dicot plants, a large percentage of the carbohydrate in the phloem exudate is found not to be sucrose, but hexoses, such as glucose [72]. However, in a recent study, Lin et al. [73] document that the transport of large quantities of hexoses via the phloem [72] is

an experimental artefact. Using radioactively labelled 14 CO2 , these authors show that the disaccharide sucrose is a ubiquitous transport sugar in seed plants, and that hexoses (glucose, fructose) are essentially absent in the phloem stream in dicot stems and petioles [73]. Therefore, the available evidence thus far indicates that sucrose is the primary (possibly only) mobile sugar in crops like sunflower. Based on these facts, we conclude that osmoregulation and hence turgor-maintenance is achieved via the vacuolar acid invertase-mechanism proposed a decade ago [25], which has been corroborated by other, more recent studies [74]. Finally, we point out that sucrose, a carbohydrate that is exclusively synthesized by oxygenic photosynthetic organisms (embryophytes [1,2]) and transported via the phloem from source to sink tissues [73], is not only a metabolite, but also a signalling molecule [75–77]. For instance, in developing Arabidopsis seedlings, root growth is triggered during photomorphogenesis by imported sucrose generated via photosynthesis within the green cotyledons [78]. We suggest that, in light-grown (de-etiolated) sunflower seedlings, sucrose imported from sink tissues within the cotyledons is the signal responsible for the development of lateral roots. However, more work is required to further corroborate this “sucrose-root-signal” hypothesis of light-induced plant development in the below-ground phytosphere. Conclusions and outlook Our results show that auxin-mediated segment elongation (Figs. 1 and 2) is not an analogue of the growth of an intact plant organ, as has been suggested [4,14,21]. Hypocotyl growth in juvenile H. annuus plants, a model organism for physiological

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and ecological studies [23–25,79], consists of three processes: (1) cell division in the apical meristem (i.e., the cell cycle: M → C → G1 → S → G2); (2) the elongation of G0-phase-cells that have left the meristem; and (3) cell maturation attended by the incorporation of extensin, the apposition of secondary wall layers and, in some cases, cell wall lignification (Fig. 10A). Our quantitative analyses show that cell division in the hypocotyledonary hook and cell elongation in the sub-apical part of the stem are both correlated with the process of organ growth during skoto- and photomorphogenetic plant development, which is primarily regulated via light perception by phytochromes and cryptochromes [80]. Moreover, we document that the plasticity of the growth-controlling peripheral wall(s) determines the rate of stem elongation, whereas turgor pressure is maintained, and finally declines during organ development. These data, which contrast with the “loss of stability concept” for cell growth, provide insights into a classical dicot model plant system that in theory might have broad applicability to other members of the embryophyta, and even perhaps charophycean algae [53,54,81]. However, it is clear from our review of the literature that care must be taken not to generalize about plant growth and development, since differences are reported even between monocots and dicots, not to mention between embryophytes and their sister lineage, the charophycean algae [1,2,81,82]. Our conceptualization of the developing sunflower seedling also draws attention to the relationship between sucrose catabolism and osmo- and turgor-regulation (Fig. 10B). We conclude that, in developing sunflower hypocotyls, organ growth consists of an interaction between cell division and subsequent cell enlargement by the plastic deformation of the growth-controlling peripheral wall(s), associated with turgor maintenance via a tightly regulated catabolism of imported sucrose. We suggest that this disaccharide serves not only as a metabolite, but also as a signalling molecule, which originates in the cotyledons and is transported via the phloem through the hypocotyl into the roots, where it triggers important aspects of organ development [83]. Finally, we point out that our data pertain to seedlings grown under controlled laboratory conditions, whereas, in nature, plants are exposed to a steadily changing environment. In addition to night/day-(variable) sunlight cycles, shading within populations is perceived via photoreceptors (phytochromes, cryptochromes) and avoided by the promotion of stem elongation [84]. For instance, sunflower hypocotyls gradually increase their rate of elongation as the red-to far-red-ratio decreases [85]. These plastic modulations of stem elongation are mediated intracellularly by interacting transcription networks involving phytochromeinteracting factors and phytohormones (auxin, gibberellins, and brassinosteroids) [86–89]. Several studies indicate that, in sunflower hypocotyls, auxin (synthesized via a simple two-step pathway [90,91]) is the dominant light-modulated growth regulator. However, more experimental work is required to further support this “active phytochrome-auxin-growth” hypothesis of stem elongation [88,92]. Acknowledgements The cooperation of the authors is supported by the Alexander von Humboldt-Stiftung (AvH), Bonn, Germany (AvH Fellowship Stanford/USA 2011/12 to U.K.). We thank Prof. Winslow R. Briggs, Department of Plant Biology, Carnegie Institution for Science, Stanford, CA 94305, USA for the provision of lab space and consultation. References [1] K.J. Niklas, U. Kutschera, The evolutionary development of plant body plans, Funct. Plant Biol. 36 (2009) 682–695. [2] K.J. Niklas, U. Kutschera, The evolution of the land plant life cycle, New Phytol. 185 (2010) 27–41.

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