Although roots are sometimes neglected by researchers and called the ‘forgotten half’ or ‘hidden half’ of plants, root apical meristems have been studied extensively for two reasons. First, roots are viewed as a simpler system than shoot meristems – the root meristem is much more accessible than the shoot meristem which is ensheathed by developing leaves. Second, complicating lateral structures arise from the terminal shoot meristem (leaf and bud initials) but not from the terminal root meristem, which produces cells solely for the primary axis. This is where the simplicity ends. A primary root meristem generates two tissues simultaneously, the main root axis extending proximally towards the shoot, and the root cap pushing relentlessly forward into the soil, succumbing to sloughing and hence rapid turnover. The detailed organisation of root meristems, which we consider here primarily from the view of the cell biologist, reveals deeper complexities and questions of cell determination.

Lateral root meristems enable generation of massive networks of fine roots. The evolutionary processes which led to root systems of a very branched nature (e.g. grasses) through to coarse unbranched root systems (e.g. orchids) are a fascinating basis for further research into control of root branching. In addition, molecular intervention is giving us new plant forms which can be used to unravel the controls on root development and branching.

Root meristem anatomy

Primary roots arise through controlled cell divisions in the apical meristem and subsequent expansion and differentiation of these cells. Root formation (rhizogenesis) is usually extremely rapid: daughter cells exiting the meristem may be found 24 h later in a fully differentiated structure (e.g. phloem), even though further modifications to cell function are still possible (e.g. formation of an exodermis).

Critical steps in setting up dimensions and thickness of the root axis, and supply of cells to the zone of elongation, are the rate and position of cell divisions in the meristem. This can be appreciated from the two-dimensional view of a developing radish root in Figure 7.3. Divisions can be in any of three planes, either anticlinal (normal to the root axis), periclinal (tangential to the root axis) or radial to the axis. These divisions will give rise, respectively, to increased root length, increased root thickness (more layers of cells through the root), or increased root circumference. The apical meristem supplies all the cells for the primary root axis and the consequences of the planes of cell division are evident long after meristematic activity ceases.

Separate cell divisions at the leading edge of the root meristem generate a root cap which extends forward as a protective structure. The central cells of the root cap are often oriented in longitudinal arrays (columella) and are destined for rapid attrition as the ‘advancing’ soil particles slough off the surface layers. These cells also fulfil a vital chemico-physical role by secreting a glycoprotein-rich mucigel which reduces friction between root and soil matrix. Root caps advance at a dramatic speed: a root might elongate by 5 cm per day and new root cap cells can be pushed in advance of the apex of the primary axis at about the same rate. This means that the leading tip of a primary root, supplied with new root cap cells, advances through the soil at up to 60 µm min^–1. The sloughing off of roughly one cell layer per hour may explain our observation that the root cap does not increase in size over time. Intriguingly, root caps are still conspicuous in roots grown in nutrient solutions but still never dominate the primary root axis, so we deduce that sloughing off may induce a feedback mechanism that upregulates root cap meristem activity.

Another remarkable feature of root apices is the quiescent centre, a paradox at the heart of the meristem (Figure 7.4). The quiescent centre is a zone of relatively inactive, slowly dividing cells, numbering about 500–600 in a mature maize root. Its discovery (Clowes 1959) involved studies on mitotic frequencies, thymidine incorporation into nuclear DNA (Figure 7.4b), and ploidy after colchicine treatment. These led to a radical change in view of plant roots. The quiescent centre functions as a reserve of cells which can survive stresses and provide cells to a regenerating meristem. Recovery from surgical removal of parts of the meristem and irradiation to destroy dividing cells supports this concept. Likewise, short determinate lateral roots often lack a quiescent centre, suggesting it is closely tied to sustained indeterminate development.

Passage of cells from meristem to differentiated structures has been studied in simple roots such as ferns in which a single apical cell can be the progenitor of all root cells. Higher plants such as maize or beans have more complex roots, but the whole root can still be traced back to as few as 12 cells in the middle of the quiescent centre (Lyndon 1990). Cell destiny appears to follow predictable patterns, suggesting the notion of clonal development in which cell fate is fixed from the first divisions in the meristem. This view is under challenge from experiments using laser ablation. Individual cells, or groups of cells, can be eliminated by laser treatment, then the behaviour of adjacent cells is followed to see how the meristem is organised. This demonstrates that cells have considerable scope for taking over the meristematic role of their nearest neighbours. However, the process depends on physical contact between dividing cells and their daughter cells, which suggests a local transfer of information. The implication is that cell fate and the asymmetric divisions which give rise to various cell lines are regulated at a tissue level, but we do not yet know the nature of the mobile signals which might program cells in the meristem. We next turn to the fate of cells in their temporal journey from division to differentiation.

Cells divide in planes which are identifiably targeted to become the various root tissues even before all cell divisions are complete. For example, cells giving rise to the stele are generally clustered around the axis of the root, proximal to the quiescent centre (Figure 7.4a), while those giving rise to outer tissues (endodermis, cortex and epidermis) are peripheral to the pre-stelar cells. In roots of Arabidopsis, which has become a favoured plant for this work, the numbers of cells which generate individual tissues (e.g. eight meristematic cells generate eight cortical files) are known and the order of divisions giving rise to tissues such as pericycle, cortex and endodermis have been defined.

The changes which cells undergo in root meristems are profound; mitotic activity is most rapid in the distal regions (with a mitotic index of up to 23% in some files) but the mitotic cycle slows dramatically within 0.5–1.0 mm from a wheat apex. Surprisingly, mitotic frequency in adjacent cell files can vary widely (Figure 7.5; Table 7.1).

The rate and plane of cell division and subsequent rate of cell elongation determine the rate of delivery of new cells to mature root tissues. The coordination of cell flux is presumably under tight control, achieving the final anatomical outcomes recognisable as mature roots – single layers of pericycle and endodermal cells, long conducting vessels and epidermal cells are some examples. The role of growth in cells exiting the meristem and the direction of expansion are major factors in rhizogenesis, with a 30- to 150-fold volume expansion required to generate the primary axis.

Lateral roots

Lateral roots are important in nutrient and water uptake (Chapter 4.1). The cellular reorganisation which leads to lateral root primordia forming and developing into new axes starts with a latent meristematic activity in the root pericycle being de-repressed and cell divisions resume. Occasionally, endodermal cells are also recruited. Periclinal divisions underlie the out-growth of cells and disruption of the outer tissues of the root. However, before the cortex and epidermis have been penetrated by the young lateral root, it has formed its own terminal meristem and root cap. The new organ is thus prepared for growth in the external matrix. Lateral roots formed from the pericycle must breach the endodermis of the parent root. How is this achieved without rupturing of the Casparian strip and preventing outflow of concentrated nutrients to the cortex? Dyes which penetrate only the apoplasm have shown that endodermal disruption is a transient feature of lateral root growth, but the consequences are not well understood.

Lateral roots generally do not form within 1 cm from the terminal apex, and almost never in the zone of elongation. This makes sense as laterals in the growing zone would act as barbs impeding growth of the primary axis through the soil. An exception which supports this view comes from Eichhornia (water hyacinth) which does produce laterals in the elongation zone very near the root tip, but because of its aquatic environment this does not interfere with growth.

Shoot apical meristems are minute yet complex structures that are ensheathed within new developing leaves or bracts. A vegetative meristem gives rise to leaves or other organs, for example thorns and tendrils. Axillary buds are themselves complete shoot meristems from which branches are produced (cf. lateral roots described above). In angiosperms, when a plant shifts from vegetative to reproductive growth some meristems undergo a transition to the reproductive state and give rise either to multiple flowers in an inflorescence, as in mango (Figure 7.6a), or to a single terminal flower, for example a poppy or waterlily (Figure 7.6b). All axial growth from meristems, be they vegetative or floral, is continuous or indeterminate until topped by the formation of a flower. When this occurs, floral organ primordia arise in whorls from the shoot meristem and differentiate into the familiar sepals, petals, stamens and carpels.

Sometimes, however, the indeterminate inflorescence meristem reverts back instead to a vegetative status. Think of a bottlebrush or a pineapple with a leafy axis extending beyond the flower or fruit (Figure 7.7). What determines whether a meristem is vegetative or floral? In many species, environmental signals cause the switching between vegetative and floral states and a picture is now emerging of the genes and molecular mechanisms responsible for defining structures that are generated by meristems.

Although meristems function as generic sources of cells for differentiation into organs, each type of meristem is programmed to produce only certain structures. Across all species, there is a small, finite range of these structures, yet we observe an amazingly diverse array of final vegetative and floral morphologies. A leaf is always recognisable as a leaf but consider the vast structural differences between a pine needle, a waterlily pad and a tree fern frond. The generation and spatial patterning of plant organs are determined by early events within the vegetative meristem. This precise positioning of organs around the shoot meristem is called phyllotaxis. Later in development, a dramatic meristematic switch will give rise to a terminal inflorescence, often with an abrupt change in patterning of organs. Phyllotaxis also applies to floral structures, for example spiral patterns of scales of a pine cone or bottlebrush flowers (Figure 7.7).

Organ spacing is a final determinant of shoot appearance. For example, leaves forming a rosette as in Arabidopsis are separated by short internodes compared with longer internodes intervening between whorls of leaves of a blue gum seedling. The resulting morphologies are strikingly different. The question of what determines phyllotaxis and internode length is discussed later.

Shoot meristem anatomy

Shoot meristems are small, with a dome typically of 100–300 µm in diameter consisting of no more than a few hundred cells. In the 1970s, Williams (1974) published elegant reconstructions of shoot meristems derived from serial sections (Figure 7.8) which revealed the extent of variability in the shape and dimensions of shoot apices. However, the overriding organisation is of a central dome with groups of cells partitioned off from its periphery to form either determinate organ primordia or secondary meristems (axillary buds). Some cells in between are not destined to become primordia and will instead later become the internodes of the axis. Different models have been proposed to describe the regions of shoot meristems. In a functional sense, vegetative meristems have three main components: the central zone, peripheral zone and the file meristem zone, all of which tend to disappear or become indistinct in infloresence meristems (Figure 7.9).

Superimposed on this functional zonation are usually three distinct cell layers which give rise to separate cell lineages (Kerstetter and Hake 1997). These cell layers, designated L1, L2 and L3, are distinguishable by their positions in the meristem and their pattern of cell divisions, and are evident in both vegetative and reproductive meristems. Surface cells of L1 divide anticlinally while within the meristem and during sub-sequent differentiation of the organs. Not surprisingly, they form the epidermis. Within the apical dome, the plane of cell division within L2 is also purely anticlinal, but later on during organ formation divisions occur in other planes. In contrast, cells of the deepest layer, L3, divide in all planes. The two inner layers, L2 and L3, contribute cells to form the body of the plant with the proportion of cells derived from each layer varying in different organ types. Although the cell lineages produced by each layer usually contribute to distinct regions within each organ, invasion of cell derivatives of one layer into another has been observed.

Invading cells differentiate in accordance with their new position, which we interpret to mean that developmental fate of cells appears to be governed more by position than by cell lineage. However, meristematic cells may already be functionally distinct as evidenced by patterns of gene expression which reflect the layered cellular organisation (Figure 7.10).

In order to maintain the very precise organisation of vegetative meristems over long periods, or to accommodate rapid changes during flower formation, some signalling process must exist to coordinate division between the cell layers. Evidence for such signalling has been established through the development of chimeras, where genetically different cell types exist together in a single apex, yet still achieve normal developmental patterns.

Phyllotaxis and internode length

We previously raised the question of what determines phyllotaxis and internode length. Organs derived from the shoot meristem can arise in whorls (two or more organs simultaneously at one node), alternately (two files displaced by 180° with a single organ at each node) or in spirals (each organ displaced from the previous one by approximately 137° with a single organ at each node). These organs may be separated by very short or long internodes. Phyllotaxis patterns are usually stable, but often change abruptly with floral induction or when seedlings undergo transition to their mature morphology. In many species of Eucalyptus, this ‘phase change’ from juvenile to adult is very striking and is accompanied by a change from whorled to alternate or spiral phyllotaxis. How is the change in phyllotaxis effected? We gain some insight from experiments on chrysanthemum meristems where application of the inhibitor of polar auxin transport TIBA (tri-iodo benzoic acid) induced changes in internode length and displacement angle between leaf primordia. The data are consistent with a change from 137° spiral (control) to alternate (50 ppm TIBA) phyllotaxis and are presumed to result from increased concentrations of auxin in the meristem resulting from inhibition of transport away from the existing primordia. Meristems are deduced to be sites of auxin synthesis (Schwabe and Clewer 1984). One interpretation is that each primordium acts as a field with a defined radius of inhibition preventing other primordia from initiating too close to it. Another option is mechanical control by pressure and tension gradients within the meristem (Green et al. 1996). Supporting evidence comes from experiments on cells in tissue culture in which applied pressure altered their morphogenetic patterns. At a whole-plant scale, tension and compression wood in trees are further examples of specific developmental responses to physical forces.

Meristems as templates for morphogenesis

The location and activity of individual meristems give rise to the diverse morphologies we recognise within the Plant Kingdom. Palms and grass trees have a distinctive morphogenesis with the entire shoot canopy produced from the activity of a single apical meristem. Removal of the crown of a coconut palm inevitably kills the whole plant. The roots of palms and grass trees are also extraordinary in that they grow and senesce in a seasonal pattern which confers tolerance to poor soils and fire.

In contrast, woody trees produce complex shoot morphologies through combined activity of terminal and lateral apices. We see the product in the height and diverse branching pattern of large trees. Australian eucalypts show a diversity of shoot forms, ranging from the single slender trunk of a mountain ash or karri, topped by a branched canopy, to the multiple trunks of mallee eucalypts. The branching form of mallee species is determined by simultaneous activity of many apical meristems. Similarly, excavation of roots of large trees has often revealed complex branching patterns which enable effective exploration of large volumes of soil and extraction of water and nutrients. In the case of Eucalyptus marginata, a root system arises from strong meristematic activity in the surface levels of the root system as well as proliferation of deep sinker roots. Some species within the Proteaceae can form proteoid (cluster) roots that are adapted to nutrient-poor soils.

Grasses have a distinctive morphology which arises from the local activity of intercalary meristems. These meristems give rise to semi-autonomous plants called tillers which comprise leaves, stems and reproductive parts and are subtended by nodal roots. Cell divisions within the intercalary meristem are developmentally responsible for the characteristic morphology of grasses, a family that is well adapted to herbivory.

Mary Byrne, School of Biological Sciences, University of Sydney

Leaves are the main photosynthetic organs of vascular plants, and their optimisation for conversion of light to chemical energy production has resulted in a striking array of shapes. There is enormous variation in the shape of leaves of different plant species, of leaves within species, and for some species there is variation between juvenile and mature leaves on an individual plant. In the most general type of dicot leaf, there is a basal stalk or petiole and a distal flat blade or lamina. The leaf forms this structure through development in three axes; the proximal-distal (tip to base), the adaxial-abaxial (top to bottom) and the medial-lateral (middle to margin) (Figure 7.11). The adaxial-abaxial axis is evident at early stages of development of the leaf primordium. The adaxial side of the leaf is the side nearest to the shoot meristem and the abaxial side of the leaf is the side further from the meristem (Figure 7.11). These will eventually become the upper and lower sides of the leaf.

The resulting basic leaf shape is a flat, planar structure but modifications during development result in variations on the basic shape. For example, other shapes include leaves that are flattened along a lateral dimension or are radial, such as tendrils of Pisum sativum (pea) leaves or spines in many cacti. Varying degrees of serrations and lobing at the leaf margins also contribute to differences in leaf shape. For instance, within Australian Banksia, B. grandis leaves have deeply indented margins, whereas B. integrifolia leaves have smooth margins (Figure 7.12).

In addition to these types of elaborations, the leaf lamina may be a simple undivided shape or may be complex and subdivided into leaflets. Complex or compound leaves are found in many species including tomato (Solanum lycopersicum) and pea (Pisum sativum) (Figure 7.13).

Different leaf shapes may be produced during the life of the plant, and may be influenced by environmental conditions. For instance many Eucalyptus species have short, broad juvenile leaves and narrow, elongated adult leaves. Leaves in Acacia species change with plant maturity where juvenile leaves have a dissected leaf lamina and adult leaves (which are a modified petiole, or phyllode), are simple in shape (Figure 7.14).

7.1-Ch-Fig-7.14.jpg

Figure 7.14 Branch of an Acacia species showing two leaf shapes on the one plant. The juvenile leaves are compound leaves and are subdivided into leaflets. The adult leaves (phyllodes) are simple in shape.

Leaf initiation

Leaves initiate in peripheral regions of the shoot meristem (Figure 7.15). Sites of initiation occur where the hormone auxin forms a localised maximum. Local high levels of auxin are achieved through the action of a membrane-localised auxin efflux carrier PIN-FORMED1 (PIN1). The distribution of PIN1 to the plasma membrane on just one side of cells leads to directed flux of auxin away from sites where organs have already initiated and toward the site where organs will next form (Figure 7.15). Coincident with high levels of auxin, class I homeodomain transcription factor KNOX genes are down-regulated in initiating organs. Class I KNOX genes are expressed in the shoot meristem and are essential to maintain growth of the shoot meristem (Figure 7.15).

Each initiating leaf primordium is flanked by a boundary of slowly dividing cells and this boundary serves to delineate the leaf from the meristem and from adjacent leaf primordia. Several members of a transcription factor gene family known as CUP-SHAPED COTYLEDON (CUC) are expressed at this boundary and downregulation of CUC genes results in fusion of adjacent organs. This reflects the importance of the meristem-organ boundary in leaf development. As we see below, auxin, as well as the KNOX and CUC genes are also involved in determining leaf shape.

Proximal-distal axis

Following initiation leaf primordia go on to develop as discrete organs through cell division, cell expansion and cell differentiation. Leaf primordia become defined morphologically through cell divisions that lead to outgrowth from the meristem and hence formation of a proximal-distal axis. In the model species Arabidopsis thaliana (Arabidopsis), as the leaf grows a gradient of maturation occurs along this axis so that cells cease division and undergo terminal differentiation firstly in the distal region and lastly in the proximal region of the leaf. The leaf proximal-distal axis may have distinguishing features of lamina and petiole as in Arabidopsis and many dicotyledonous plants. One variation of this shape occurs in monocotyledons, which includes the grasses such as wheat, barley, rice and maize. Strap-like leaves of many monocotyledons have a distal blade and a proximal sheath, which wraps around the stem. The blade and sheath are separated by specialised cells that form a hinge, and this hinge allows the blade to bend away from the sheath and stem to be exposed to light.

Adaxial-abaxial axis

The adaxial-abaxial axis is evident at early stages of development of the leaf primordium. The adaxial side of the leaf is the side nearest to the shoot meristem and the abaxial side is furthest from the meristem (see Figure 7.11 above). Experimental separation of initiating leaf primordia from the shoot meristem results in loss of adaxial fate. Amazingly this loss of adaxial fate leads to development of an abaxial, radial leaf. This indicates that adaxial fate is determined either by contact with the meristem, or by signalling from the meristem.

Most flat leaves have adaxial-abaxial polarity where the two sides of a leaf have distinct morphological features. On the leaf surface, adaxial and abaxial epidermal cell size and shape, stomata density, or the presence of elaborated cell types such as hairs may be different. Vasculature often has a discrete adaxial-abaxial polar arrangement with adaxial water-conducting xylem and abaxial nutrient-conducting phloem (Figure 7.16a). Internal mesophyll cells of the leaf may also display differential arrangement along the adaxial-abaxial axis. In some species, the adaxial domain has closely aligned and chloroplast-rich palisade mesophyll cells whereas the abaxial domain has dispersed spongy mesophyll cells. Mature leaves are typically oriented with the adaxial side facing the light and the abaxial side away from the light. As such, distinct cell types along the adaxial-abaxial axis serve to optimize light capture and gas exchange for photosynthesis.

On top of this layer of regulation, adaxial and abaxial fates are each determined by specific genes. A reduction in expression of genes required for either adaxial or abaxial fate results in development of leaves that comprise only the opposite fate. Furthermore these leaves are radial. Together such observations have led to a fundamental tenet of leaf development, whereby adjacent adaxial and abaxial tissue types are essential to development of an adaxial-abaxial axis and hence to formation of a flattened leaf.

Key genetic determinants of adaxial fate include class III HD-ZIP transcription factors (Figure 7.16b). These proteins have a homeodomain, which is involved in binding DNA, a leucine zipper domain, which is involved in protein-protein interaction, and a sterol-binding domain that is predicted to bind small lipid molecules. In Arabidopsis, there are five genes encoding class III HD-ZIP transcription factors. Three of these genes, PHABULOSA (PHB), PHAVOLUTA (PHV) and REVOLUTA (REV), have a significant role in leaf development. Transcripts of all three genes are confined to the adaxial domain through transcriptional and post-transcriptional repression in the abaxial domain. For example, PHB, PHV and REV transcripts are all targets of the microRNAs miR165 and miR166, which are expressed on the abaxial side of the leaf. In mutants with reduced class III HD-ZIP activity adaxial fate diminishes. Conversely, gain of class III HD-ZIP expression in the abaxial domain results in conversion of the abaxial side of the leaf to adaxial fate.

Two other transcription factors, ASYMMETRIC LEAVES1 (AS1) and ASYMMETRIC LEAVES2 (AS2), form heterodimers and promote adaxial fate, in part through repression of miR165/166. Abaxial fate is promoted by additional factors, including KANADI (KAN) proteins, which are part of a plant-specific group of transcription factors. In Arabidopsis, four KAN genes have overlapping roles in leaf development. KANADI proteins are expressed on the abaxial side of the leaf and are necessary and sufficient for specifying the abaxial fate. It is also known that KAN1 directly represses AS2. Other abaxial factors are AUXIN RESPONSE FACTOR (ARF) proteins, which bind to promoter sequence elements in auxin responsive genes and regulate plant response to auxin. The two ARF genes, ARF3 and ARF4, are repressed on the adaxial side to the leaf by a set of small RNAs known as tasiR-ARF. The AS1-AS2 module directly represses ARF3. Thus the emerging picture is that repressive interactions between adaxial and abaxial genes helps to set up the adaxial-abaxial axis of the developing leaf (Figure 7.16).

Mediolateral axis

The mediolateral axis of leaves extends from the margin to the mid-region of the leaf, which is characterised by the presence of the main vein or midrib. Most leaves are bilaterally symmetrical and the two halves of the leaf, from margin to midrib, form mirror images. Leaf width is determined by the degree of lateral expansion along this axis. However, the relationship between the leaf margin and the midrib depends on cell recruitment from the shoot meristem during the process of initiation. Leaves of dicots initiate from a limited region of the shoot meristem so that the margins arise from cells close to the mid-region and lateral expansion of the leaf occurs post-initiation. On the other hand, leaves of monocots such as grasses, recruit cells from around the entire circumference of the shoot meristem. In this case, the leaf margins are derived from cells that are distant from the mid-region and consequently this early lateral expansion results in the leaf primordium wrapping around the entire shoot meristem.

Margins

Relatively simple modifications of the basic leaf shape are achieved through changes in growth dynamics at the leaf margins. In this way, leaves can develop marginal teeth or serrations or may be lobed. Leaf shape is more dramatically modified in compound leaves, where the blade is divided into reiterating units or leaflets. Multiple orders of subdivision occur when leaflets themselves are further subdivided into leaflets, for example in the compound leaves of tomato, which may have primary and secondary leaflets. Surprisingly, studies on the simple, serrated leaf of Arabidopsis, the lobed leaf of the Arabidopsis close relative Cardamine hirsuta, and the compound leaf of tomato have found that genes involved in setting up leaf initiation from the shoot meristem also function to modify the shape of the leaf lamina. The class I KNOX genes are important for modified leaf shape in many species. These genes are not expressed in the simple leaf of Arabidopsis but are expressed in the lobed leaf of Cardamine and the compound tomato leaf. Misexpression of class I KNOX genes in Arabidopsis leaves generates lobed leaves and increasing expression of these genes in Cardamine and tomato results in more lobing or more leaflets. Likewise, decreasing class I KNOX gene levels in Cardamine leads to a more simple leaf shape. Thus the degree of leaf complexity can depend on the expression of growth promoting class I KNOX genes. In addition to KNOX genes, auxin and CUC genes play central role in elaboration of leaves. In compound leaves of tomato and lobed leaves of Cardamine, high levels of auxin occurs at discrete sites along the developing leaf margin, where leaflets or lobes will form. Furthermore CUC gene expression marks the boundaries between initiating leaflets. At these sites CUC genes act to repress growth thereby promoting separation of leaflets. Altering the spatial distribution of auxin or CUC in the young developing leaf leads to reduced leaflet production. Less intricate elaborations of leaf shape, such as leaf serrations, also involve auxin and CUC genes. In developing leaves of Arabidopsis, the margins form periodic auxin maxima flanked by boundaries of CUC gene expression (Figure 7.17).

Together this promotes the formation of outgrowths at the leaf margin and so modulates leaf shape. Differences between the growth potential and intricacy of interactions between transcription factors and hormones likely determine whether the leaf will develop with modest outgrowths in the form of serrations, more prominent lobes or establish a compound leaf with distinct leaflets.

Further reading on leaf development

Bar M, Ori N (2015) Compound leaf development in model plant species. Curr Opin Plant Biol 23: 61-69

Bowman JL, Floyd SK (2008) Patterning and polarity in seed plant shoots. Ann Rev Plant Biol 59: 67-88

Braybrook SA, Kuhlemeier C (2010) How a plant builds leaves. Plant Cell 22: 1006-1018

Byrne ME (2005) Networks in leaf development. Curr Opin Plant Biol 8: 59-66

Byrne ME (2006) Shoot meristem function and leaf polarity: the role of class III HD-ZIP genes. PLoS Genet 2: e89

Chitwood DH, Nogueira FT, Howell MD et al. (2009) Pattern formation via small RNA mobility. Genes Dev 23: 549-554

Efroni I, Eshed Y, Lifschitz E (2010) Morphogenesis of simple and compound leaves: a critical review. Plant Cell 22: 1019-1032

Fukushima K, Hasebe M (2014) Adaxial-abaxial polarity: the developmental basis of leaf shape diversity. Genesis 52: 1-18

Hay A, Tsiantis M (2010) KNOX genes: versatile regulators of plant development and diversity. Develop 137: 3153-3165

Hepworth SR, Pautot VA (2015) Beyond the divide: Boundaries for patterning and stem cell regulation in plants. Front Plant Sci 6: 1052

Machida C, Nakagawa A, Kojima S et al. (2015) The complex of ASYMMETRIC LEAVES (AS) proteins plays a central role in antagonistic interactions of genes for leaf polarity specification in Arabidopsis. WIRES Develop Biol 4: 655-671

Mentink RA, Tsiantis M (2015) From limbs to leaves: common themes in evolutionary diversification of organ form. Front Genet 6: 284

Sluis A, Hake S (2015) Organogenesis in plants: initiation and elaboration of leaves. Trends Genet 31: 300-306.

The cambial zone includes a tier of meristematic cells between the developing wood (xylem) towards the inside of the stem and bark (phloem) tissue towards the outside (Figure 7.19). Cell divisions in this region either occur in parallel to the stem axis (periclinal) adding cells to a radial file of cells, or perpendicular to the stem axis (anticlinal) adding new cells to the initiating layer and thereby allowing for the establishment of new radial files. The cambium proper is formed by a layer or layers of initiating cells (cambial initials) that undergo mostly periclinal, but also anticlinal division to give rise to radially aligned files of secondary tissue. These initials are enclosed on both radial walls by xylem and phloem mother cells, respectively. Mother cells have a higher differentiation state than initials as they are destined to become either xylem or phloem elements, whereas true cambial initials retain the ability to produce both. Xylem and phloem mother cell divisions make up the majority of periclinal divisions in this zone and cambial initials replenish mother cells when those are lost from the cambial zone. Most anticlinal divisions are contributed by cambial initials resulting in the formation of new radial files and supporting the increase in girth of the growing tree stem.

Meristematic cells in the cambium include fusiform initials and ray cell initials, which are responsible for the formation of specific cell types in both xylem and phloem tissues (Figure 7.19b). Fusiform initials give rise to longitudinally aligned cells including vessels, tracheids and fibres towards the inside of the stem (in gymnosperms only tracheids are formed) and sieve tube elements towards the outside. They are large (compared to ray initials), longitudinally elongated and highly vacuolated. Ray initials on the other hand give rise to transversely aligned cells, primarily ray parenchyma cells, and occur as scattered aggregations within the cambial zone. The ratio of fusiform to ray initials can vary within the cambial zone depending on plant species, age, environmental factors and growth rates. For example, in angiosperms, fusiform initials on average make up 60-90% of the initials within the cambium. While in some species the cambium is comprised entirely of fusiform initials in others they make up as little as 25%.

Fusiform initials in the cambium occur either in a storied arrangement, where cells are laterally aligned, or non-storied, where they are arranged irregularly and often overlap. Divisions in a storied cambium generally occur synchronously resulting in lateral alignment and are typical of some small woody plant species, whereas divisions in a non-storied cambium occur asynchronously and are typical in trees. A storied cambium is believed to represent a more advanced evolutionary arrangement. Fusiform cell derivatives undergo variable amounts of cell expansion during differentiation with radial width often doubling and longitudinal length remaining more or less similar except for some longitudinal extension at the tip (expansion zone in Figure 7.19b). Longitudinal extension, particularly in xylary elements, occurs in the form of intrusive (or sliding growth) which involves the extension of tapered end walls of derivatives into the available space around them. Consequently, the end walls intertwine with end walls of other cells forming strong bonds that increase the mechanical strength of the stem.

The arrangement of rays in angiosperms is highly variable and can be uniseriate (1 cell wide), biseriate (2 cells wide) or multiseriate (3 or more cells wide) while in gymnosperms rays are exclusively uniseriate. Ray initials do not undergo much elongation during differentiation and have been observed to unite frequently, both vertically and laterally to form aggregate rays as well as to split by cell invasion of fusiform initials. The size of ray initials tends to remain constant throughout the life of the plant but their derivatives show a gradual increase in size with age and distance from the pith. Trans-differentiation from a fusiform initial to ray initials can occur in the cambium to maintain a balance between derivatives. This takes place either by segmentation or by progressive shortening of a fusiform initial, whilst trans-differentiation from a ray to a fusiform initial involves elongation via intrusive growth. The mechanisms that drive this process are largely unknown but proximity to other ray initials as well as the effects of plant hormones such as auxin and ethylene have been suggested.

Further differentiation of cambial cells results in the formation of vascular tissues, which are involved mainly in longitudinal transport of liquids and solutes up and down the plant stem. This is facilitated by the creation of long, unbroken tubular structures. Vascular tissue is made up of two distinct elements, xylem and phloem, with xylem being involved in the transport of water, soil derived nutrients and plant hormones from the roots up to the shoots, and phloem in the transport of organic material such as photosynthetic products, proteins, hormones and other regulatory molecules from the leaves to the roots. During secondary growth, these tissues, in particular xylem, are also involved in providing mechanical strength to the stem and as a storage sink for metabolites produced in other parts of the plant. Production of these conduits from the cambium is polarised with xylem derivatives produced centripetally (toward the inside of the meristematic layer) and phloem derivatives centrifugally (toward the outside of the meristematic layer).

Secondary vascular tissue, produced by fusiform initials in the cambium consists of axially aligned tracheary elements (fibres, tracheids, vessels) involved in xylem transport and axially aligned sieve elements (sieve tube cells, companion cells) involved in phloem transport. Ray and axial parenchyma cells are also products of meristematic activity in the cambium and occur in both xylem and phloem tissues. Ray parenchyma cells are aligned radially and axial parenchyma cells are aligned longitudinally. Both parenchyma tissues are involved in transport and storage.

In angiosperms longitudinal transport in the xylem occurs in vessels, which are specialised for liquid transport and arranged end on end to form a continuous, longitudinally aligned tube system. Vessel elements exhibit tapered end walls and thickened lignified cell walls. Prior to maturation and becoming functional, vessel elements undergo programmed cell death (PCD), a process during which cellular content is actively removed, which together with specialised end structures known as perforation plates allows for unimpeded flow between individual vessel elements. Vessels do not form completely straight parallel tubes but rather deviate from their axial path slightly to come in contact with other vessels creating a network of interconnected tubes that allow for movement between vessels through cell wall pits.Vessels are surrounded by longitudinally aligned fibre cells which have a role in providing strength to the plant stem. Compared to vessels, fibres are longer and narrower, have thicker, usually more heavily lignified cell walls, increased tapering of their end walls and they lack a perforation plate. Fibres also undergo PCD as they mature and contain simple pits on their cell walls that allow for minimal transport of liquids between adjacent fibres. Fibres are found most frequently in the xylem and, to a lesser extent, in the phloem where they are arranged in small bundles. In gymnosperms, both transport and strength are provided by a single xylary cell type, tracheids, which, in terms of morphology, are an intermediate form between a fibre and a vessel element. Tracheids are also found in angiosperm stems but to a much lesser degree. Transport of liquids between tracheids is via specialised cell wall openings called bordered pits which through passive opening or closing via acentral, thickened area of pit membrane (torus)can prevent gas or water movement (for example to prevent possible embolism).

Parenchyma cells are rectangular to almost isodiametric in shape and alive at maturity. They are involved in a variety of roles such as transport, signaling, storage and wound responses. Walls of parenchyma cells have a characteristically high number of plasmodesmata that allow for efficient cell to cell communication and transport. Ray parenchyma cells are the only cells within a tree stem that are aligned radially and play an important role in transport and communication between xylem derivatives, cambial initials and phloem derivatives. Similarly, axial parenchyma cells are the only live cells aligned longitudinally in the xylem. They are important for storage and transport in this direction while in the phloem they more often fulfill a storage function.

In the phloem, longitudinal transport occurs via sieve tube elements, which like vessels, are aligned end to end and form continuous tubes. Individual elements are connected by structures called sieve plates. Sieve tube elements undergo some elongation and cell wall thickening during differentiation but differ from vessels and fibres as they do not undergo lignification. Also sieve tube elements are alive at maturity and have adapted a highly modified cytoplasm to allow for free movement of solutes between cells. Here, so called P-proteins internally line all sieve tubes and in the event of wounding can seal sieve plates and allow for wound closure. Cytoplasmic modifications occur via the removal of large organelles including the nucleus, ribosomes, golgi bodies, the tonoplast (vacuoles) and a cytoskeleton as well as links of plasma membranes between longitudinally aligned cells. Sieve tube elements form an association with companion cells, which in angiosperms are derived from a single division of a sieve tube element during differentiation while in gymnosperms they are derived from separate lineages. Companion cells are believed to produce and secrete information molecules via plasmodesmata to associated sieve tube elements.

The most prominent cell division type in the cambial zone, making up approximately 90% of divisions in the cambial zone, is periclinal or additive division, which leads to the formation of highly organised files of radially aligned cells. Unlike in most cell divisions in plants where the cell plate forms across the shortest distance of the cell, cells undergoing periclinal division in the cambium have a cell plate that forms across its long axis. In this process the cell plate begins formation around the mid point of the cell and develops longitudinally until it connects with two lateral cell walls at the end of a cell, resulting in the formation of two nearly identically sized daughter cells. Anticlinal divisions or multiplicative divisions do not occur as frequently as periclinal divisions and are responsible for the formation of new radial files within the cambial zone capable of further periclinal divisions. Anticlinal divisions account for the increasing stem circumference as a result of radial growth as cell wall expansion of initials and their derivatives is not sufficient to compensate for this. This results in a higher frequency of anticlinal divisions during times of accelerated plant growth. The number of anticlinal divisions occurring in fusiform initials is often higher than the number required to compensate for an increase in stem circumference. Superfluous cells are either lost or squeezed out from the cambial zone or undergo trans-differentiation to form ray initials.

Wood formation (xylogenesis) comprises cell division, cell expansion, secondary cell wall deposition, lignification and finally programmed cell death, which occur in highly coordinated and defined ways in response to the environmental conditions imposed on the plant. In mature tree stems, fusiform cambial initials give rise to fibres, tracheids and vessel cells (only tracheids are formed in conifers) while ray cell initials give rise to ray parenchyma cells which, combined, form the ray system. Following division, but prior to the commencement of secondary cell wall deposition, the immediate xylogenic derivatives of fusiform cambial initials (i.e. xylem mother cells) undergo cellular expansion to reach their final form. The extent of this turgor driven expansion is regulated by a range of enzymes that assist in the association and disassociation of primary cell wall components including pectin, randomly aligned cellulose microfibrils (MF) and other non-cellulosic polysaccharides.

The beginning of secondary cell wall formation marks the end of cellular expansion with the production of new cell wall layers leading to cell wall thickening. This stage is characterised by the deposition of large amounts of aligned cellulose microfibrils to the inside of the primary wall in three layers, S1, S2, S3. These differ in their thickness and cellulose MF alignment, and also by the development of cell wall pits. The S2 layer is the most dominant of these layers, accounting for approximately 75-85% of the cellulose content in the secondary cell wall, and as far as wood properties are concerned, has the largest influence. For example ‘Microfibril Angle’ (MFA), one of the most important complex wood quality traits, refers specifically to the cellulose microfibril orientation in this layer of the fibre (or tracheid) cell wall. MFA is known to influence the stiffness and elasticity of individual cells as well as wood and wood products. During this process, the cell wall lignifies and finally undergoes programmed cell death (PCD) during which all cellular content is lost through the action of proteolytic enzymes.

In many tree species, a final developmental process, heartwood formation, occurs later during development. With progressing age, and with the continuous differentiation of some fusiform cambial initials into new vessels, older vessels towards the center of angiosperm tree stems no longer transport water and nutrients and the actively conducting sapwood develops into heartwood. The only cells that are still alive in the sapwood/heartwood transition zone are ray parenchyma cells. Here, they are believed to undergo programmed cell death, translocating polyphenolic substances into surrounding cell walls, leading to the discoloration of heartwood. Where ray cells of some angiosperm trees are directly associated with larger vessels, as a final developmental process, they can produce tylose plugs, decreasing wood permeability and preventing microorganisms from entering the center of the stem. A similar process occurs in sapwood in response to wounding in order to avoid embolism (Figure 7.20).

The deposition of lignified secondary cell walls marks the end of cellular expansion and is central to the process of xylogenesis and ultimately determines the functional properties of wood. Secondary cell walls consist of cellulose, lignin and other non-cellulosic polysaccharides which typically represent 90% of the wood dry weight and provide support for the cell. Other minor yet important contributions to cell wall structure and function are made by cell wall proteins and other compounds. Quantitative and qualitative changes in the composition of these elements can lead to changes in cell wall properties and are responsible for the large variation in wood properties observed within and between tree species.

Throughout the year, the cambium undergoes cyclic or induced dormancy and activity periods and wood properties show high levels of variability in response to seasonal, developmental, genetic and abiotic processes. In temperate regions, typically in the northern hemisphere, the vascular cambium undergoes seasonal or periodic dormancy in response to unfavorable and/or detrimental environmental conditions. This dormancy period is usually triggered by a decrease in photoperiod, temperature and water availability during the autumn months preparing the plant for adverse winter conditions and in deciduous plants is closely linked with leaf shed. Prior to the onset of dormancy as conditions begin to decline, cambial activity as well as the number of dividing cells is reduced and cell wall thickness in fusiform initial derivatives in the differentiation zone increases. These seasonal differences in the formation of woody tissues enable the distinction between earlywood (larger cell lumen and thinner cell walls) and latewood (smaller cell lumen, thicker cell walls) leading to the creation of growth rings. Ray initial derivatives on the other hand do not undergo such strong seasonal changes in morphology but their number in the cambial zone decreases during dormancy. Changes in wood properties are also observed between wood produced from early stem growth, juvenile wood, and wood produced later from an older cambium, mature wood. Abiotic stress can lead to reaction wood formation (for example in response to non-vertical growth; referred to as tension wood in angiosperms and compression wood in gymnosperms) displaying altered secondary cell wall characteristics and wood properties to ‘normal’ wood in an attempt by new cambial growth to realign the stem back to a vertical position.

As the plant stem increases in girth during secondary growth, the outermost protective layer of the stem, the epidermis, is gradually shed and replaced by a periderm. Peridermal tissue is produced by the cork cambium (or phellogen) which, like the vascular cambium, comprises a meristematic layer that produces derivatives, both centrifugally and centripetally, and undergoes periclinal and anticlinal division forming radial files of cells. Derivatives from the phellogen differ from those of the cambium both structurally and functionally and the rate of division from this meristem is often much lower than that observed in the vascular cambium. The periderm is comprised of three different tissue types; 1) the phellogen or cork cambium, the meristematic tissue of the periderm, 2) phellum or cork, the outermost protective layer of tissue, and 3) phelloderm, the living parenchyma formed toward the inside of the phellogen. The phellogen is made up of a single layer or tier of cells that are radially flattened, rectangular to polygonal in shape, with thin primary walls, dense cytoplasm and, in some cases, chloroplasts. Phellum cells divide centrifugally from the phellogen and are highly specialised, creating an imperious barrier to gases and liquids on the plant stem. Phellum cells are usually polyhedral in shape, but can be radially compressed and/or elongated in several planes (radial, tangential and longitudinal) and are arranged compactly, lacking intercellular spaces. Phellum cells develop a layer of suberin, an unsaturated fatty acid that impregnates the cell wall during development and gives them their impervious properties. Maturation of these cells ultimately results in cell death. In some genera, e.g. Eucalyptus, phellum cells can also be lignified. Phelloderm cells are living parenchyma cells that divide centripetally from the phellogen and are similar in appearance to the adjacent cortical tissue, being only distinguishable by their radial alignment with cells in the phellogen.

The periderm is continuously replaced during growth and the site and tissue type from which it develops differ between species. Initiation of a periderm occurs either uniformly as a ring around the plant, or in scattered groups which link up later via lateral spreading.

Once the first periderm is ruptured, initiation of the next periderm usually occurs from either cortical tissue or phloem parenchyma whereas later periderms form exclusively from phloem parenchyma. This process is repeated continuously during the life of the plant. The formation of a new periderm displaces a large number of cells, which die and are shed from the stem, thus a tree continues to increase in diameter.

The wall is an intricate network of polymers that, when capable of expanding, is termed the primary wall. The secondary wall is formed underneath the primary wall when it has finished expanding, and is thicker. The secondary wall is elastic and can shrink or swell but not expand irreversibly like the primary wall.

a) Structure of the primary wall – growing cells

Primary walls are composed predominantly of a complex array of polysaccharides (~90%) and some protein (~10%). In all cell types, rigid cellulose microfibrils are embedded in a gel-like matrix of non-cellulosic polysaccharides and pectins. These polysaccharides are intimately associated with one another, both non-covalently and covalently, and often with proteins and lignins. Walls are complex, diverse and dynamic, changing throughout the processes of cell division, growth and differentiation. The types of wall polysaccharides vary depending on the plant species, cell type and developmental stage (Doblin et al. 2010). In some woody tissues, the secondary wall can be very thick and make up more than half the total volume of the cell.

Growing cells are surrounded by thin walls (100 nm or less) that are sufficiently flexible to yield to the hydrostatic forces that drive growth. The primary walls are highly hydrated (~60% of wet weight) and in dicots and gymnosperms consist of a cellulosic network embedded in a matrix of complex polysaccharides, of which xyloglucans and pectic polysaccharides are most abundant (Figure 7.22). Primary walls of most monocots are organised in essentially the same way except glucans and glucuronylarabinoxylans predominate in the matrix phase and may have properties similar to xyloglucans and pectins.

The main wall constituent that allows the primary wall to expand is the gel-like matrix in which the cellulose microfibrils are embedded. Typically consisting of xyloglucans and pectins, attention is increasingly directed toward the pectins because mutant Arabidopsis plants (xxt1 and xxt2) still can grow even though they totally lack xyloglucans (Cavalier et al. 2008). On the other hand, the genes for pectins have proliferated during the evolution of land plants and Arabidopsis has 15 involved in the synthesis of linear pectins, and 10 more that resemble these genes. Some of their mutants are lethal, especially mutants in GAUT1 and GAUT4 that are the major ones coding for enzymes synthesising linear pectins (Caffal et al., 2009). This suggests a central role for the pectins in the wall. The cellulose does not contribute directly to the growth process and instead mostly strengthens the wall. As a polysaccharide consisting of a linear chain of several hundred to many thousands of β(1–4) linked D-glucose units, it spontaneously crystallises in cellulose microfibrils having about 20 cellulose chains assembled into long cables a few nanometers in diameter. These are cross-linked by hydrogen bonds to a matrix of hemicellulose and pectin along with a small amount of structural protein. The microfibrils have a high tensile strength approaching that of steel, and their orientation in the wall determines the shape of the expansion. When laid down in an orientation perpendicular to the long axis of the cell, lateral expansion is inhibited and the wall extends mostly lengthwise. The longitudinal extension stretches the matrix between the microfibrils, spreading the microfibrils apart. The orientation of the microfibrils thus controls cell shape while the matrix controls growth rate.

b) Structure of the secondary wall – fully grown cells

The secondary wall is laid down underneath the primary wall when it has finished expanding, that is, when the cell has stopped growing. The secondary wall is elastic, it can shrink or swell and undergo reversible but not irreversible expansion. In some woody tissues, the secondary wall can be quite thick and be over half the total volume of the cell.

Cells that have ceased enlarging and are required to withstand large compressive forces mature by depositing secondary walls up to several micrometers thick that increase the wall strength and reduce flexibility (Figure 7.23). During secondary wall development cellulose and matrix phase polysaccharides with a lower degree of backbone substitution, such as heteroxylans and heteromannans, are deposited in a highly ordered pattern. Together with the deposition of lignin this results in a dehydration of the wall material, which becomes increasingly hydrophobic in nature. In some cell types, lignin is also deposited throughout the wall during secondary development. Hydrophobic lignins overlie and encrust the cellulose microfibrils and matrix polysaccharides, and can also be covalently complexed with wall polysaccharides (Doblin et al. 2010). The character of these cells controls woodworking properties of stems of trees and can ward off pathogens that might attack the wood.

Osmosis makes cell expansion possible. Osmotic concepts were first understood by J. Willard Gibbs (1875-76) and demonstrated experimentally soon thereafter by Wilhelm Pfeffer (1876). Neither of these scientists knew how the process caused growth but Pfeffer (1900) sensed that the critical feature was turgor pressure (P). It is now known that P has to be low enough to create a water potential to bring water into the growing cell while at the same time being high enough to expand the wall. This dual role can be understood from the theoretical basis of osmosis (Figure 7.24).

Osmosis in a constrained compartment like a plant cell causes P to build up inside the cell until it becomes high enough to prevent net water entry. At that stage, the cell is in equilibrium with the solution in its surroundings. Inside the cell, P is the turgor pressure and it can be high (positive pressures of 0.5 MPa are common). Outside of the cell in the wall that is hydraulically connected to the xylem, P can vary considerably and tends to be negative during the day because the xylem water is under tension (negative pressure). The tension in xylem is of course transmitted throughout the apoplast. Bearing this in mind, it is possible to express mathematically the osmotic forces and flows across a membrane as:

\[ \frac{dV}{dt} = AL_{p}[(P_i - P_o) - (\Pi_i - \Pi_o)] \tag{1} \]

where V is the net volume of water moving osmotically across the membrane, \(t\) is the time, \(A\) is the area of the membrane, \(P\) is the pressure, \(\Pi\) is the osmotic effect of the solute concentration, \(L_{p}\) is the hydraulic conductivity of the membrane, and subscripts \(i\) and \(o\) refer to the interior (cytoplasm) and exterior (apoplast) of the cell.

It is important to point out that the water potential is \( \Psi = P – \Pi \) and Eq. 1 simplifies to:

\[ \frac{dV}{dt} = AL_{p} \Delta \Psi \tag{2} \]

where \( \Delta \Psi \) is the water potential difference between the inside and outside of the cell. At equilibrium, \(P\) builds until the net flux is zero and Eq. 1 becomes:

\[ (P_i - P_o) = (\Pi_i - \Pi_o) \tag{3} \]

Let’s see how Eq. 3 fits plant cells whose apoplast is hydraulically connected to the xylem. Starting in the xylem where the solution is very dilute (say \(\Pi_o\) = 0) and the wall solution is under tension (say \(P_o\) = -0.7 MPa), osmosis will move water into the cell as long as the cytoplasm is more concentrated (say \(\Pi_i\) = 1 MPa). As water moves in, \(P_i\) builds inside until it prevents water from entering. Ultimately, the \(P_i\) reaches 0.3 MPa whereupon the cell is equilibrated with its surroundings, as shown in Eq. 3 but expressed numerically in Eq. 4:

\[ 0.3 - (-0.7) = (1 - 0) \tag{4} \]

Notice in Eq. 4 that the P difference across the membrane equals the \(\Pi\) difference across the membrane. In other words, the pressure difference is in equilibrium with the osmotic effect of the solute. Of course, if insufficient water can enter the cell to build its turgor as high as 0.3 MPa, \(P_i\) decreases. If it falls near zero, we sometimes see the leaf “wilting”.

A major advantage of the water potential is that it describes the direction of water movement, taking osmotic properties and pressures into account in one symbol. According to the example above (Eq. 4), the water potential outside of the cell is -0.7 MPa (-0.7 + 0.0). Because outside and inside are in equilibrium, the water potential inside of the cell is also -0.7 MPa (0.3 - 1.0). In both places, \(\Psi\) is a negative number (like temperatures below zero).

Osmosis is the key to water entry into the plant. It not only causes water to enter the cells of the plant but also creates the tension in the xylem transmitted to the cell wall (apoplast) that pulls water through the root from the soil. In fact, this ability to pull water from the soil has to balance transpiration. In other words, water uptake by osmosis is necessary to prevent the plant from drying out. Larger \(\Delta\Pi\) create the potential for greater pull extending out into the soil. Wheat typically contains large \(\Pi_i\) (2.0 to 2.5 MPa) that allows it to obtain water from relatively dry soil. Maize generally has small \(\Pi_i\) (1.2 to 1.8 MPa) that makes it less able to obtain water from dry soil. Maize tends to be less drought tolerant than wheat. The reasons for the \(\Pi_i\) differences are unknown.

These ideas work for osmosis in plant cells that discriminate perfectly between solute and water (water moves through the membrane but solute does not). In healthy cells, most membranes are nearly perfect and negligible solute moves through (solute actually moves through on specific carriers but the amount is negligible osmotically). If solute crosses the membrane, it acts essentially the same as a water molecule, and the osmotic force Π is diminished. Places where this may happen are phloem termini where large amounts of solute are delivered to the developing embryo along with water. Developing grain would be an example. Another place is in air-dry seeds whose membranes have been dehydrated and are leaky to solute until the cells are rehydrated during imbibition, allowing the membranes to re-form.

So far, we have only considered the hydraulic conductivity of membranes of individual cells. Sometimes it might be useful to determine the conductivity of a whole organ such as a root system or leaf. For that an abbreviated form of Eq. 1 is typically used:

\[ \frac{dV}{dt} = L(\Delta\Psi) \tag{5} \]

where \(L\) is called the water conductance of the organ and the definition of the other terms remains unchanged. The reason we cannot use Eq. 2 is that the area for flow is not the plasma membrane of the cell. In effect, \(A\) is undefined and is included in \(L\) but the general principle governing flow is otherwise like that in Eq. 2. As you can see, the bigger the root system the higher the flow and thus \(L\) will tend to get larger as the organ grows.

Plants grow mostly by increasing the size of cells in enlarging regions. For a cell to become larger, the wall becomes irreversibly extended by \(P\). In effect, the wall in a growing cell is constructed so the polymers slip a little at high \(P\). In that situation, as \(P\) builds from osmosis, it never reaches a level where equilibration can occur. Instead, \(P\) can only build enough to cause irreversible slippage (yielding) of the wall and the wall compartment becomes bigger. This can be demonstrated by lowering \(P\). Even if \(P\) decreases as low as zero, the compartment remains bigger than it was.

Typically the \(P\) must be higher than a minimum before yielding occurs. It is possible to express this wall property with an equation similar to Eq. 2. Lockhart (1965) was the first to do so and described cell enlargement for \(P\)-driven growth as:

\[\frac{dV}{dt} = A \phi (P - P_{th}) \tag{6} \]

where \(\phi\) is the wall yielding coefficient, sometimes referred to as wall extensibility (m Pa^–1 s^–1) and \(P_{th}\) is threshold turgor pressure or yield threshold (\(P_{th}\)) above which the cell wall yields. The other terms are defined as before. It is obvious in Eq. 6 that \(P\) must be above \(P_{th}\) in order for the cell to enlarge. Also, as \(\phi\) becomes larger, the cell grows faster.

In tissues instead of individual cells, a similar equation applies because all the cells enlarge in concert. The enlargement of the whole organ is then described by:

\[\frac{dV}{dt} = m (P - P_{th}) \tag{7} \]

where everything is the same as in Eq. 6 except that \(m\) contains the wall yielding attributes for all the enlarging cells as well as the \(A\) term. Examples where this equation applies would be tissues of enlarging stem and root tips, hypocotyls and coleoptiles of geminating seeds, flower buds, and even growing fruits (grapes, tomatoes, maize kernels, etc).

Nevertheless, the inability of \(P\) to build enough for equilibration keeps the cell water potential lower than it otherwise would be. As a result the yielding walls create a lower water potential that is called “growth-induced” because mature cells that do not have such yielding walls (Figure 7.25). The growth-induced \(\Psi\) can move water from the mature cells into the elongating ones if no external supply is available. This probably explains how a stored potato can sprout in a cupboard because the cells in the bud develop yielding walls that create a growth-induced \(\Psi\) that moves water from the mature cells into the sprouts.

Usually the water source is in the soil or xylem (\(\Psi_o\)), and a slight modification to Eq. 5 can show this effect:

\[\frac{dV}{dt} = L (\Psi_o - \Psi_{growth-induced}) \tag{8} \]

Because the cells with yielding walls have turgor that is \(P = \Pi + \Psi_{growth-induced}\) (remember \(\Psi\) is negative), Eq. 7 becomes:

\[\frac{dV}{dt} = m (\Pi + \Psi_{growth-induced} - P_{th}) \tag{9} \]

and solving for \(\Psi_{growth-induced}\) in Eqs 8 and 9 gives:

\[\frac{dV}{dt} = \frac{mL}{m + L} (\Psi_o + \Pi - P_{th}) \tag{10} \]

This shows that growth depends on a water supply function (Eq. 8) and a water demand function (Eq. 7) that can be combined to give Eq. 10. A quick look at this latter expression shows that the growth rate of an organ is determined by the water potential of the supply, the osmotic effect of the solute minus the threshold turgor (which is too low to contribute to the growth rate) multiplied by a coefficient. In other words, the difference in water potential that we’ve seen before in Eq. 5 diminished by the threshold turgor that is inactive in the growth process provides the driving force for growth. When multiplied by the coefficient, \(\frac{mL}{m + L}\), the equation gives the growth rate of the organ.

The reason Eq. 10 is important is that it shows a plant organ grows not only because of the turgor properties of the tissue but also because water moves through all the small cells (\(L\)) of the entire organ. Meristems have dividing cells that are necessarily small. There are many wall and membrane barriers to traverse before water reaches all of the cells and allows them to grow simultaneously at a similar rate. The yielding of the walls controls the growth-induced \(\Psi\) needed for this water. In effect the turgor must be high enough for the walls to yield but low enough to create the growth-induced potential to supply water to the enlarging cells. The supply is often quite far away depending on the anatomy of the meristem and enlarging tissues. Consequently, large distances may be travelled by water for growth and the anatomy plays a large part in the growth rate of the organ.

It is worth looking at the coefficient a little more deeply. If the wall yields easily and m is large (say \(m\) = 10 units and \(L\) = 1 unit), the coefficient is 10/11 and the growth rate is controlled mostly by the ability of water to move into the enlarging tissues. If the reverse occurs (\(m\) = 1, \(L\) = 10), the coefficient is still 10/11 but the growth rate is controlled mostly by the yielding properties of the walls. Therefore, comparing the magnitude of \(m\) and \(L\) is the key to determining whether the demand (\(m\)) or the supply (\(L\)) function dominates the coefficient. A picture of these effects is shown in Figure 7.26 and methods are available for measuring not only \(m\) and \(L\) but also all the other terms in these equations (Boyer et al., 1985).

It is pretty obvious from this behaviour that simply putting water around a tissue creates a system with several interacting factors. The growth rate could be affected not only by \(P\) but also by \(m\), \(L\), \(\Pi\), or \(P_{th}\). In an effort to simplify this system, scientists have sought single cells that could be surrounded by water. From a practical point of view this allows \(P\) and \(m\) to be the main factors controlling growth and minimises the effect of \(L\), as shown in Figure 7.27.

One approach has been to use single algal cells large enough to measure \(P\) and growth (\(\frac{dV}{dt}\)) simultaneously. Chara corallina or Nitella flexilis are candidates because they have cells large enough for the measurements. They are naturally surrounded by fresh or brackish water and have rhizoids resembling roots. Gametes form in structures in the axils of branches analogous to flowers or cones in their land counterparts. In fact, genomic and morphological analyses consider these algae to be among the closest relatives of the progenitors of land plants. In the internode cells of Chara or Nitella, microfibrils are oriented normal to the cell axis and the walls expand mostly in length like many plant tissues (roots, stems, grass leaves).

Using the internodes of these species, \(P\) and growth rate (\(\frac{dV}{dt}\)) can be monitored simultaneously and changed so quickly that \(\Pi\) and \(P_{th}\) remain constant. This allows the \(P\) response to be rigorously determined. Moreover, the walls can be isolated without leaving the medium in which the algae are grown. The same measurements can be repeated without the cytoplasm. This is a great tool for observing the response to \(P\). When the growth of the live cells was compared with that in the isolated walls, they were similar but only for the first hour or so. After that, growth ceased in the walls but continued in the live cells even though the walls and cells had the same \(P\). Something was missing in the isolated wall that was being supplied by the live cells.

Considering that new wall material is supplied by the cytoplasm and missing in the isolated walls, if seemed reasonable to supply new wall constituents as though the cytoplasm had done so. Supplying pectin (a wall polymer) to the growth medium returned the growth rate of isolated walls to the rate in the live cells! This was unexpected but indicated the wall needed a supply of pectin in order to continue growing. The active pectin was a linear unbranched polymer of α-1,4-D-galacturonic acid sometimes with a small amount of rhamnose (usually 1-2%) that is normally synthesised in the cytoplasm and released to the wall by exocytosis. It becomes a prominent member of the wall matrix and forms a gel embedding the cellulose microfibrils.

The pectin gels because calcium ions bind to neighbouring pectin polymers. The cross-bridging forms junction zones with the polymers that are strong enough for the pectin to form a gel solid. The gel gets stronger with more cross-bridges. The new pectin from the cytoplasm removed some of the cross-bridges from the wall, weakening the wall gel and allowing the polymers to slip a little. This action occurred only when \(P\) was above \(P_{th}\), and only when temperatures were warm enough for growth. Figure 7.28 compares the turgor pressure and temperature responses of the live algal cells with those of land plants tissues.

A chemical mechanism has been proposed to account for this behaviour and is called a “calcium pectate cycle” (Figure 7.29). It seems possible that the chemistry might also occur in land plants. As long as pectin, calcium, and sufficient turgor pressure are present, the cycle should occur in pectin-containing walls. Pectins are among the most conserved components of cell walls during plant evolution, and the similarity in pressure and temperature response in these algae and land plants suggests a common mechanism in both. However, despite these intriguing similarities, definitive tests remain for the future.

Expansins

In other experiments a class of cell wall proteins, expansins, are proposed as potential agents for catalysing yielding in vivo (McQueen-Mason 1995). Figure 7.30 shows sharp gradients in growth along the hook of a cucumber hypocotyl that are paralleled by a gradient in extension of these tissues when stretched under acid conditions (Figure 7.30b) but not at neutral pH (Figure 7.30c). When tissues were killed by boiling, extension was blocked (Figure 7.30d). From these results, it seems that hypocotyl extension requires acid pH and non-denatured proteins. However, all the experiments were done in an extensometer with a uni-axial pull substantially less than the multi-axial tension exerted by \(P\). Since growth requires \(P\) above a threshold in order for walls to yield (Eq. 7), it is difficult to interpret this proposal. When greater uni-axial pull was used by Ezaki et al. (2005) to study hypocotyl growth in soybean hypocotyls, pectin chemistry appeared to determine growth rate. In fact, Zhao et al. (2008) give evidence that pectins may be the target of expansin action.

Notice that the growth mechanism in Figure 7.29 is entirely chemical, with no role for enzymes. It is difficult to hypothesize an enzymatic mechanism because of the requirement for \(P\) above a threshold. Enzyme activity is generally unaffected by these pressures and would continue acting regardless of \(P\). What is clear is that the biophysical consequences of instantaneous changes in \(P\) will be followed by a phalanx of biochemical events including wall polymer synthesis and altered gene expression, and rigorous methods will be required to distinguish enzymatic from biophysical hypotheses. For instance, sustained expansion of plant cell walls cannot be explained simply by inexorable wall hydrolysis; if it were, cell walls would weaken to breaking point during growth. The ‘setting’ of long-term cell expansion rates is likely to hinge on biochemical and chemical events underlying wall relaxation and reinforcement.

Cessation of cell wall expansion

Molecular events leading to cessation of wall expansion are even less well understood than those which initiate growth. For example, part of the growing region stops growing when water deficits occur around maize roots (Figure 7.31).

Clearly, the region farthest from the tip has stopped growing but \(P\) remains uniform throughout the zone. \(P\) is lower in the water deficient roots presumably because less water can be absorbed from the water deficient soil. A common view is that sufficient cross-linking develops to limit the extension of the matrix around cellulose microfibrils and prevent further wall expansion. Essentially, when a cell has reached its final dimensions its wall is ‘locked’ into a final, hardened conformation. From the description above, molecules with a specific role in growth cessation are thought to be exocytosed into cell walls, providing either substrates for cross-linkage reactions or enzymes catalysing cross-linkage of pre-existing wall polymers. Identification of cross-linkage reactions have led to a search for their presence in vivo. For example, ferulic acid residues in grass cell walls can cross-link to produce di-ferulic acid and potentially stiffen walls through formation of a polysaccharide-lignin network. Unfortunately, in rice coleoptiles the abundance of the di-ferulic form bore no relation to growth cessation. Also this form of stiffening might be difficult to distinguish from secondary wall deposition.

Secondary cell walls generally form after primary walls have ceased to grow but the familiar rigidity of secondary cell walls (e.g. wood) is mostly viewed as distinct from stiffening of primary walls. Lignification of primary walls commences earlier than once thought and is a possible factor in growth cessation (Müsel et al. 1997). Such a response might be controlled through release of peroxide into walls in much the same way as seen in walls subject to fungal attack. Peroxidase enzymes are candidates for the catalysis of these reactions. Understanding rigidification of this complex matrix of polymers demands input from the disciplines of biology, chemistry and physics. Combining established techniques with novel approaches to the study of individual cells (e.g. Fourier-Transform Infra-red microspectroscopy and the cell pressure probe) will bring new insights to the molecular basis of wall expansion.