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Journal of Experimental Botany, Vol. 54, No. 387, pp. 1585-1595,
June 1, 2003
© 2003 Oxford University Press
Growth and morphogenesis at the vegetative shoot apex of Anagallis arvensis L.
Received 29 October 2002; Accepted 5 March 2003
1 Institute of Plant Biology, Wroc
aw University, Kanonia 6/8, 50–328 Wrocaw, Poland
2 Department of Applied Mathematics and Theoretical Physics, Centre for Mathematical Sciences, University of Cambridge, Wilberforce Road, Cambridge CB3 0WA, UK
3 To whom correspondence should be addressed. Fax: +48 71 375 41 18. e-mail: dorotak{at}biol.uni.wroc.pl
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Abstract |
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A non-destructive replica method and a 3-D reconstruction algorithm are used to analyse the geometry and expansion of the shoot apex surface. Surface expansion in the central zone of the apex is slow and nearly isotropic while surface expansion in the peripheral zone is more intense and more anisotropic. Within the peripheral zone, the expansion rate, expansion anisotropy, and the direction of maximal expansion vary according to the age of adjacent leaf primordia. For each plastochron, this pattern of expansion is rotated around the apex by the Fibonacci angle. Early leaf primordium development is divided into four stages: bulging, lateral expansion, separation, and bending. These stages differ in their geometry and expansion pattern. At the bulging stage, the site of primordium initiation shows an intensified expansion that is nearly isotropic. The following stages develop sharp meridional gradients of expansion rates and anisotropy. The adaxial primordium boundary inferred from the surface curvature is shifting until the separation stage, when a crease develops between the primordium and the apex dome. The cells forming the crease, i.e. the future leaf axil, expand along the axil and contract across it. Thus they are arrested in this unique position.
Key words: Shoot apical meristem, strain anisotropy, strain rates, surface curvature, surface expansion.
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Introduction |
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Primary shoot structures originate from the shoot apical meristem, and all the cells of the shoot are derived from putative stem cells located at the top of the apex dome. Growth at the shoot apex is unique among plant meristems because it integrates two fundamental processes: the self-perpetuation of the apex dome and the initiation of lateral organs. Self-perpetuation requires the maintenance of meristem size despite the continuous flow of cells through the meristem. The initiation of lateral organs depends on the specification of groups of cells and their gradual separation from the meristem proper. The superposition of these two processes at the shoot apex leads to a complex, yet predictable, sequence of shapes. The complexity of meristem morphogenesis combined with the technical difficulties involved in observing the meristem explain why a detailed analysis of shoot apex morphogenesis has yet to be performed (Erickson, 1976; Silk, 1984).
Some significant features of shoot apex morphogenesis are nevertheless known. As a rule, all cells in the meristem are dividing (Clowes, 1959, 1961). However, the expansion and division rates of these cells vary over the apex volume. In the case of vegetative apices, growth is less intensive at the distal portion of the dome (the central zone) and more intensive at the periphery (the peripheral zone), especially where lateral organs are initiated (Romberger et al., 1993; Laufs et al., 1998a; Lyndon, 1998). The expansion of the shoot apical meristem is therefore inhomogeneous. Moreover, cells do not expand at the same rate in all directions. Instead, many cells show a main axis of extension while little extension is observed in the perpendicular direction (Hernández et al., 1991; Tiwari and Green, 1991). Meristem expansion is therefore anisotropic.
Little is known about the quantitative aspect of meristem morphogenesis beyond the general trends mentioned above. Perhaps the most fundamental reason why a detailed analysis of meristem morphogenesis is necessary is that a particular sequence of shapes can result from a wide range of growth patterns at the cell level (Hejnowicz and Nakielski, 1979; Nakielski, 1982, 1987). The only way to address the role of individual cells in morphogenesis is to quantify their contribution directly. Such quantitative data about meristem morphogenesis would be an important addition to the molecular knowledge of this process. They would help us to understand how known cytohistological zones (Clowes, 1961; Romberger et al., 1993) and gene expression domains (Bowman and Eshed, 2000; Brand et al., 2001; Traas and Doonan, 2001) are maintained, even if the cells that compose these zones and domains are continually changing. In particular, quantitative data on the rate of cell migration from one expression domain to the next would indicate how quickly cells need to acquire new ‘identities’.
With these applications in mind, what would constitute an adequate quantitative description of morphogenesis at the shoot apical meristem? Ideally, morphogenesis should be described in 3-D and with high spatial and temporal resolution. The two fundamental variables for any quantitative analysis of morphogenesis are the surface curvature and the surface strain rates. Curvature is a measure of the local geometry of the surface, while the strain rates measure its relative rate of expansion. If surface curvature and strain rates are to be compared with gene expression data, cellular resolution of these variables is needed since the transition between gene expression domains can occur over one cell length. Morphogenesis at the shoot apical meristem repeats itself at a regular time interval, the plastochron, corresponding to the time required for the initiation of successive leaves or groups of leaves at the apex (Erickson and Michelini, 1957). While, for the root meristem, a description of geometry at one instant is sufficient to apprehend the entire morphogenetic process (Silk et al., 1989), for the shoot apical meristem the description must span the whole plastochron. Therefore, protocols to investigate morphogenesis at the shoot apex must be able to resolve morphogenesis within a plastochron, which, in apices producing relatively large leaf primordia, usually lasts for a couple of days (Dale and Milthorpe, 1983; Lyndon, 1998).
This paper presents an analysis of surface expansion at the shoot apical meristem. The protocol uses a non-invasive replica method (Williams and Green, 1988) and a new computational approach that includes 3-D reconstruction of the meristem surface (Dumais and Kwiatkowska, 2002). This protocol allows the quantification of the geometry and expansion of the apex with relatively high temporal and spatial resolution. The data collected are used to address two fundamental questions concerning morphogenesis at the shoot apex: (i) what is the surface expansion pattern leading to the self-perpetuation of the apical dome; and (ii) what is the surface expansion pattern leading to partitioning of new leaf primordia from the meristem proper. The investigation is performed on vegetative shoots of Anagallis arvensis L. (Primulaceae) exhibiting spiral Fibonacci phyllotaxis.
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Plant material
Four shoot apices of A. arvensis were used in this investigation. With this material, meristem morphogenesis could be followed over seven plastochrons. Two apices (the first and the second apex) came from a plant collected on the Stanford University campus and kept in a growth chamber under a 16/8 h day/night cycle, a 15 W m–2 illumination, and a temperature of 20–21 °C (Dumais and Kwiatkowska, 2002). These conditions were non-inductive despite the fact that A. arvensis is a long day plant (Brulfert et al., 1985), presumably because the light spectrum did not include the inductive wavelengths. The other two apices (including the third apex presented here) originated from a clone derived by means of vegetative propagation from a wild plant growing in the Sudety Mts., Poland (the same clone as used by Kwiatkowska, 1997). These plants were kept at a temperature ranging from 18–24 °C, in short days (10/14 h day/night) to prevent flowering, with an illumination of 9 W m–2.
Data collection and analysis
A non-destructive replica method (Williams and Green, 1988; Green et al., 1991; Hernández et al., 1991) was used to obtain a developmental sequence for each apex. To expose the meristem surface, thin threads were glued to the tip of the young leaves overarching the meristem so that these leaves could be temporarily pulled away. The leaves were returned to their original position once the replica was completed. Replicas were made at 12–24 h time intervals and examined with scanning electron microscopy (Philips SEM505 and LEO435VP).
Computer tools developed earlier (Dumais and Kwiatkowska, 2002) were used to collect and analyse the morphogenetic data. The cell vertices, i.e. the points where three cells come into contact, were used as surface markers. For each member of a developmental sequence, two scanning electron micrographs were taken, one tilted by 10° with respect to the other. The x–y position of vertices was collected manually from one of the images, while for the other image the vertex locations were found with an area matching protocol. Since the two images offer slightly different perspectives of the same structure, the distances between vertices differed in these images. This shift in the x–y position of vertices was used to determine the elevation (the z position) of the vertices.
The curvature of the meristem surface was computed from the reconstructed meristem. A quadratic surface was first fitted to the vertices of a given cell and its contacting neighbours. The local curvature could then be computed from the coefficients of this quadratic surface. The computation of strain rates requires the identification of corresponding vertices on consecutive replicas of the growing meristem. This was done with a clonal analysis in which cells and their progeny were identified in consecutive images, like the exemplary cells highlighted in Fig. 1A–E. The strain rates at a given vertex were then computed from the deformation of the triangle defined by the three vertices in direct contact with that vertex. The overall strain rates for the cell were computed as the weighted sum of the strain rates at the cell’s vertices. A graphical user interface and computer programs written in Matlab (The Mathworks, Natick, MA, USA) can be downloaded at: http://culex.biol.uni.wroc.pl/instbot/dorotak.
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Results |
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Similarity of the developmental sequences confirms the validity of quantitative data
Although the manipulations of the meristem required to make replicas do not involve any wounding, they could nevertheless disrupt morphogenesis. Inspection of the material collected indicates that the geometry of the meristem has not been affected. For example, casts of apices at the same plastochron stage are strikingly similar. This is true whether replicas from the same apex, further referred to as members of the same developmental sequence (e.g. Fig. 1A, E), or from different apices (e.g. Figs 1A, 2C) are compared. The meristem manipulations could also have slowed down the rate of growth, which would lead to a longer plastochron. Such an effect on growth was indeed observed in some sequences excluded from this investigation. An increase in the length of the plastochron can also be seen in the sequence published in Fig. 5 of Green et al. (1991). A careful comparison of the observed plastochron is therefore in order. The estimated plastochrons for the sequences presented in this paper fall between 40 h and 45 h, irrespective of the origin of the plant material and the growth conditions. The material presented is therefore consistent at that level. Precise measurements of the plastochron were also published by other authors (Ballard and Grant Lipp, 1964; Brulfert et al., 1985), for decussate vegetative shoots of A. arvensis kept in the growth conditions similar to the plants used here. Again, this plastochron is consistent with the measured plastochrons for decussate shoots from which replicas were made (results not shown). The plastochron for shoots with decussate phyllotaxis is more than twice as long as shoots with spiral phyllotaxis, but not surprisingly since in decussate phyllotaxis two leaf primordia are initiated simultaneously. Therefore, half rather than the whole plastochron characteristic for decussate shoots should be compared with that of spiral ones (Rutishauser, 1998).
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Variables describing the geometry and expansion of the shoot apex surface
The geometrical analysis yields the principal curvatures and the Gaussian curvature for each cell on the reconstructed meristem surface. It should be emphasized that the curvature values reported do not refer to individual outer walls of the cells but rather to the curvature of the smooth surface defining the overall geometry of the meristem. The principal curvatures are the extreme curvature values, attained on a surface along principal curvature directions. In plots of the meristem curvature (the second column in Figs 1–3), the principal curvature directions for each cell are illustrated by the two arms of a cross. The length of these arms is proportional to the curvature in their direction. If in the arm direction the surface is convex, the cross arm appears in black; if the surface is concave, the arm is red. The Gaussian curvature is the product of the principal curvatures and shows how much a surface differs from being flat. If a surface is flat or can be flattened without any tears or folds, its Gaussian curvature is zero. A concave or convex surface that tears when flattened, for example, a hemisphere, has a positive Gaussian curvature. A surface shaped like a saddle, which will form folds, if flattened, has a negative Gaussian curvature. In curvature plots, the Gaussian curvature is indicated by the colour assigned to each cell (Figs 1–3). The warmer the colour, the higher is the curvature.
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The growth analysis yields the principal strain rates (
and 
) and directions, areal strain rate, and strain rate anisotropy. The strain rates are the relative rates of extension (Erickson, 1966, 1976; Hejnowicz and Romberger, 1984; Silk, 1984; Goodall and Green, 1986; Dumais and Kwiatkowska, 2002). If surface expansion is anisotropic, these rates assessed in different directions are not the same. There are two unique mutually orthogonal directions on the surface, in which strain rates attain extremal, minimal and maximal, values. These directions are the principal strain rate directions. The extremal values are called principal strain rates. The crosses plotted for each cell in the third column in Figs 1–3 indicate the principal strain rate directions. The length of cross arms is proportional to the corresponding principal strain rate. Positive strain rates (extension) are indicated with black cross arms, negative strain rates (contraction) are in red. The principal strain rate directions used in the present paper are the same as directions pointed by the ‘strain crosses’ introduced by Goodall and Green (1986), and theoretically defined principal directions of growth (Hejnowicz and Romberger, 1984). The areal strain rate is computed as the sum of the principal strain rates and is indicated in the strain rate plots by the cell colour. The warmer the colour, the higher is the areal strain rate. The strain rate anisotropy, given by the equation ( 
), shows how much the two principal strain rates differ from each other.
Geometry and growth of the apical dome are not uniform and change during the plastochron
The apical dome is always in contact with three leaf primordia of various ages. The Gaussian curvature of the central zone of the dome is positive but close to zero (the second column in Figs 1, 2). The curvature of the peripheral zone depends on the plastochron stage and the age of the contacting leaf primordium. At the beginning of the plastochron, a new leaf primordium becomes discernible within the peripheral zone as a region where the Gaussian curvature is locally higher and minimal and maximal curvatures are very similar (e.g. P1 in Fig. 2F). Leaf initiation sites become apparent before any signs of bulging can be seen on scanning electron micrographs of the apex surface (compare P1 in micrograph in Fig. 2C with the curvature plot in Fig. 2F). During the second half of the plastochron, a narrow band, one to three cells wide, becomes visible within the peripheral zone along the recently initiated primordium (e.g. along P3 in Fig. 1G). These cells are curved almost exclusively in the direction of the primordium margin. Thus a typical cross of principal curvatures is reduced to a latitudinal line segment. At the same time a part of the peripheral zone where the next primordium will be initiated attains a slightly higher curvature (e.g. the upper right portion of the dome in Fig. 2E). During the following plastochron the cells forming the band of zero Gaussian curvature give rise to the primordium axil (compare cells located along the P3 margin in Fig. 2D, E). In the present paper the term axil cells is referring to cells, which have the most negative meridional curvature of all the cells located near the primordium margin. In the curvature plots they have the longest red arms, such as the cells on the margin of primordium P3 in Fig. 2F. The parts of the peripheral zone contacting the axil (e.g. at the contact with P4 in Fig. 1F and G) are usually saddle-shaped: convex along the dome circumference (black cross arms), and slightly concave along its meridian (red cross arms). Between the regions contacting leaf primordia, the peripheral zone cells are curved predominantly in the meridional direction (between P2 and P3 in Fig. 1H).
The expansion of the apex dome surface is non-uniform and generally anisotropic (the third column in Fig. 1 and 2). In terms of surface expansion, the central zone and the peripheral zone merge into each other gradually. Areal strain rates are the lowest and strain is nearly isotropic in the central zone, while in the peripheral zone both strain anisotropy and areal strain rates are generally higher. However, expansion of the peripheral zone is not uniform. It is slow and not strongly anisotropic in the region adjacent to the recently formed primordium (e.g. the cells adjacent to P2 in Fig. 1L–M), where maximum strain rates are latitudinal, i.e. in the direction parallel to the primordium margin. The above-mentioned band of cells having nearly zero Gaussian curvature gives rise to the leaf axil expanding along the primordium margin (latitudinally) and contracting across it (meridionally, like the cells along P3 in Fig. 1L). The expansion of the formed axil cells is similar (like cells of the axil of P3 in Fig. 1M or P3 in Fig. 2H). The area of these cells is almost not changing, or slightly diminishing. Where the dome flanks are rebuilding at the contact with the primordium axil (for example, adjacent to the axil of P3 in Fig. 1M), areal strain rates and anisotropy are high. There the maximal strain rate is in the meridional direction. This direction is perpendicular to the maximal strain rate direction of the adjacent axil cells. The region contacting the axil is the fastest growing portion of the peripheral zone during the first half of the plastochron (Fig. 1M). In the second half of the plastochron, an additional region of increased areal strain rate appears at the initiation site of the next primordium. This region covers a portion of the peripheral zone larger than is actually contributing to the formation of a new primordium (e.g. around P1 in Fig. 1L). When the next plastochron begins, the cycle is repeated but rotated by the Fibonacci angle around the apex, e.g. by about 137.5° clockwise between members of the first sequence shown in Fig. 1K and M.
Emerging leaf primordia show increasing gradients of curvature and strain rate
Four developmental stages can be distinguished in the course of the primordium formation covered in this study. These are: (i) the bulging stage; (ii) the lateral expansion stage; (iii) the separation stage; and (iv) the bending stage. The first two stages cover about one plastochron. The third one lasts nearly half of a plastochron; whereas the fourth one extends beyond the period covered in this study. Primordium P2 from the first sequence (Fig. 1) and primordium P1 from the third one (Fig. 3) can illustrate all the stages. In the first stage, the leaf buttress is characterized by locally elevated Gaussian curvature (P2 in Fig. 1F–H). The buttress surface is expanding with high areal strain rates and nearly isotropically (P2 in Fig. 1K, L). The region of increased expansion sometimes covers more than the surface of the emerging primordium (e.g. P2 in Fig. 1L). During the second stage, the buttress attains an oval shape (P2 in Fig. 1I; P1 in Fig. 3E). Cells on the primordium surface expand mostly latitudinally, while some contraction may take place in the meridional direction (P2 in Fig. 1M). The cell area, however, is still increasing since the latitudinal expansion exceeds the meridional contraction. During the third stage (P2 in Fig. 1J; P1 in Fig. 3F), the buttress becomes separated from the dome by the axil. At the same time, the primordium surface expands similarly to the preceding stage. Finally in the fourth stage (P1 in Fig. 3G, H), a flattened primordium is formed with an adaxial surface of negative Gaussian curvature and an upper portion of high positive curvature. The adaxial surface is concave meridionally and convex latitudinally. At this stage the primordium starts to overarch the apex (like P1 in Fig. 3G, H). The fourth stage is characterized by differential growth. The expansion of the upper leaf portion where the Gaussian curvature becomes relatively high is intensive and not strongly anisotropic (for example, the outermost cells of P1 in Fig. 3J, K). The adaxial surface of the primordium bends towards the dome. Its cells expand relatively slowly and with high strain rate anisotropy (see remaining cells of the above-mentioned primordium). Some meridional contraction is observed on the adaxial side of the primordium, while the maximal strain rate direction is parallel to the leaf axil.
Leaf primordium boundaries inferred from the surface curvature are unstable with respect to cells during early leaf development, i.e. during the bulging and lateral expansion stages. The boundaries are shifting during these two stages by one or two rows of cells. Both an incorporation of new cells into the primordium (e.g. P1 in Fig. 4A) and, less often, a reduction of the primordium area can take place (P2 in Fig. 4A). Starting from the third developmental stage, the displacements of the primordium boundaries are significantly reduced. This corresponds to the time when the leaf axil is formed between the apical dome and the leaf primordium. Subsequently, shifting of the primordium boundaries is restricted to the lateral sides of the primordium (right side of P1 in Fig. 4C), where the Gaussian curvature is negative but closer to zero than in the middle of the axil. Sometimes, starting from the separation stage, the boundaries do not move at all (P3 in Fig. 4B).
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Discussion |
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A non-invasive replica method and a 3-D reconstruction algorithm were used to quantify the geometry and surface expansion of the shoot apex of A. arvensis. The results were analysed in the light of two fundamental processes occurring at the shoot apex: the self-perpetuation of the apical dome and the formation of leaf primordia. The observed changes in the geometry and expansion of a leaf primordium surface were characteristic enough to differentiate early leaf development into distinct stages. These stages could be regarded as representative for the early development of dicot leaves, of which the leaf of A. arvensis is a typical example.
Self-perpetuation of the vegetative shoot apical dome
The overall pattern of expansion in the shoot apex is characterized by a gradual increase in the rate of expansion and expansion anisotropy as one goes from the central zone to the peripheral zone. These observations are in agreement with earlier studies of shoot apex growth (Newman, 1956; Soma and Ball, 1963; Hussey, 1971; Green et al., 1991; Lyndon, 1998; Laufs et al., 1998a). This particular pattern of expansion, however, is not unique in its ability to maintain the size and dome-like shape of the shoot apical meristem (Hejnowicz and Nakielski, 1979; Nakielski, 1987). Opposite gradients of expansion were observed on the dome of tip-growing cells (Hejnowicz et al., 1977; Shaw et al., 2000) and there is no geometrical reason why a similar expansion pattern would not work in the shoot apical meristem. A possible biological explanation for the slow growth of the central zone is that it helps the plant preserve the genetic material of reproductive cells which will originate from this region (Hejnowicz and Nakielski, 1979; Klekowski, 1988). If the rate of expansion is low, the cells will divide less frequently thus reducing the likelihood that replication errors will occur. An alternative explanation is that the high rate of growth in the peripheral zone results from the rapid initiation of lateral organs. This explanation is incomplete since in the pin1 mutant of A. thaliana the inflorescence apex does not initiate lateral organs and yet the mitotic index remains relatively high in the peripheral zone (Vernoux et al., 2000).
The peripheral zone is differentiated into regions whose distribution is related to the youngest three leaf primordia. Similar sectors, comprising the peripheral zone of the shoot apex of Clethra barbinervis Sieb. et Zucc. exhibiting spiral phyllotaxis, were reported by Hara (1971) who determined them based on the anticlinal cell walls pattern. In the apex of A. arvensis, the geometry and growth of these sectors were shown to depend on the age of the adjacent leaf primordium. The sectors differ in their Gaussian curvature and in their expansion rates and anisotropy. Even the direction of maximal strain rate in adjacent sectors may be entirely different. The site of leaf primordium initiation is characterized by locally intensified growth. Interestingly, the region of high strain rate also includes cells that will not contribute to the new primordium. For each plastochron, the growth pattern is rotated around the apex by a fixed angle (the Fibonacci angle). The temporal and spatial complexity of the dome expansion is a consequence of the fact that the continuous initiation of leaf primordia in a regular but asymmetric pattern is superimposed on the self-perpetuation of the dome.
Formation of the leaf axil—partitioning of the shoot apex
The leaf axil is unique among the different regions of the meristem, both in its geometry and growth. The meridional gradients of curvature, areal strain rate, and strain rate anisotropy are especially sharp across this region. The expansion observed in the leaf axil is similar to the expansion observed during the formation of the radial creases between stamens in A. arvensis flowers (Hernández et al., 1991), and also the creases delimiting organs in adventitious shoots of Graptopetalum paraguayense E. Walther (Tiwari and Green, 1991). The expansion of the future axil cells is very slow, and takes place only along the axil as observed also by Green et al. (1991). Thus, the cells forming the axil are arrested in this particular position. This observation may explain why Soma and Ball (1963), as well as Hussey (1971), noted that some of the markers applied to the apex surface were not moving out of the young leaf axils. Another consequence of such a pattern of cell expansion is that its appearance on the adaxial margins of a leaf primordium significantly reduces the displacement of the primordium boundary, i.e. it partitions the shoot apex surface into the dome and new leaf primordium.
Early development of the leaf primordium
In this study, the course of leaf primordium formation is divided into four stages. These stages cover the first (‘organogenesis’) stage and a part of the second (‘early growth of the primordium’) stage of leaf development defined by Sylvester et al. (1996). Distinct changes in primordium geometry and growth allow the first, ‘organogenesis’ stage to be resolved. It comprises the bulging, the lateral expansion, and the separation stages of the present paper, while the bending stage is at the beginning of the stage of ‘early growth’. During the bulging stage, growth of the primordium is relatively intensive and isotropic. The cytoplasm of cells comprising the primordium at this stage, visible in longitudinal sections of the apex, stains more intensely than the cytoplasm of other meristem cells (Fontaine, 1972). The intense staining is another manifestation of the high meristematic activity of these cells. By contrast with the bulging stage, lateral expansion of the primordium surface relies on relatively low areal strain rates, but high strain rate anisotropy. Later, during primordium development, sharp meridional gradients of strain rates and anisotropy appear, and the primordium surface becomes differentiated into regions of various expansion and geometry.
The control of areal strain rates and strain rate anisotropy
One important conclusion from this work is that morphogenesis at the shoot apex depends on sharp gradients of areal strain rates as well as on anisotropy of strain rates. One should therefore ask how these gradients and anisotropy are created and maintained at the cell and molecular levels.
Areal strain rate is scalar, that is, any given point of the meristem surface has a single areal strain rate value. Its level could be directly regulated by a scalar factor, such as the concentration of gene products or growth regulators (Hejnowicz and Romberger, 1984). In the apical dome, the distribution of such factors can be regulated by genes whose expression is specific to the peripheral or central zone (Nishimura et al., 1999; Zondlo and Irish, 1999). Some of these genes, such as CLAVATA and MGOUN, are known to regulate the size of these meristem zones (Laufs et al., 1998a, b; Traas and Doonan, 2001). The primordium initiation site, in turn, shows an elevated expression of the expansin gene (Reinhardt et al., 1998). Also, exogenous application of expansin to the apex surface can induce leaf-like outgrowths (Fleming et al., 1999), while local induction of expansin expression can induce the entire process of leaf formation (Pien et al., 2001). The intensified areal strain rate observed at leaf initiation may result from a local loosening of cell walls on the apex surface by secreted expansin proteins (Fleming et al., 1999).
Future axil cells presumably also express specific genes, although the exact spatial and temporal relationships between gene expression domains and future axil cells remain unknown. The expression domains of CUP-SHAPED COTYLEDON in Arabidopsis thaliana (L.) Heynh. (Aida et al., 1999; Brand et al., 2001; Traas and Doonan, 2001), NO APICAL MERISTEM in Petunia sp. hybrids (Souer et al., 1996), and various OSH genes in Oryza sativa L. (Sentoku et al., 1999) are correlated with primordium boundaries. The CUP-SHAPED COTYLEDON2 gene is thought to act as a local growth suppressor (Traas and Doonan, 2001). Theoretically, a crease can be formed by locally lowering the areal strain (Todd, 1986). Therefore, some of the above-mentioned genes may be directly involved in the control of axil formation by locally inhibiting cell expansion.
Unlike areal strain rates, the strain rate anisotropy must be regulated by factors of a similar, anisotropic nature (Hejnowicz and Romberger, 1984). The significance of anisotropic factors in morphogenesis is supported by the fact that dividing plant cells seem to ‘perceive’ stress or growth anisotropy existing in their environment (Miller, 1980; Hejnowicz, 1984; Lintilhac and Vesecky, 1984; Lynch and Lintilhac, 1997). Neither the concentration of gene products (scalars), nor the concentration gradients of such gene products (vectors) are anisotropic factors. The latter have only a single direction defined together with a corresponding scalar value. By contrast, cellular structures such as the cytoskeleton or the cell wall often show anisotropy. A connection between cytoskeletal arrangement, cell wall reinforcement, and anisotropic cell expansion has been shown (see for example Selker and Green, 1984; Tomos et al., 1989; Green, 1992; Bichet et al., 2001). In the case of cell walls, a non-random alignment of cellulose microfibrils results in differences in mechanical properties of walls along different directions. Cell walls are mechanically reinforced in the direction of cellulose alignment. Since the outer walls of tunica cells in the shoot apical meristem are the thickest of all the meristem cell walls, the reinforcement of these particular walls is a good candidate for the direct control of the apex surface growth (Green, 1986; Green and Selker, 1991). Combining the study of cell wall mechanics with quantitative data on the morphogenesis at the shoot apical meristem may help to understand how meristem growth is controlled.
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Acknowledgements |
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The authors thank Dr Zygmunt Hejnowicz (Silesian University, Poland) for critical reading of the manuscript and Drs Zofia Czarna and Krystyna Heller (Electron Microscopy Laboratory, Wroc
aw University of Agricultural Sciences, Poland) for help in preparing some of the scanning electron micrographs used in this work. Part of this research was financed by a research grant No. 3P04C 015 22 from the Polish Committee for Scientific Research to DK and a NSF grant to the late Paul Green. DK acknowledges additional support from the Fulbright Foundation while visiting Stanford University. JD acknowledges support from the Center for Computational Genetics and Biological Modeling (Stanford University).
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