- Research article
- Open Access
The development and geometry of shape change in Arabidopsis thalianacotyledon pavement cells
© Zhang et al; licensee BioMed Central Ltd. 2011
Received: 11 October 2010
Accepted: 1 February 2011
Published: 1 February 2011
The leaf epidermis is an important architectural control element that influences the growth properties of underlying tissues and the overall form of the organ. In dicots, interdigitated pavement cells are the building blocks of the tissue, and their morphogenesis includes the assembly of specialized cell walls that surround the apical, basal, and lateral (anticlinal) cell surfaces. The microtubule and actin cytoskeletons are highly polarized along the cortex of the anticlinal wall; however, the relationships between these arrays and cell morphogenesis are unclear.
We developed new quantitative tools to compare population-level growth statistics with time-lapse imaging of cotyledon pavement cells in an intact tissue. The analysis revealed alternating waves of lobe initiation and a phase of lateral isotropic expansion that persisted for days. During lateral isotropic diffuse growth, microtubule organization varied greatly between cell surfaces. Parallel microtubule bundles were distributed unevenly along the anticlinal surface, with subsets marking stable cortical domains at cell indentations and others clearly populating the cortex within convex cell protrusions.
Pavement cell morphogenesis is discontinuous, and includes punctuated phases of lobe initiation and lateral isotropic expansion. In the epidermis, lateral isotropic growth is independent of pavement cell size and shape. Cortical microtubules along the upper cell surface and stable cortical patches of anticlinal microtubules may coordinate the growth behaviors of orthogonal cell walls. This work illustrates the importance of directly linking protein localization data to the growth behavior of leaf epidermal cells.
The elaboration of blade shaped organs is a common morphological process in the plant kingdom. It is also quite plastic. Developmental gradients and environmental inputs can generate highly variable leaf shapes over the lifespan of the plant [1, 2]. An important challenge is to understand the complex interplay of cell number and the geometry of cell growth at regional scales that can dictate the spatial patterns of organ formation . In the leaf, the epidermis is an important architectural control element. Genetic mosaics indicate that the genotype of the epidermis has a major impact on the growth properties of underlying tissues and the overall form of the organ [4–6]. Therefore, the morphogenesis of epidermal pavement cells is of particular interest. As in other tissues, both cell division and irreversible cell expansion in the epidermis contribute to tissue morphology. However, cell size increase is the dominant factor during organ expansion. For example, epidermal pavement cells in the dicot Arabidopsis thaliana undergo multiple rounds of endoreduplication , and simultaneously increase in cell volume by almost 2 orders of magnitude compared to their protodermal precursors [8–11]. As pavement cells increase in size they remain highly vacuolated, and the thickness of the cell wall does not increase significantly [8, 10]. Therefore pavement cell size increase is true cell growth that includes the balanced synthesis of new vacuole, plasma membrane, and cell walls. Unlike animal cells , the shape changes of plant cells during cell growth are defined by the mechanical properties of the cell wall [13, 14]. In the epidermis, the thick external cell wall impedes expansion perpendicular to the leaf surface ; consequently cell size increase occurs preferentially within the plane of the epidermis.
Pavement cell expansion in the lateral dimension often occurs in a sinusoidal pattern, generating highly interdigitated cells . The striking undulation of the cell wall is widespread in the plant kingdom and is not limited to epidermal cell types. For example, in the fern Adiantum capillus-veneris, leaf mesophyll cells that are in physical contact with one another initiate lobes that are in direct opposition . Polarized expansion of the opposing lobes generates air spaces between cells that facilitate efficient gas exchange between the plant and the environment. In the epidermis adjacent pavement cells initiate protrusions that are offset from one another. The subsequent pattern of cell expansion generates an interdigitated, mechanically stabilized tissue.
There is a correlation between the occurrence of localized anticlinal (perpendicular to the leaf surface) microtubule bundles (AMBs) and the presence of cell indentations that form a local concave shape [18–21]. In concave regions of the growing pavement cells there also is a correlation between the location of AMBs and the presence of dense pads of cellulose microfibrils at the interface of the anticlinal and outer periclinal (parallel to the leaf surface) cell walls . This activity is significant because cellulose microfibrils are the primary load-bearing polymer in the plant cell wall and their pattern of deposition at the plasma membrane is dictated by cortical microtubules [22–24]. However, the morphogenesis of lobed cells is complicated and includes many cellular activities in addition to those that directly affect cellulose deposition. For example, mutations that affect the actin cytokeleton, targeted vesicle secretion, and non-cellulosic components of the extracellular matrix cause pavement cell growth defects [rev. in: [16, 25]].
Despite genetic and ultrastructural descriptions of pavement cell growth there is still very little clear knowledge about the geometry and cellular dynamics of pavement cell shape change. Current models of the growth process are varied, and are derived from static images collected from populations of cells. Some models propose that pavement cell growth includes sequential phases of cell expansion along the proximo-distal and lateral leaf axes , with selective expansion in lobes driving cell expansion primarily in the lateral dimension . Other models propose a continuous and iterative lobe initiation process during cell morphogenesis [20, 27]. The role of AMBs in the epidermal tissue is also unclear. These specialized microtubule zones are presumed to direct the synthesis of oriented cellulose microfibrils. Based on ROP small GTPase and AMB localization in cells that had a lobed morphology, it was hypothesized that localized synthesis of parallel arrays of cellulose microtubules in the anticlinal wall locally restricts protrusive growth perpendicular to the cellulose microfibril network, initiates lobe formation, and promotes polarized lobe expansion [rev. in: [16, 26, 27]]. The analogy to the restriction of radial expansion of cylindrical cells is valid for pavement cells only if parallel arrays of microfibrils in the anticlinal wall extend into the periclinal wall. In addition, the restricted growth model cannot explain persistent interdigitating growth during which the protrusive (convex geometry) growth of one cell must be accommodated by the complimentary growth of the concave indentation of the neighboring cell. The model above also does not account for the detection of AMBs within the lobes of cotyledon pavement cells , which is presumed to be a subcellular domain of accelerated growth [26, 27].
In this paper we take advantage of the developmental synchrony and simplicity of cotyledon development to monitor the microtubule organization and cell shape changes that occur during pavement cell morphogenesis. Time series images of cotyledon pavement cells and the use of fiduciary extracellular marks reveal distinct phases of lobe initiation and subsequent uniform cell expansion in the plane of the epidermis. Our microtubule localization experiments during the lateral isotropic growth phase confirm previous reports of clustered anticlinal microtubules along cell indentations [16, 26, 28] and within lobes . In this paper we demonstrate that asymmetric patterns of cortical microtubules persist for days, but are not necessarily associated with polarized growth.
Size and geometry of pavement cells at different stages of cotyledon development
Number of Skeleton Ends
Growth Rate (%/hour)
2 (N = 41)
2169 ± 597 (1)
279 ± 66 (2)
0.35 ± 0.08 (3)
8 ± 2 (4)
5 (N = 44)
3756 ± 1973
401 ± 175
0.30 ± 0.09
11 ± 4
1.02 ± 0.53(5)
12 (N = 43)
16160 ± 4434
1181 ± 278
0.15 ± 0.05
18 ± 4
1.97 ± 0.54(6)
18 (N = 35)
15399 ± 4476
1070 ± 253
0.17 ± 0.04
15 ± 4
Lobe initiations and splits at different time points during cotyledon pavement cell development
Cells with lobe
% of cells with
28 (N = 4)(1)
17 (N = 3)
22 (N = 5)
Linear regression analysis of cell area, perimeter and single segment changes from 3 DAG to 5 DAG using time-lapse images
IF (%) (2)
Growth rate (%/hour) (3)
1 (N = 6)
0.963 ± 0.030 (4)
91 ± 3 (5)
1.89 ± 0.26 (6)
2 (N = 4)
0.991 ± 0.007
89 ± 4
1.11 ± 0.18
3 (N = 5)
88 ± 2
1.42 ± 0.24
In many lobed cell types, parallel arrays of AMBs are distributed unevenly along the cell perimeter and are thought to have a strong influence on the morphogenesis process [19, 20, 26, 27]. In both 3 and 5 DAG cells, many but not all cell indentations corresponded to sites where periclinal microtubules coalesced with clearly resolved AMBs (Figure 2A,F). A region from a confocal image of two such indentations was digitally resliced to examine the AMBs in xz and yz views (Figure 2B,D,G, and 2I). The AMBs had a clear parallel alignment, and intensity profiles across the region demonstrated our ability to resolve distinct microtubule structures (Figure 2C,E,H and 2J). Although the lifetime of individual bundles was not measured, specific domains of the cortex of individual cells were populated by AMBs over a 2 day period. For example, cortical domains inside the anticlinal wall that were populated by AMBs at 3 DAG were also enriched in AMBs at the 5 DAG time point. At 5 DAG, the AMBs had increased in number and occupied a more extended domain of the cortex (Figure 2C,E,H, and 2J). Although zones populated by anticlinal bundles persisted for days, the closely associated microtubule network on the periclinal cell surface was obviously reorganized during the same growth interval. For example, in the inset, red-boxed region of cell 4, many microtubules coalesced at or emanated from an indentation (Figure 2A, inset), but at 5 DAG the periclinal microtubules in the same region had no clear pattern (Figure 2F, inset).
Visual comparisons of individual pavement cells at 3 and 5 DAG made it seem impossible that uniform cell growth restricted to the cell periphery could explain the observed shape transitions from 3 to 5 DAG cells. We tested an alternative growth model of uniform lateral isotropic expansion of periclinal cell wall surfaces by digitally magnifying the thresholded image of a 3 DAG cell (Figure 4G,K and 4O) by a constant so that its final area (Figure 4H,L and 4P) was equal to the measured area for that same cell at 5 DAG (Figure 4I,M and 4Q). The digitally magnified cell was rotated to maximize the overlap of the magnified image with the real 5 DAG cell. An overlay of the 2 images (Figure 4J,N and 4R) was used to measure the ratio of overlapping pixels (Figure 4J,N and 4R, yellow) to the total number of pixels for the real 5 DAG cell (Figure 4J,N and 4R, green). This ratio, which can be interpreted as an "isotropy factor", would be equal to 1 if the overlap was perfect. In three independent fields of pavement cells, the mean isotropy factor ranged from 0.88 to 0.91 (Table 3).
The extent of isotropic lateral growth was independent of cell size, because small (cell 1, Figure 4G-J), medium (cell 6, Figure 4K-N), and large (cell 4, Figure 4O-R) cells at the 3 DAG time point had very similar isotropy factors. An isotropy factor value less than 1 could be caused by human error during the digital cell dissection protocol. To characterize this error, 6 cell images were repetitively dissected, digitally magnified, and the overlap between all possible cell pairs was calculated. For the repeat dissections, the measured overlap value of 0.97 ± .01 (mean ± SD, n = 6) was close to the expected complete overlap. The ~3% error in dissection accuracy cannot explain the isotropy factor values calculated for growing cells (Table 3). Using time-lapse images, we also analyzed the circularity values for cells at 3 and 5 DAG. The mean circularity values of 3 (0.26 ± 0.08, mean ± SD, N = 15) and 5 (0.24 ± 0.08, mean ± SD, N = 15) DAG cells were clearly higher than those of fully expanded cells (Table 1). These findings suggest that an additional phase of polarized cell growth occurs at later stages of cotyledon development. Pair-wise comparisons of the circularity values of 3 and 5 DAG cells did not detect significant differences. However, there was a clear trend toward lower values in 5 DAG cells; because 80% of the 5 DAG cells had a circularity value that was lower than the corresponding 3 DAG cell. Based on the significant increase in cell shape complexity and the number of skeleton ends between 5 and 12 DAG (Table 1), additional lobe initiation events are likely to be common at later times of cotyledon development.
The size and shape of aerial organs in plants can be understood as an emergent property that arises from complex interactions between tissues [4, 5] and regional differences in the growth behavior of sectors of cells . The epidermis features prominently in growth control models, and yet there is a lack of basic knowledge about the morphogenesis of pavement cells, which are the fundamental building blocks of the tissue. This paper provides important new methods to analyze the morphogenesis and cell biology of the epidermal tissue and its constituent pavement cells. These data provide specific geometric rules that govern a persistent maintenance phase of pavement cell growth that contributes significantly to the size increase of the cotyledon.
Our time course observations of developing pavement cells reveal an initial wave of lobe initiation followed by an extended phase of isotropic cell expansion. This differs from previous models of pavement cell shape change that were based on static images and population-level sampling [10, 20, 26]. The population-level measurements here are also misleading, and depict lobe initiation and growth as a continuous process (Table 1). This is clearly not the case. Lobe formation in cotyledons, like cell division rates, metabolism, and stomatal development [35–37], undergoes a sharp transition at or near the 2 DAG stage (Table 2). Sequential images of developing pavement cells clearly revealed an early phase of growth and lobe initiation that was completed at or near 3 DAG, and a subsequent period of diffuse growth from 3 to 7 DAG during which lobe formation was rare. Sequential patterning and maintenance phases of growth are also observed in trichomes, a highly branched unicellular epidermal cell type [38, 39]. In future experiments we will try to learn more about the symmetry break that occurs during lobe initiation and the extent to which the similar genetic control of pavement cells and trichome shape  reflects a common usage of patterning and growth control machineries.
Because of its importance during organ expansion, we focused our analyses on the growth phase that occurs in the absence of frequent lobe initiation. As expected for cells that use a diffuse growth mechanism, the amount of cell growth in the 3 to 5 DAG interval was related to the initial cell area, because the magnitude of surface area increase is positively correlated with cell size. In three independent fields of cells, when cell size at 5 DAG is plotted as a function of initial cell surface area, the data points define a straight line, with extremely high R2 values (Table 3). Therefore, within the sampled fields of cells, growth is uniform and independent of cell boundaries. This coordinated growth behavior would minimize shearing forces between cells that are physically coupled by the cell wall, and is expected if groups of cells employ a uniform diffuse growth mechanism and all expanding surfaces experience an equal strain.
Detection of equal growth rates among fields of cells does not address the geometric path of the cell shape change. To learn about the spatial dynamics of growing pavement cells we used three-way cell wall junctions as fiduciary marks to monitor the spatial behavior of the cell anticlinal wall, which unambiguously defines the leading lateral edge of the growing cell. In 3 independent populations of cells (Figure 4F), increases in anticlinal wall length were remarkably uniform along the cell perimeter (Figure 4F, Table 3, Figure S1). This would be expected for uniform diffuse growth of the ribbon of anticlinal wall within the plane of the leaf. Height increases in the anticlinal wall are unpredictable (Figure 5) and the behavior of this cell surface requires further study.
In terms of lateral cell growth, the low spatial resolution of our fiduciary marks cannot detect micro-heterogeneity in growth at micron or nanometer scales. However, the perimeter segments did resolve lobes and indentations within individual pavement cells. Previous localization data on lobed epidermal cells led to the idea that lobed regions expand at a greater rate compared to indentations and more central domains of the cell [16, 26]. To the contrary, our findings indicate that the entire anticlinal cell wall grows at similar rates that are independent of cell shape (Figure 4F). In fact, the entire lateral surface of the cell expands more or less isotropically (Figure 4G-R). Our analysis of cell growth behavior in three independent fields of cells is consistent with this idea. Regardless of their size or shape, the mean isotropy factors ranged from 0.88 to 0.91 (Table 3) and circularity measurements of the pavement cells at the two time points were very similar. Therefore, during this maintenance phase of pavement cell morphogenesis, fields of expanding cells follow a previously defined pattern and accommodate the growth of their neighbors: indentations, protrusions, and midzones of adjacent cells expand in harmony. This contrasts with intrusive growth behavior of fusiform cambial initials, in which the growth of one cell occurs at the expense of its neighbor . The detection of equal cell expansion rates within sectors of the leaf that span ~ 6 cell diameters (Figure 4A-C) suggests that the growth control occurs at a regional scale in the tissue.
The regional growth behavior of sectors within an organ contributes to macroscopic asymmetry . In our case the Arabidopsis cotyledon is very symmetrical, and this geometry may be the emergent property of isotropic lateral expansion in populations of pavement cells. However, we do not want to gloss over the fact that the lateral expansion during the 3 to 5 DAG interval is not completely uniform. Real cells consistently displayed local deviations from isotropic lateral expansion (Figure 4G-R) that could not be explained by measurement error. These local deviations may simply reflect random variability in the geometric path of lateral diffuse growth. Alternatively, it may reflect a distinct mechanism for local asymmetric growth. Regardless of the mechanism, local asymmetry in cell growth patterns can contribute to different tissue and organ geometries. In the future it will be important to develop cell wall marking techniques  that will allow us to monitor the surface behavior of pavement cells at a high resolution.
The cytoplasmic control of cell lobing is complex . Genetic and cytological data point to the involvement of microtubules and AMBs during local cellulose synthesis and cell shape control [19, 20, 22, 23, 26–28, 43, 44]. Furthermore, the ability of AMBs to localize the cellulose biosynthetic machinery has been shown , although this was in the context of localized wall synthesis in developing xylem cells that are no longer expanding. Although the involvement of AMBs in pavement cell shape control and wall extensibility has not been proven, it is reasonable to consider a mechanism that includes microtubule-dependent templating of cellulose microfibril synthesis. This cellular control mechanism is easiest to understand in the context of uniform diffuse growth along the periclinal surface of the pavement cells. The periclinal cell wall is thick and contains cellulose microfibrils of mixed orientations  and correlates with the variable configurations of periclinal cortical microtubules that have been reported [20, 27, 33, 46]. In fields of cells undergoing isotropic lateral expansion, we detect periclinal cortical microtubule networks whose alignments vary greatly between and within cells (Figure 2). Given the nearly isotropic growth of these cells (Table 3), the organization of the periclinal microtubule network at a particular moment [27, 46–48] has little predictive value with respect to the growth trajectory of the cell. Instead, this variability reveals a cell autonomous control of the microtubule array that could include modulation of the KATANIN-dependent severing of intersecting microtubules  and the dynamic remodeling of the inner-most network of cellulose microfibrils that determine the elastic properties of the wall.
The relationships between AMBs and pavement cell expansion are less obvious, and may vary depending on the cell type and/or the particular stage of pavement cell morphogenesis. In some cell types, lobe formation is associated with cell wall detachment and the localized expansion of protrusions that create air spaces within the internal tissues of the leaf . This cellular organization and shape change can be explained by a model in which the parallel alignment of microtubules and microfibrils locally restricts lateral expansion perpendicular to the cellulose microfibrils [reviewed in:]. Over time, uneven growth along the cell perimeter could generate a narrow indentation as cell expansion preferentially occurs in the developing lobes. In lobed epidermal cells from a variety of species, clustered anticlinal microtubules coincide with active sites of cell wall formation , and a modified version of this local microtubule growth restriction model has been adopted to explain lobe formation and polarized outgrowth in Arabidopsis leaf pavement cells [26, 27]. Although it is not known if cotyledon and leaf pavement cells adhere to same morphogenetic rules, it is possible that AMBs are patterning elements that define the positions of lobe initiation [Table 2, ]. However, during lobe initiation turgor pressures between two cells cancel along the anticlinal wall in regions of cell-cell contact. Therefore, modification of the local strain behavior of the cell wall alone is unlikely to be sufficient for lobe initiation.
The concept of persistent differential growth at the interface of a lobe and an indentation is also problematic because normally there are no gaps and very little overlap between pavement cells. Instead, the complementary cell expansion within the lobe of one cell and the indentation of its neighbor is required to preserve the integrity of the tissue . Consistent with this model of cell mechanics, we find that cotyledon pavement cells within a field display nearly equal length increases along the entire anticlinal wall, and the growth is independent of the local contour of the cell (Figure 4F). In the lateral dimension, the anticlinal wall responds uniformly to wall tension that is likely generated by the periclinal cell wall. A mechanical coupling of the anticlinal wall with an expanding periclinal wall could generate this tension.
Regardless of the mechanism, it is clear from our analysis of the lateral isotropic growth phase that anticlinal wall strain in the plane of the leaf is quite uniform and also includes a growth vector that is perpendicular to the anticlinal wall. At first glance this seems to be at odds with the patchy distribution of AMBs (Figure 2) and their presumed involvement in the synthesis of parallel arrays of cellulose microfibrils that would resist radial expansion of the cell perpendicular to the microfibril network. However, this growth control model assumes that cellulose microfibrils in the anticlinal wall are physically coupled to aligned microfibrils in the periclinal cell wall that resist radial expansion. In contrast to typical cylindrical cells that have a net transverse orientation of cortical microtubules (and microfibrils) at a whole cell scale, pavement cells only occasionally display aligned microtubules that span the anticlinal and periclinal walls (Figure 2A,F, insets). It may be that the physical coupling of the periclinal and anticlinal wall is regulated during growth, and that forward progression of the anticlinal boundary may not always be restricted by linkages with the periclinal wall. We speculate that phenomena such as regulated microtubule-dependent nucleation [49, 50] at the junctions of anticlinal and periclinal walls could, via the local activity of CESA, modulate the resulting physical connectivity of cellulose microfibrils between these two cell surfaces.
Time-lapse live cell imaging and new quantitative analyses of the growing epidermis allowed us to study the dynamic process of pavement cell morphogenesis and its relationship to the microtubule cytoskeleton. During pavement cell development, there are distinct phases of lobe initiation punctuated by lateral growth that is highly isotropic. During lateral isotropic growth cortical AMBs are found both along cell indentations and within lobes. In some cases cortical domains of AMBs spread and persist for days. Although it is clear that AMBs do not restrict cell expansion, their importance during the symmetry breaking events of lobe initiation and the coordination of isotropic growth within and between cells is unknown. Further integration of live cell imaging, computational tools, and genetics can provide a way to dissect morphogenesis at spatial scales that span from the initiating pavement cell lobe to the macroscopic features of an expanding leaf.
Seedling growth conditions and cell staining
Arabidopsis thaliana (Col-0) seedlings were grown in 0.5 × MS (Casson, North Logan, UT) media in a Percival chamber at 22°C under continuous illumination (90 μmol m-2 sec-1). Seedlings that germinated at 36 h after transfer from cold treatment to the growth chamber were used in subsequent analyses. To obtain static images of cells from synchronized populations, whole seedlings were stained with 1 μM FM4-64 as described previously . For time lapse imaging ~1 cm2 of agar was cut around each 3 DAG plant and the TUB6:GFP-expressing seedlings were aligned on a petroleum jelly chambered slide and mounted in water. After one round of imaging the seedlings were transferred to humidified chambers and remounted 2 days later in the same manner.
Immunolocalization and particle bombardment
Seedlings were staged as described above and processed for immunolocalization using the freeze shattering technique and the DMIA monoclonal antibody as previously described . For particle bombardment 2 DAG seedlings were bombarded using the PDS-1000 helium particle delivery system (DuPont, Biotechnology Systems Division, Wilmington, DE) as previously described . Briefly, 2 DAG seedlings were planted at high density on 1/2 × MS plates and bombarded with 0.7 μg of 1 μm gold particles that were coated with 2 μg of GFP:TUB6 expression plasmid . Cells were imaged 36 to 48 h after bombardment.
FM4-64 stained samples were imaged using a Spot RT CCD camera mounted on a Nikon Eclipse E800 fluorescence microscope using the filter set 532-587 nm excitation, 595 nm long pass dichroic mirror, 608-683 nm emission. Excised cotyledons were pressed firmly within a chambered slide. A 40X 0.75 NA objective was used for 2 and 5 DAG fields, and a 20X 0.5 NA objective was used for 12 and 18 DAG cells. Intact GFP-TUB6-expressing seedlings were mounted in water in chambered slides. Samples were imaged using a Bio-Rad 2100 laser scanning confocal microscope mounted on a Nikon eclipse E800 stand. Images were obtained with a 60X 1.2 NA water immersion lens. Samples were excited with 488 nm light and fluorescence signal was collected using a 490 nm long pass dichroic, and a 500-560 nm band pass emission filter was used for detection. The xy pixel size was 0.4 μm and the z-step size was 1.2 μm. Two examples of the raw Biorad *.pic files from Figure 2 and the associated metadata are included as additional data (Additional file 2 and Additional file 3)
Morphometry and image analysis
Three cotyledon fields from three different seedlings were imaged at 3 and 5 DAG. To test the rate of cell area expansion in the same imaging field during two point time-lapse imaging, cell outlines were drawn manually in ImageJ (http://rsbweb.nih.gov/ij/) software and the cell areas were measured. To measure perimeter segment growth for cells, maximum Z-projections of confocal images from two point time-lapse imaging were used to obtain the cell outline. The three-way cell wall junctions were used as fiduciary marks to follow perimeter segments. Cell segments were individually marked and measured in 3 and 5 DAG cells using ImageJ. To measure the height of the cell wall at three-way cell wall junctions, confocal image stacks were resliced perpendicular to the measured wall. Cell height was measured from a maximum projection of the resliced image and included only the wall domain where two adjacent cells were in contact at three-way junctions. Cell areas, cell segments and cell wall heights at 3 DAG and 5 DAG were plotted and subjected to least squares linear regression analysis using Minitab software (Minitab, Quality Plaza, PA). To calculate the isotropy factor for the 3 to 5 DAG growth interval, manually extracted 3 DAG cells were digitally magnified to yield a cell area that was equal to the real 5 DAG cell. After rotation to optimize overlap, the percent of overlapping pixels of the digitally amplified 3 dag cells and actual 5 dag cells were quantified by ImageJ software. The growth of cells from 3 DAG to 5 DAG was calculated as (5 DAG area - 3 DAG area)/(3 DAG area)/48 hr*100%.
The work was supported by NFS MCB Grant No. 0640872 to D.B.S. Thanks to David Umulis and Dan Cosgrove for helpful discussions. Thanks to Eileen Mallery for editorial assistance.
- Tsiantis M, Langdale JA: The formation of leaves. Curr Opin Plant Biol. 1998, 1: 43-48. 10.1016/S1369-5266(98)80126-X.PubMedView ArticleGoogle Scholar
- Fleming A: The control of leaf development. New Phytol. 2005, 166: 9-20. 10.1111/j.1469-8137.2004.01292.x.PubMedView ArticleGoogle Scholar
- Coen E, Rolland-Lagan A-G, Matthews M, Bangham JA, Prusinkiewicz P: The genetics of geometry. PNAS USA. 2004, 101 (14): 4728-4735. 10.1073/pnas.0306308101.PubMedPubMed CentralView ArticleGoogle Scholar
- Savaldi-Goldstein S, Peto C, Chory J: The epidermis both drives and restricts plant shoot growth. Nature. 2007, 446: 199-202. 10.1038/nature05618.PubMedView ArticleGoogle Scholar
- Marcotrigiano M: A role for leaf epidermis in the control of leaf size and the rate and extent of mesophyll division. Am J Bot. 2010, 97: 224-233. 10.3732/ajb.0900102.PubMedView ArticleGoogle Scholar
- Bai Y, Falk S, Schnittger A, Jakoby MJ, Hulskamp M: Tissue layer specific regulation of leaf length and width in Arabidopsis as revealed by the cell autonomous action of ANGUSTIFOLIA. Plant J. 2010, 61: 191-199. 10.1111/j.1365-313X.2009.04050.x.PubMedView ArticleGoogle Scholar
- Szymanski DB, Marks MD: GLABROUS1 overexpression and TRIPTYCHON alter the cell cycle and trichome cell fate in Arabidopsis. Plant Cell. 1998, 10 (12): 2047-2062. 10.1105/tpc.10.12.2047.PubMedPubMed CentralView ArticleGoogle Scholar
- Pyke KA, Marrison JL, Leech RM: Temporal and spatial development of the cells of the expanding first leaf of Arabidopsis thaliana (L.) Heynh. J Exp Bot. 1991, 42: 1407-1416. 10.1093/jxb/42.11.1407.View ArticleGoogle Scholar
- Tsukaya H, Tsuge T, Uchimiya H: The cotyledon: A superior system for studies of leaf development. Planta. 1994, 195: 309-312. 10.1007/BF00199692.View ArticleGoogle Scholar
- Tsuge T, Tsukaya H, Uchimiya H: Two independent and polarized processes of cell elongation regulate leaf blade expansion in Arabidopsis thaliana (L.) Heynh. Development. 1996, 122: 1589-1600.PubMedGoogle Scholar
- Le J, Mallery EL, Zhang C, Brankle S, Szymanski DB: Arabidopsis BRICK1/HSPC300 is an essential WAVE-complex subunit that selectively stabilizes the Arp2/3 activator SCAR2. Curr Biol. 2006, 16: 895-901. 10.1016/j.cub.2006.03.061.PubMedView ArticleGoogle Scholar
- Machacek M, Hodgson L, Welch C, Elliott H, Pertz O, Nalbant P, Abell A, Johnson GL, Hahn KM, Danuser G: Coordination of Rho GTPase activities during cell protrusion. Nature. 2009, 461: 99-103. 10.1038/nature08242.PubMedPubMed CentralView ArticleGoogle Scholar
- Cosgrove DJ: Growth of the plant cell wall. Nat Rev Mol Cell Biol. 2005, 6 (11): 850-861. 10.1038/nrm1746.PubMedView ArticleGoogle Scholar
- Szymanski DB, Cosgrove DJ: Dynamic coordination of cytoskeletal and cell wall systems during plant cell morphogenesis. Curr Biol. 2009, 19: R800-R811. 10.1016/j.cub.2009.07.056.PubMedView ArticleGoogle Scholar
- Esau K: Plant Anatomy. New York: John Wiley & Sons; 1965.Google Scholar
- Panteris E, Galatis B: The morphogenesis of lobed plant cells in the mesophyll and epidermis: organization and distinct roles of cortical microtubules and actin filaments. New Phytol. 2005, 167: 721-732. 10.1111/j.1469-8137.2005.01464.x.PubMedView ArticleGoogle Scholar
- Panteris E, Apostolakos P, Galatis B: Microtubule organization and cell morphogenesis in two semi-lobed cell types of Adiantum capillus-veneris L. leaflets. New Phytol. 1993, 125: 509-520. 10.1111/j.1469-8137.1993.tb03899.x.View ArticleGoogle Scholar
- Wernicke W, Jung G: Role of cytoskeleton in cell shaping of developing mesophyll of wheat (Triticum aestivum L.). Eur J Cell Biol. 1992, 57: 88-94.PubMedGoogle Scholar
- Panteris E, Apostolakos P, Galatis B: Sinuous ordinary epidermal cells: behind several patterns of waviness, a common morphogenetic mechanism. 1994, New Phytol, 127: 771-780.Google Scholar
- Qiu JL, Jilk R, Marks MD, Szymanski DB: The Arabidopsis SPIKE1 gene is required for normal cell shape control and tissue development. Plant Cell. 2002, 14: 101-118. 10.1105/tpc.010346.PubMedPubMed CentralView ArticleGoogle Scholar
- Frank MJ, Smith LG: A small, novel protein highly conserved in plants and animals promotes the polarized growth and division of maize leaf epidermal cells. Curr Biol. 2002, 12 (10): 849-853. 10.1016/S0960-9822(02)00819-9.PubMedView ArticleGoogle Scholar
- Paredez AR, Somerville CR, Ehrhardt DW: Visualization of cellulose synthase demonstrates functional association with microtubules. Science. 2006, 312 (5779): 1491-1495. 10.1126/science.1126551.PubMedView ArticleGoogle Scholar
- Gutierrez R, Lindeboom JJ, Paredez AR, Emons AM, Ehrhardt DW: Arabidopsis cortical microtubules position cellulose synthase delivery to the plasma membrane and interact with cellulose synthase trafficking compartments. Nat Cell Biol. 2009, 797-806. 10.1038/ncb1886.Google Scholar
- Crowell EF, Bischoff V, Desprez T, Rolland A, Stierhof YD, Schumacher K, Gonneau M, Hofte H, Vernhettes S: Pausing of Golgi bodies on microtubules regulates secretion of cellulose synthase complexes in Arabidopsis. Plant Cell. 2009, 21 (4): 1141-1154. 10.1105/tpc.108.065334.PubMedPubMed CentralView ArticleGoogle Scholar
- Szymanski DB: Plant cells taking shape: new insights into cytoplasmic control. Curr Opin Plant Biol. 2009, 12: 735-744. 10.1016/j.pbi.2009.10.005.PubMedView ArticleGoogle Scholar
- Fu Y, Li H, Yang Z: The ROP2 GTPase controls the formation of cortical fine F-actin and the early phase of directional cell expansion during Arabidopsis organogenesis. Plant Cell. 2002, 14: 777-794. 10.1105/tpc.001537.PubMedPubMed CentralView ArticleGoogle Scholar
- Fu Y, Gu Y, Zheng Z, Wasteneys GO, Yang Z: Arabidopsis interdigitating cell growth requires two antagonistic pathways with opposing action on cell morphogenesis. Cell. 2005, 11: 687-700. 10.1016/j.cell.2004.12.026.View ArticleGoogle Scholar
- Panteris E, Apostolakos P, Galatis B: Microtubules and morphogenesis in ordinary epidermal cells of Vigna sinensis leaves. Protoplasma. 1993, 174: 91-100. 10.1007/BF01379041.View ArticleGoogle Scholar
- Zhang C, Mallery EL, Schlueter J, Huang S, Fan Y, Brankle S, Staiger CJ, Szymanski DB: Arabidopsis SCARs function interchangeably to meet actin-related protein 2/3 activation thresholds during morphogenesis. Plant Cell. 2008, 20: 995-1011. 10.1105/tpc.107.055350.PubMedPubMed CentralView ArticleGoogle Scholar
- Russ JC: The Image Processing Handbook fourth edition. Boca Raton: CRC Press; 2002.Google Scholar
- Nakamura M, Naoi K, Shoji T, Hashimoto T: Low concentrations of the propyzamide and oryzalin alter microtubule dynamics in Arabidopsis epidermal cells. Plant and Cell Physiol. 2004, 45: 1330-1334. 10.1093/pcp/pch300.View ArticleGoogle Scholar
- Xu T, Wen M, Nagawa S, Fu Y, Chen JG, Wu MJ, Perrot-Rechenmann C, Friml J, Jones AM, Yang Z: Cell surface- and rho GTPase-based auxin signaling controls cellular interdigitation in Arabidopsis. Cell. 2010, 143 (1): 99-110. 10.1016/j.cell.2010.09.003.PubMedPubMed CentralView ArticleGoogle Scholar
- Verbelen J-P, Vissenberg K, Kerstens S, Le J: Cell expansion in the epidermis: microtubules, cellulose orientation and wall loosening enzymes. J Plant Physiol. 2001, 158: 537-543. 10.1078/0176-1617-00277.View ArticleGoogle Scholar
- Rolland-Lagan A-G, Bangham JA, Coen E: Growth dynamics underlying petal shape and asymmetry. Nature. 2003, 422-Google Scholar
- Mansfield SG, Briarty LG: The dynamics of seedling and cotyledon cell development in Arabidopsis thaliana during reserve mobilization. Int J Plant Sci. 1996, 157: 280-295. 10.1086/297347.View ArticleGoogle Scholar
- Geisler MJ, Sack FD: Variable timing of developmental progression in the stomatal pathway in Arabidopsis. New Phytol. 2002, 153: 469-476. 10.1046/j.0028-646X.2001.00332.x.View ArticleGoogle Scholar
- Masubelele NH, Dewitte W, Menges M, Maughan S, Collins C, Huntley R, Nieuwland J, Scofield S, Murray AH: D-type cyclins activate division in the root apex to promote seed germination in Arabidopsis PNAS USA. 2005, 102: 15694-15699.Google Scholar
- Szymanski DB, Marks MD, Wick SM: Organized F-actin is essential for normal trichome morphogenesis in Arabidopsis. Plant Cell. 1999, 11: 2331-2347. 10.1105/tpc.11.12.2331.PubMedPubMed CentralView ArticleGoogle Scholar
- Mathur J, Spielhofer P, Kost B, Chua N: The actin cytoskeleton is required to elaborate and maintain spatial patterning during trichome cell morphogenesis in Arabidopsis thaliana. Development. 1999, 126 (24): 5559-5568.PubMedGoogle Scholar
- Smith LG, Oppenheimer DG: Spatial control of cell expansion by the plant cytoskeleton. Annu Rev Cell Dev Biol. 2005, 21: 271-295. 10.1146/annurev.cellbio.21.122303.114901.PubMedView ArticleGoogle Scholar
- Jura J, Kojs P, Iqbal M, Szymanowska-Pulka J, Wloch W: Apical intrusive growth of cambial fusiform initials along the tangential walls of adjacent fusiform initials: evidence for a new concept. Aust J Bot. 2006, 54: 493-504. 10.1071/BT05130.View ArticleGoogle Scholar
- Shaw SL, Dumais J, Long SR: Cell surface expansion in polarly growing root hairs of Medicago truncatula. Plant Physiol. 2000, 124 (3): 959-970. 10.1104/pp.124.3.959.PubMedPubMed CentralView ArticleGoogle Scholar
- Ambrose JC, Shoji T, Kotzer AM, Pighin JA, Wasteneys GO: The Arabidopsis CLASP gene encodes a microtubule-associated protein involved in cell expansion and division. Plant Cell. 2007, 19: 2763-2775. 10.1105/tpc.107.053777.PubMedPubMed CentralView ArticleGoogle Scholar
- Kirik V, Herrmann U, Parupalli C, Sedbrook JC, Ehrhardt DW, Hulskamp M: CLASP localizes in two discrete patterns on cortical microtubules and is required for cell morphogenesis and cell division in Arabidopsis. J Cell Sci. 2007, 120 (Pt 24): 4416-4425. 10.1242/jcs.024950.PubMedView ArticleGoogle Scholar
- Wightman R, Marshall R, Turner SR: A Cellulose Synthase-Containing Compartment Moves Rapidly Beneath Sites of Secondary Wall Synthesis. Plant and Cell Physiology. 2009, 50 (3): 584-594. 10.1093/pcp/pcp017.PubMedView ArticleGoogle Scholar
- Wightman R, Turner SR: Severing at sites of microtubule crossover contributes to microtubule alignment in cortical arrays. Plant J. 2007, 52: 742-751. 10.1111/j.1365-313X.2007.03271.x.PubMedView ArticleGoogle Scholar
- Yang Z: Small GTPases: versatile signaling switches in plants. Plant Cell. 2002, S375-S388. SupplementGoogle Scholar
- Fu Y, Xu T, Zhu L, Wen M, Yang Z: A ROP GTPase signaling pathway controls cortical microtubule ordering and cell expansion in Arabidopsis. Curr Biol. 2009, 19 (21): 1827-1832. 10.1016/j.cub.2009.08.052.PubMedPubMed CentralView ArticleGoogle Scholar
- Murata T, Sonobe S, Baskin TI, Hyodo S, Hasezawa S, Nagata T, Horio T, Hasebe M: Microtubule-dependent microtubule nucleation based on recruitment of γ-tubulin in higher plants. Nat Cell Biol. 2005, 7: 961-968. 10.1038/ncb1306.PubMedView ArticleGoogle Scholar
- Chan J, Sambade A, Calder G, Lloyd C: Arabidopsis cortical microtubules are initiated along, as well as branching from, existing microtubules. Plant Cell. 2009, 21: 2298-2306. 10.1105/tpc.109.069716.PubMedPubMed CentralView ArticleGoogle Scholar
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