Flowers are the most complex structures of plants. Studies of Arabidopsis thaliana, which has typical eudicot flowers, have been fundamental in advancing the structural and molecular understanding of flower development. The main processes and stages of Arabidopsis flower development are summarized to provide a framework in which to interpret the detailed molecular genetic studies of genes assigned functions during flower development and is extended to recent genomics studies uncovering the key regulatory modules involved. Computational models have been used to study the concerted action and dynamics of the gene regulatory module that underlies patterning of the Arabidopsis inflorescence meristem and specification of the primordial cell types during early stages of flower development. This includes the gene combinations that specify sepal, petal, stamen and carpel identity, and genes that interact with them. As a dynamic gene regulatory network this module has been shown to converge to stable multigenic profiles that depend upon the overall network topology and are thus robust, which can explain the canalization of flower organ determination and the overall conservation of the basic flower plan among eudicots. Comparative and evolutionary approaches derived from Arabidopsis studies pave the way to studying the molecular basis of diverse floral morphologies.
1. INTRODUCTION: WHEN DID THE FLOWER EVOLVE?
The flower is the most complex structure of plants. Flowers distinguish the most recently diverged plant lineage, the angiosperms or flowering plants, from the other land plants (Figure 1). Embryophytes originated approximately 450 million years before present (MYBP) and have as distinctive features a thick cuticle resistant to desiccation, sporopollenin, pores or true stomata that aid in gas exchange, a glycolate oxidase system that improves carbon fixation at high oxygen tensions, and importantly, distinctive multicellular diploid (sporophytic) and haploid (gametophytic) stages within their life cycles (Judd et al., 2002). The major extant land plant lineages are Bryophytes (Liverworts, Hornworts and Mosses), which do not have a vascular system, and Tracheophytes, vascular plants. Within the large latter group, Lycophytes, ferns, and seed bearing plants (Spermatophytes) can be distinguished. The Spermatophyte group has been further divided into Gymnosperms (originating 380–325 MYBP) and Angiosperms. According to the fossil record, flower-like structures originated 160–147 MYBP (Frohlich, 2006). A general trend within land plant evolution is the appearance of heterospory: the existence of a megagametophyte, including the female gametes, and a microgametophyte, including the male gametes, a progressive reduction in gametophyte size (sexual reproductive structures), and within the seed plants, the presence of a diploid embryo. While these characteristics are shared among both extant and extinct seed plant lineages, the defining features of the angiosperm flower are: (1) a closed carpel bearing the ovules, which are each generally comprised of two integuments and (2) a nucellus that contains the embryo sac within which, after double fertilization, a diploid embryo and a triploid endosperm (nutritional tissue for the embryo) will develop to form a seed (Judd et al., 2002). Another characteristic of angiosperms is true hermaphroditism (Judd et al., 2002; Frohlich, 2006).
Flower structure has been studied in a variety of ways. Studies of the natural history and evolutionary biology of flowers have emphasized understanding the ultimate (evolutionary) causes of the wide range of variants such as color, symmetry, meristic arrangements (e.g. flower organ number), size, pollination syndrome, etc. Other studies have addressed the cellular, tissue type, morphological and physical factors that can account for both the phenotypic plasticity and developmental constraints in flower form (for a review of the developmental framework of angiosperm morphology, see Endress, 2006). A different approach flourished in the late 1980s and early 1990s, the molecular genetics of flower development in two model eudicot species: Arabidopsis thaliana and Antirrhinum majus (see reviews in: Jack, 2004; Kaufmann et al., 2005; Krizek and Fletcher, 2005; Theissen and Melzer, 2007).
Genetic studies of floral homeotic mutants in both plant species yielded the now classic combinatorial ABC developmental model for floral organ determination (Bowman et al., 1989; Coen and Meyerowitz, 1991). While much work has been and continues to be done in Antirrhinum and other eudicot species, including Petunia hybrida, the genomic and life-cycle characteristics of Arabidopsis make it the preferred experimental system for in-depth studies on the molecular components underlying cell differentiation and morphogenesis during flower development.
The basic floral architecture is mostly conserved among the so-called core eudicots, that make up over 73% of extant flowering plants (Drinnan et al., 1994) including Arabidopsis. Flowers within this group generally have four concentric whorls of organs that are specified, from the outside to the center of the flower, in the sequence: sepals, petals, stamens, and carpels. Arabidopsis has this typical floral architecture. An interesting exception to the conserved floral ground plan of eudicots is found in a Mexican rainforest monocotyledon, Lacandonia schismatica (Triuridaceae), which bears central stamens surrounded by carpels (Martínez and Ramos, 1989; Vergara-Silva et al., 2003; Ambrose et al., 2006).
Even though the basic floral architecture is overall conserved among core eudicots, variation in the symmetry and size of flowers, the number of whorls of each organ type, the number of organs per whorl, and their arrangement, size, shape and color is common (e.g., Judd et al., 2002).
The overall conservation of the flower plan suggests that robust gene regulatory network (GRN) modules controlling the basic features of flower development were established early in the evolution of angiosperms and have persisted in the great majority of lineages throughout 140 million years of flower evolution. Recent integrated approaches to study the concerted action of the molecular components in flower development (Mendoza and Alvarez-Buylla, 1998; Espinosa-Soto et al., 2004), have led to a hypothesis that helps explain such robustness and conservation at the level of the GRN underlying floral organ specification. However structural (e.g., mechanical) constraints could also be involved in conserving floral architecture (see section 4). Approaches that integrate genetic and structural aspects of flowers should be pursued further to understand flower development in Arabidopsis and other angiosperms.
2. STRUCTURAL ASPECTS OF ARABIDOPSIS FLOWER DEVELOPMENT
In this section, we provide a summary of structural features of Arabidopsis flower development. This is essential background to the molecular genetics reviewed in section 3.
2.1 Structural Organization of the Inflorescence Meristem and Origin of the Flower Meristem
During the vegetative phase of the Arabidopsis life cycle, the shoot apical meristem (SAM) produces leaves on its flanks and on transition to flowering, the shoot bolts and the SAM becomes the inflorescence shoot apical meristem (IM). On bolting, some of the pre-existing leaf primordia become cauline leaves subtending lateral inflorescence shoots (paraclades) and the shoot apex starts to produce flowers (Hempel and Feldman, 1995). A primary IM produces lateral meristems that may go on to produce flowers or secondary inflorescences. Arabidopsis inflorescences are subtended by fully developed bracts, but flowers only by rudimentary ones. It is generally said that the IM generates the floral meristems (FM) on its flanks, but to be more precise, Arabidopsis FM are formed in the axils of the rudimentary bracts (Figure 2A; Long and Barton 2000; Hepworth et al., 2006; Kwiatkowska, 2006; reviewed in Kwiatkowska, 2008).
The SAM of the Arabidopsis inflorescence consists of a small dome of cells organized into different regions (Figure 2B) with different gene expression profiles (see section 4.1), cellular behaviors and structures. The tunica at the SAM surface and corpus are distinguished on the basis of cell division planes. In Arabidopsis, the tunica consists of two clonally distinct cell layers called L1 and L2 (Vaughan, 1952; Steeves and Sussex, 1989). Cell divisions within these meristem layers are exclusively anticlinal and the new cell walls are formed perpendicular to the surface of the meristem. The progeny of cells in the L1 will therefore remain in this same layer within the meristem similar to the underlying L2 progeny. Since outside the meristem the L1 derived cells continue to divide only anticlinally the L1 eventually gives rise to epidermal cells. The cells originating from L2 also divide periclinally (outside the SAM) and contribute for example to the leaf mesophyll or stem ground tissue formation during organogenesis. This is also the germ line in the angiosperm SAM (Ruth et al., 1985, Klekowski, 1988; Kwiatkowska, 2008). Below the tunica, cell divisions are both anticlinal and periclinal. This region of the SAM is the corpus or L3 from which the innermost tissues, like vascular tissues, are formed (Figure 2B; Brand et al., 2001).
The SAM is also organized into three different cytohistological zones each with characteristic cytoplasmic densities and cell division rates: the central zone (CZ), the peripheral zone (PZ) surrounding the CZ and the rib zone (RZ) underneath the CZ (Figure 2B; Bowman, 1994; Bowman and Eshed, 2000).
Flower primordia are derived from the PZ of the IM and are initiated from a block of four so-called founder cells (Bossinger and Smyth, 1996; Reddy et al., 2004). This estimate was based on sector boundary analysis. However, using a non-invasive replica method and a 3-D reconstruction algorithm, Kwiatkowska (2006; 2008) argues that more cells are assigned to the flower primordium, and this is consistent with the observations by Grandjean et al. (2004). The difference could be due to the fact that not all of the cells estimated to be involved in the latter approaches are incorporated into the flower meristem proper. Some of them may form a part of the subtending rudimentary bract (Figure 2A; see next section for further discussion).
The first cells produced by the RZ following the transition to flowering are rectangular with their long axis perpendicular to the major axis of the stem, but the subsequent elongation of these cells reverses this situation (Vaughan, 1955). The RZ gives rise to stem tissue. The CZ encompasses the reservoir of stem cells that divide less frequently than cells at the periphery (Grandjean et al., 2004; Reddy et al., 2004). The CZ maintains itself and yields daughter cells that form both the PZ and RZ (Bowman and Eshed, 2000). Fifteen stages of Arabidopsis flower development have been distinguished (Smyth et al., 1990). The first stages of flower meristem development are: stage 1, when a flower buttress arises, stage 2 when the flower meristem is formed and stage 3 when sepal primordia appear. Recently researchers have been able to study early flower meristem development in greater detail (Reddy et al., 2004; Kwiatkowska, 2006; reviewed by Kwiatkowska, 2008) and have proposed subdividing stage 1 (see section 2.3).
2.2 Floral Organ Primordia
Once a flower primordium is initiated, the geometry changes and a rapid and coordinated burst of cell expansion and division occurs in three dimensions generating a concentric group of cells as an almost spherical flower primordium, from which all floral tissues are derived (Bossinger and Smyth, 1996; Reddy et al., 2004; Kwiatkowska, 2006). Jenik and Irish (2000) found that the regulation of cell divisions during early and late stages of flower development seems to depend upon different mechanisms. Early in flower development, when the floral meristem of Arabidopsis is divided into four concentric rings (each with a characteristic multigenic expression profile; see section 3.3), cell division patterns depend upon the cell's radial position in the floral meristem, and not on the future identity of the floral organ to be formed in each ring. After stage 6, during organogenesis, the ABC homeotic genes (see section 3.3) seem to control the rate and orientation of cell divisions. As a result, the continuity of the concentric rings is broken giving distinct floral organ primordia within each whorl, then cells subdifferentiate into distinct types within each organ. The initiation and identity of floral organs are also regulated by different and largely independent molecular modules. This is suggested, for example, by the fact that conversion of petals into sepal-like organs in mutant plants does not alter the number of cells involved in their initiation (Crone and Lord, 1994; Bossinger and Smyth, 1996).
Tissues of floral organs are organized according to coordinated patterns and rates of cell division in the different cell layers of the meristem that dynamically acquire distinct fates. Clonal analysis shows that L1 contributes to the epidermis, the stigma, part of the transmitting tract and the integument of the ovules, while L2 and L3 contribute to the mesophyll and other internal tissues (Jenik and Irish, 2000).
Sector boundary analysis of surface cells has shown that sepals and carpels are initiated from eight cells, stamens from four cells, and petals from two cells (Bossinger and Smyth, 1996). Each organ primordium arises as a set of cells separated by boundary regions of slow-dividing cells (Figure 2C and section 3.4.2; Breuil-Broyer et al., 2004). Flower development ends when mature organs are formed and all the flower meristem cells are used up (Takeda et al., 2004; Krizek and Fletcher, 2005).
2.3 Stages of Flower Development
We provide an illustrated description of 20 states of floral development and fruit formation (Figures 2–7), mostly based on Bowman (1994), Smyth et al. (1990), Ferrándiz et al. (1999) and Roeder and Yanofsky (2006), with updates and substages as proposed by Long and Barton (2000), Reddy et al., 2004; Hepworth et al. (2006), Kwiatkowska (2006) and Kwiatkowska (2008).
STAGE 1: The first sign of flower primordium formation is the bulging of the peripheral surface of the IM in a lateral direction. This stage was referred to as P1 by Reddy et al. (2004). It is hypothesized that a lateral protrusion formed during bulging is a rudimentary bract (Figure 2A; Kwiatkowska, 2006). At this early stage, growth is fast and strongly anisotropic, with maximal growth in a meridional (i.e. radial when viewed from the top of the meristem) direction (Kwiatkowska, 2006) eventually leading to formation of a shallow crease, which corresponds to the first visible groove and to the P2 stage (according to Reddy et al., 2004) of flower development (Figure 2A). This shallow crease corresponds to the axil of the putative rudimentary bract (Kwiatkowska, 2006, 2008). Soon after the bract is formed, another bulge occurs in its axil in an upward direction. This second bulging corresponds to the formation of a flower primordium proper and to stage P3 according to Reddy et al. (2004). This stage corresponds to stage 2 according to Smyth et al. (1990). Hence, during early stages of flower development in Arabidopsis, two types of primordia (bract and flower primordium proper) and organ boundaries are observed. The first boundary is the adaxial boundary of the rudimentary bract, while the second is the boundary between the IM and the flower primordium proper (Figure 2A). The expression patterns of several genes confirm the developmental stages distinguished here (see more data on gene expression in the next section).
A significant increase in mitotic activity is observed upon formation of the primordium. The mitotic activity can be estimated as the increase in the number of cells per 24 h or the accompanying area growth rates on the condition that the mean cell size does not increase (Grandjean et al., 2004; Kwiatkowska, 2006; Reddy et al., 2004). During these early stages of flower development, periclinal cell divisions occur in the corpus while L1 and L2 cells only divide anticlinally (Vaughan, 1955). Hence, the two-layered tunica organization is maintained in the flower meristem, but all of its cells are mitotically active.
STAGE 2: During this stage, the hemispherical primordium continues to grow forming almost a right angle with the surface of the SAM, which itself lengthens and widens rebuilding the portion of the periphery that has been used for primordium formation (Figures 3 and 4A–B). At this stage the flower primordium becomes clearly delimited from the IM, and starts to grow larger very quickly in all directions (Figures 3 and 4A–B; Reddy et al., 2004; Kwiatkowska, 2006).
STAGE 3: This stage begins when sepal primordia become visible. By now the flower primordium is 30–35 µm in diameter and is becoming stalked with an incipient pedicel. It has also started to grow vertically. The two lateral (I) sepal primordia appear first, but are soon outgrown by the abaxial (ab) then the adaxial (ad) sepal primordia. Sepal primordia arise initially as ridges that lengthen and curve inwards until they begin to overtop the remaining dome-shaped portion of the flower primordium (Figures 3 and 4C–D).
STAGE 4: During this stage, the elongation of the pedicel continues concurrently with an increase in the diameter of the developing flower primordium to 65–70 µm. The medial sepal primordia have already partly overtopped the remaining floral meristem (Figure 4E).
STAGE 5: This stage is when the petal and stamen primordia become visible. Primordia of the four medial (long) stamens are first seen as wide outgrowths on the flanks of the central dome of the FM. The four petal primordia that arise between the sepals close to their base are just visible during this stage. The two lateral (short) stamens develop from primordia that appear later during this stage (Figures 3 and 4F–G).
STAGE 6: The sepals grow to completely cover the floral bud and the primordia of the four long stamens bulge out and become distinct from the central dome of cells that comprise the FM. The two lateral stamen primordia arise slightly lower on the dome and develop later. The petal primordia grow somewhat but are still relatively small. A rim around the central dome of the flower primordium now begins to grow upward to produce an oval tube that will become the gynoecium (Figures 3 and 4H–I).
STAGE 7: This stage begins when the growing primordia of the long stamens become stalked at their base. The stalks give rise to the filaments, and the wider upper region to the anthers. By this stage, petal primordia have become hemispherical although they are still relatively small (ca. 25 µm in diameter; Figures 3 and 5A–B).
STAGE 8: The beginning of stage 8 is defined by another landmark in stamen development: anther locules are visible as convex protrusions on the inner (adaxial) surface of the long stamens. At this stage stamens are 55–60 µm long most of which is the developing anther. Locules also appear soon after in the short stamens. Petal growth now accelerates and petal primordia become apparent (Figures 3 and 5C–E).
STAGE 9: This stage begins when the petal primordia elongate. There is a rapid lengthening of all organs especially of petals that acquire a tongue-like shape and increase in length from about 45 µm to up to 200 µm. Nectary glands appear and the stamens grow rapidly. By the end of stage 9, the medial stamens are around 300 µm long. Most of this growth occurs in the anther region, which still accounts for over 80% of their total length. At this stage the floral bud remains completely closed (Figures 3 and 5F–G).
STAGE 11: This stage begins when the upper surface of the gynoecium develops stigmatic papillae (Figures 3 and 6A–C) although their outward growth is limited at first to regions not in contact with the overlapping sepals. By the end of this stage petal primordia reach the top of the medial stamens.
STAGE 12: Petals continue to lengthen relatively rapidly. Lateral sepals continue to grow while the stamens and gynoecium lengthen coordinately. The anthers have almost reached their mature length of 350–400 µm and the filaments now lengthen rapidly. The upper part of the gynoecium differentiates into the style (Figure 6D) and a sharp boundary separates it from the cap of stigmatic papillae. Stage 12 ends when the sepals open (Figures 3 and 6D–F).
STAGE 13: Petals become visible between the sepals and continue to elongate rapidly. The stigma is receptive at this stage (Figures 3 and 6G–H). Stamen filaments extend even faster so the stamens outstrip the gynoecium in length and self pollination takes place. The gynoecium is now mature and its three distinct regions can be distinguished: an apical stigma, a style, and a basal ovary. After pollination, pollen tubes grow to fertilize the ovules, the stamens extend above the stigma, and furrows at both valve/replum boundaries appear.
STAGE 14: This is also defined as the stage zero hours after flowering (0 HAF), and it marks the beginning of silique (the fertilized pistil or fruit) and seed development. Cells in the exocarp continue to divide anticlinally and expand longitudinally in the replum and the valve, where there is also some expansion in other directions. There is also division and expansion in the mesocarp and many chloroplasts develop (Figures 3 and 6I).
STAGE 15: The stigma extends above the long anthers. In the carpel walls, cell division and expansion continue. The medial vascular bundles continue to grow and xylem lignifies, while the lateral bundles branch out through the mesocarp (Figures 3 and 6J–K).
STAGE 17: This stage is defined by the abscission of the senescent floral organs from the silique, ∼2 days after fertilization. The green silique grows to reach its final length and matures, a phase lasting about 8 days making this the longest stage. The dehiscence zone also differentiates (Figures 3, 7A and 7E; Substages 17A and 17B, see Roeder and Yanofsky, 2006).
STAGE 18: The silique begins to yellow from the tip to the base. One of the endocarp cell layers (the second from the inside) lignifies further, and the inner endocarp cell layer disintegrates, while the mesocarp begins to dry out. It has been suggested that lignification may contribute to the silique shattering process, acting in a springlike manner to create mechanical tensions (Figures 3 and 7B).
2.4 Morphology, Histology and Development of Floral Organs
Sepals: In sepals L1 -derived cells form the epidermis, the mesophyll originates from the L2, and the L3 contributes to the vascu-lature in the basal part (Jenik and Irish, 2000). Sepals and petals together form the perianth. Both organ types have a simple laminar structure, consisting of an epidermis, mesophyll and rather delicate vascular bundles (veins). The four sepal primordia (the abaxial, adaxial, and two lateral sepal primordia) are the first floral organ primordia to appear. They arise at stage 3 of flower development in a cruciform pattern (Figures 4C–D; Smyth et al., 1990; Bowman, 1994). Whether all four sepals occupy one whorl or the two lateral sepals occupy a separate outer whorl, has been the subject of discussion (Figure 4E; Smyth et al., 1990; Bowman, 1994; Choob and Penin, 2004), but all sepal primordia are formed at around the same time, shortly after they are specified (Figure 4E; Bowman, 1994).
The adaxial and abaxial surfaces of the sepal epidermis are different (Figures 8B and 8D–E). On the abaxial surface, cells have irregular shapes and sizes with some quite long cells (with nuclei of various sizes) and fringes of smaller cells. Unlike the adaxial surface, the abaxial surface has stomata and may have unbranched trichomes (Figure 8E; Smyth et al., 1990; Bowman, 1994; Hase et al., 2000; Krizek et al., 2000).
Petals: In the petal primordium the meristematic layer L1 contributes to the epidermis and L2 to the mesophyll; as yet cells originating from L3 have not been found to form part of the petal (Figure 2B; Jenik and Irish, 2000). These primordia become apparent almost at the same time as stamen primordia at stage 5 of flower development. Visible signs of petal differentiation are seen by stage 9 (Figure 5F; Smyth et al., 1990; Bowman, 1994). The four petals of Arabidopsis are white and flat and approximately the same size and shape. They are narrower and greenish toward the base (Figure 5C; Takeda et al., 2004).
Cells on the adaxial surface are conical with epicuticular thickenings running from the cell base to the apex, whereas those on the abaxial surface are flatter and more cobblestone-like with cuticular thickening (Figures 8F–G). Stomata are absent from both petal surfaces (Bowman, 1994; Krizek et al., 2000). Cells toward the base of petals resemble those of stamen filaments (Bowman, 1994).
Stamens: Primordia appear at stage 5 of flower development (Figure 4F) due to periclinal divisions in the subprotodermal cell layer (L2) and sometimes in L3 (Crone and Lord, 1994; Jenik and Irish, 2000). Stamen primordia are visible at stage 6. By stage 7, differentiation can be observed and long stamen primordia appear stalked at their bases (Figures 4I and 5A–B; Bowman, 1994; Smyth et al., 1990). At this stage stamen primordia are composed of an L1 -derived epidermis, one layer of L2-derived subepidermis, and an L3-derived core (Figure 2B; Jenik and Irish, 2000). Locules appear in the anthers by stage 8 (Figure 5C). Growth of the internal anther tissue at this stage is due to divisions of L2-derived cells (Jenik and Irish, 2000). At stage 14, anthers extend above the stigma (Figure 5I; Bowman, 1994). In the mature anther, the L3 cells contribute only to the vasculature (Jenik and Irish, 2000). Stamens of the Arabidopsis flower are not formed simultaneously: four long medial stamens arise a little earlier than the two short lateral ones (Smyth et al., 1990).
Each stamen consists of two distinct parts, the filament and the anther. At the tip of the filaments, the anther develops both reproductive and non-reproductive tissues that produce, harbor, and release pollen grains upon maturity (Goldberg et al., 1993). The anther is a bilocular structure with longitudinal dehiscence (Figure 6G; Bowman, 1994). Each locule develops from successive divisions of subprotodermal archesporial cells formed in the anther primordium that gives rise to three morphologically distinct layers: the endothecium, the middle layer, and the tapetum which surrounds the pollen mother cells (PMCs). The PMCs undergo meiosis and form the haploid microspores. The tapetum is a source of nutrients and is indispensable for microspore maturation (Xing and Zachgo, 2008). Anther development and microspore formation in Arabidopsis is a complex process that has been divided into 14 stages (See also section 3.4.5; Sanders et al., 1999).
Once formed, PMCs are surrounded by a layer of callose. After meiosis, the anther contains most of its specialized cells and tissues, and tetrads of microspores are present within the pollen sacs; with microspores in each tetrad surrounded by a callose wall. Callose dissolves and microspores are released. As pollen grains develop, the anther enlarges and is pushed upward in the flower by the elongating filament (Scott et al., 2004).
Carpels: The fourth and innermost whorl is occupied by the gynoecium that is composed of two fused carpels. Carpel primordia start to form at stage 6 of flower development (Figure 4I) due to periclinal cell divisions in the L3 layer (Jenik and Irish, 2000). Carpels enclose and protect the developing ovules, mediate pollination, and after fertilization develop into a fruit within which fertilized ovules develop into seeds (Bowman et al., 1999). The gynoecium consists of two valves separated by a false septum with ovules arising from parental placental tissue on each side of the septum (Bowman, 1994). The valves grow upward from the flower meristem to form a closed cylinder. At early stage 8, the walls of the cylinder are composed of an L1 -derived epidermis, one L2-derived subepidermal layer and a two-cell thick, L3-derived core. At this stage the distal L2 cells start to divide periclinally (with respect to the top surface of the cylinder), contributing to the longitudinal growth of the carpel (Figure 2B; Jenik and Irish, 2000). Later the inner surfaces of septal outgrowths within this cylinder will fuse, the tip will close and ovules will develop along the margins of the fused walls (placenta) of the bilocular chamber (Bowman, 1994; Sessions and Zambryski, 1995). The gynoecium is oriented in the flower so that the septum coincides with the medial plane (Figures 4D–E and 4G; Sessions and Zambryski, 1995).
At the distal end of the gynoecium, the stigma, an epidermal structure composed of stigmatic papillae (bulbous elongated cells), functions in pollen binding and recognition and participates in the induction of pollen germination (Figures 6A–B). After germination, the pollen tubes will grow between the papillar cells into the transmitting tract at the center of the style and the septum of the ovary (Bowman, 1994; Sessions and Zambryski, 1995).
At about stage 11, the inner and outer integuments of the ovule are formed. By stage 12, the integuments of the developing ovule grow to cover the nucellus and megagametogenesis occurs (Figures 6E–F; Bowman, 1994).
Nectaries: These organs produce and secrete nectar. Nectar is a protein- and carbohydrate-rich solution, which varies in composition among different plant species (Davis et al., 1998). Nectar may be a reward for pollinators or for insects that protect the plant against herbivores, or even a lure for animal prey in carnivorous plants (Davis et al., 1998; Baum et al., 2001; Lee et al., 2005a).
In Arabidopsis, the nectarium (multiple nectary) found in individual flowers (Davis et al., 1998) is composed of two parts: nectary glands that form below the stamen filament, and the connective tissue linking the glands in a continuum around the androecium (Bowman, 1994; Baum et al., 2001). The nectarium is always situated in the third whorl of the flower and its location is independent of the identity of the other organs occupying this whorl. These glands are formed from stage 9 to 17 of flower development (Figure 5G; Bowman, 1994; Bowman and Smyth 1999; Baum et al., 2001 ; Tapia-López et al., 2008).
3. MOLECULAR GENETICS OF ARABIDOPSIS FLOWER DEVELOPMENT
Plant organogenesis, including flower formation, occurs from actively proliferating meristems over the entire life cycle. In the next section we provide a very brief summary of the molecular mechanisms that maintain an active SAM. In section 3.2, we explain how the flower meristem is specified and becomes determinate after the flower organs are formed.
3.1 Shoot Apical Meristem Proliferation and Maintenance
The balance between cell proliferation and cell recruitment to differentiated tissues in the SAM is dependent on mechanisms regulated by WUSCHEL (WUS; Laux et al., 1996; Sablowski, 2007). The homeodomain-containing WUS transcription factor has the role of the maintaining the identity of stem cells in the organizing center of the CZ; wus mutants lack stem cells in the SAM (Mayer et al., 1998). WUS expression is limited to the cells immediately below the stem cells, an expression domain regulated by the receptor-kinase signaling system that includes the CLAVATA1, 2 and 3 (CLV1, 2, 3) gene products (Mayer et al., 1998; Brandetal., 2000; Schoof et al., 2000). CLV1 is expressed in most L3 stem cells while CLV3 is expressed in all three stem cell layers but mostly in L1 and L2 stem cells (see Figure 2B; Clark et al., 1997; Fletcher et al., 1999). In civ mutants, there is an imbalance between cells retained within meristems versus those recruited to form lateral organs, clv mutations cause an expansion of the WUS expression domain resulting in an enlarged stem cell niche. CLV3 expression is, in turn, positively regulated by WUS, suggesting that meristem size depends greatly on a WUS-CLV regulatory loop (Clark et al., 1993, 1995; Kayes and Clark, 1998; Brand et al., 2000). Overexpression of CLV3 represses WUS expression and decreases meristem activity, suggesting that CLV3, a secreted CLE-domain peptide, is the signal that regulates WUS expression via the CLV1/CLV2 LRR protein-kinase transduction complex (Fletcher et al., 1999; Jeong et al., 1999; Trotochaud et al., 1999; Clark, 2001a and 2001b; Ni and Clark, 2006). It has been shown that other LRR-protein kinases closely related to CLV1 like BARELY ANY MERISTEM1 and 2 (BAM1, 2) are also involved in meristem maintenance possibly by sequestering CLV3 on the flanks of the meristem where they are expressed (DeYoung et al., 2006; DeYoung and Clark, 2008).
SHOOT MERISTEMLESS (STM) is a KNOTTED1-like homeobox (KNOX) gene that encodes a protein expressed in the SAM's CZ, RZ and regions of the PZ that have not been assigned to a primordium, i.e. it is expressed throughout the meristem except for anlagen, the sites of primordium formation (Figure 2B). STM promotes the profileration of stem cell derivatives until a critical cellular mass is attained sufficient to form either leaves or floral primordia. It also inhibits the expression of ASYMMETRIC LEAVES1 and 2 (AS1, 2) genes in the SAM, preventing these cells from undergoing premature differentiation (Byrne et al., 2000; Byrne et al., 2002). Thus, the STM gene is considered to play a pivotal role in meristem maintenance (Long et al., 1996; Carles et al., 2004). ULTRAPETALA1 (ULT1) encodes a cysteine-rich protein with a B-box like domain that restricts the size of shoot and floral meristems. It functions antagonistically to the proliferative roles of WUS and STM during most of the Arabidopsis life cycle but it in an independent genetic pathway (Carles et al., 2004).
3.2 Floral Meristem Specification and Determination
The changes in cellular characteristics, growth and geometry observed in the transition of the SAM to an IM (Kwiatkowska, 2006) are correlated with dynamic changes in the spatial and temporal expression of certain genes. The Arabidopsis IM produces rudimentary bracts in whose axils flower meristems emerge. STM and AINTEGUMENTA (ANT) expression patterns correlate with the development of this rudimentary bract primordium (Long and Barton, 2000).
The expression of LEAFY (LFY), a transcription factor found only in plants (Schultz and Haughn, 1991; Waigel et al., 1992; Maizel et al., 2005; Weigel, 2005) and ANT has been used in order to trace the cells that form the flower primordium (Grandjean et el., 2004). First, tens of cells are rapidly recruited to those already committed to become part of the flower meristem. This stage may correspond to the upward bulging at the shallow crease formed between the rudimentary bract and the IM described by Kwiatkowska (2006). These cells which express LFY then continue to proliferate. Interpretating this, the first cells that express LFY would correspond to the rudimentary bract (but not its axil or shallow crease), and later the domain of LFY expression would expand to include the cells committed to the flower primordium proper (Kwiatkowska, 2006). This interpretation can explain the discrepancy in the number of founder cells estimated using sector boundary analysis (Bossinger and Smyth, 1996) and using in vivo LFY expression patterns (Grandjean et al., 2004). Bossinger and Smyth (1996) concluded that a FM derives from four founder cells directly on the surface of the IM (or SAM). In support of this, evidence from confocal laser scanning microscopy indicates that flower primordia are formed from two rows of cells in a radial arc (Reddy et al., 2004). In contrast, the number of cells expressing LFY at these early stages (Reddy et al., 2004) suggest that a flower meristem has more founder cells. An explanation that resolves the discrepancy is that the LFV-expressing cells could include those that eventually form the rudimentary bract, as well as those which form the flower primordium (Kwiatkowska, 2006; reviewed in Kwiatkowska, 2008)
The gene CUP-SHAPED COTYLEDON2 (CUC2) is expressed in the slow-dividing cells that expand in a latitudinal direction (Reddy et al., 2004) to define the second boundary between the floral primordium proper and the IM (Breuil-Broyer et al., 2004). Several regulators of CUC including a miRNA have been described as important components of the GRN involved in this developmental process (Laufs et al., 2004; Aida and Tasaka, 2006a).
Flower versus inflorescence meristem identity is controlled by a complex GRN that integrates environmental and internal cues (Figure 9). On induction of flowering, the IM genes, such as TERMINAL FLOWER 1 (TFL1, Alvarez et al., 1992; Shannon and Meeks-Wagner, 1991 and 1993, Ohshima et al., 1997) and EMBRYONIC FLOWER 1 and 2 (EMF1, 2; Chen et al., 1997; Aubert et al., 2001), are repressed in the FM, while the floral meristem identity (FMI) genes, mainly LFY, APETALA1 (AP1), APETALA2 (AP2), and CAULIFLOWER (CAL), are upregulated (Figure 10; Blazquez et al., 1997; for review Mandel et al., 1992; Waigel et al., 1992; Bowman et al., 1993; Kempin et al., 1995; Mandel and Yanofsky, 1995a; Blazquez et al., 2006).
Mutual repression of the IM and FMI genes seem to underlie the co-existence, identity and boundaries of both types of meristem in the SAM in the transition to flowering (Chen et al., 1997; Liljegren et al., 1999; Ratcliffe et al., 1999). For example, if genes such as TFL1 or EMF1 or 2 are mutated, LFY and/or AP1 are ectopically expressed in the IM that is then transformed into a FM (Shannon and Meeks-Wagner. 1991; 1993; Weigel et al., 1992, Bowman et al., 1993; Gustafson-Brown et al., 1994; Bradley et al., 1997; Chen et al., 1997; Moon et al., 2003). On the contrary, if AP1, CAL and LFY are repressed, the FM attains IM identity (Ratcliffe et al., 1998; Ratcliffe et al., 1999). TFL1 is an important regulator of inflorescence development (Alvarez et al., 1992; Ratcliffe et al., 1998; Parcy et al., 2002). It encodes a phosphatidyl ethanolamine-binding protein (PEBP) that is transcribed in the center of the IM but the protein moves to other cells where AP1 and LFY are downregulated (Bradley et al., 1997; Conti and Bradley, 2007). EMF genes are required for vegetative growth, but they seem to regulate flowering time and inflorescence development too (Sung et al., 1992; Aubert et al., 2001 ; Yoshida et al., 2001). Loss-of-function mutants in these genes produce flowers immediately after germination skipping the vegetative phase (Yang et al., 1995; Chen et al., 1997). EMF1 encodes a transcription factor that represses AP1 but not LFY, and EMF2 encodes a novel zinc finger protein related to the polycomb group (Aubert et al., 2001; Yoshida et al., 2001).
LFY is necessary and sufficient to specify FMI (Weigel et al., 1992; Weigel and Nilsson, 1995). In lfy mutants, leaves and secondary shoots are produced instead of flowers (Schultz and Haughn, 1991; Weigel et al., 1992) and LFY overexpression causes the conversion of leaves and axillary meristems to flowers (Weigel and Nilsson, 1995). LFY is expressed in the leaf primordia during vegetative growth, but when induced by external (vernalization and light) and/or internal (gibberellins) signals, it is strongly expressed and relocates to the SAM flanks where floral meristems are formed (Figure 9; Blazquez et al., 1997; Hempel et al., 1997; for LFY regulation see: Nilsson et al., 1998; Blazquez and Weigel, 2000; Liu et al., 2008). LFY expression persists at high levels in the FM until stage 3 of development and then diminishes in the center of the flower (Figure 10; Blazquez et al., 1997; Wagner et al., 2004). LFY protein abundance, however, is homogenous in the FM, probably because it moves between cells (Parcy et al., 1998; Sessions et al., 2000; Wu et al., 2003).
LFY and AP1 have overlapping functions in establishing the FM; while the ap1 mutant has shoots with inflorescence characteristics, the lfy ap1 double mutant has an almost complete conversion of flowers into shoots (Huala and Sussex, 1992; Bowman et al., 1993). Both genes when overexpressed cause a terminal flower phenotype suggesting that each one is sufficient to determine the IM (Mandel and Yanofsky, 1995a; Weigel and Nilsson, 1995). CAL, the closest paralogue of AP1, and FRUITFULL (FUL) from the same gene clade within the MADS-box phylogenetic tree (Alvarez-Buylla et al., 2000; Martínez-Castilla and Alvarez-Buylla, 2003; Parenicová et al., 2003), may also act redundantly to AP1 in FM specification. Single cal and ful mutants do not show any FMI disorders, but in combination with ap1 in double or triple mutants, the ap1 phenotype is greatly intensified (Bowman et al., 1993; Kempin et al., 1995; Ferrándiz et al., 2000a). FUL is expressed at the same time as LFY during the establishment of the FMI (Mandel and Yanofsky, 1995b; Hempel et al., 1997), but is mostly localized in the IM (Figure 10). Later during flower development, FUL is expressed again during carpel and silique development where it plays an important role (Gu et al., 1998). Despite its close similarity to AP1, overexpression of CAL is not able to determine the IM as does overexpression of AP1, indicating that CAL does not interact with the same partners as AP1. The unique functions of AP1 rely on residues within the K and COOH domains that are not found in CAL (Alvarez-Buylla et al., 2006).
LFY directly regulates AP1 and CAL transcription by binding to the consensus sequence CCANTG (CArG-box; Parcy et al., 1998; Wagner et al., 1999; Wagner et al., 2004; William et al., 2004). However, expression reminiscent of AP1 is seen in the lfy mutant, while it is completely abolished in the double mutant lfy ft (flowering locus t, Ruiz-Garcia et al., 1997; Schmid et al., 2003; Wigge et al., 2005).Thus FT, a homolog of TFL1 (Koornneef et al., 1991; Kardailsky et al., 1999), together with FD, a bZip transcription factor (Abe et al., 2005), redundantly regulate AP1 with LFY. AP1 and CAL in turn regulate LFY by positive feedback, allowing it to exert its transcriptional regulation during flower development (Bowman et al., 1993; Liljegren et al., 1999). Recently, additional LFY targets have been found (William et al., 2004), among them LATE MERISTEM IDENTITY1 (LMI1), which encodes a homeodomain leucine-zipper transcription factor and functions as a FMI gene. Interestingly, LMH acts together with LFY to activate CAL expression (Figure 9; Saddic et al., 2006).
AP2 encodes a putative transcription factor of a plant-specific gene family (AP2/EREBP) with diverse functions (Riechmann and Meyerowitz, 1998). Mutations in AP2 enhance both ap1 and lfy mutant phenotypes, indicating that AP2 also plays a role in specifying FMI (Huala and Sussex, 1992; Schultz and Haughn, 1993; Shannon and Meeks-Wagner, 1993; Simpson et al., 1999).
MADS-box genes are key components of the regulatory module that integrates flowering transition signaling pathways (for review see Jack, 2004; Parcy, 2005, Blazquez et al., 2006), IM and FM identities (Mandel et al., 1992; Bowman et al., 1993; Mandel and Yanofsky, 1995a, 1995b), and floral organ specification (see section 3.3; Coen and Meyerowitz, 1991). To specify the FM, LFY and/or AP1 are also required to downregulate flowering induction genes such as AGAMOUS-LIKE24 (AGL24), SUPPRESSOR OF OVEREXPRESSION OF CO 1 (SOC1), SHORT VEGETATIVE PHASE (SVP), and FUL (Figures 9 and 10). Overexpression of any of these genes causes FM to revert to IM-like structures as when LFY and/or AP1 are mutated (Mandel and Yanofsky 1995b; Yu et al., 2004a; Liu et al., 2007).
Floral reversion is often found in plants heterozygous for lfy6 (LFY/lfy) and homozygous for agamous-1 (ag-1), suggesting a key role for LFY and AG in the maintenance of determinate floral meristems (Okamuro et al., 1996). The reason for this is that late in floral organogenesis AG, induced by WUS, LFY and PERIANTHIA (PAN) among others, positively regulates KNUCKLES (KNU) which in turn represses WUS expression to terminate the stem cell niche after a limited number of organs have been formed (Parcy et al., 1998; Busch et al., 1999; Lenhard et al., 2001; Lohmann et al., 2001 ; Das et al., 2009; Maier et al.. 2009; Sun et al., 2009). In fact, while WUS expression declines after stage 6 in wild-type flowers, it persists in pan or ag flowers (Lenhard et al., 2001; Lohmann et al., 2001; Das et al., 2009; Maier et al., 2009). ULT also participates in meristem determinancy together with AG downregulating WUS (Carles et al., 2004).
Although it is very rare to observe spontaneous or induced reversion from FM to IM, a set of genes that actively maintain FM identity could conform to a “flower developmental module” that prevents reversion. The genetic mechanisms involved in maintaining FMI are closely linked to hormone balance and environmental factors (Tooke et al., 2005). For example, we now know that STM is a positive regulator of local cytokinin (CK) biosynthesis and accumulation (Jasinski et al., 2005; Yanai et al., 2005), and a repressor of gibberellin (GA) production (Jasinski et al., 2005). On the other hand, WUS enhances CK activity by repressing ARABIDOPSIS TYPEA RESPONSE REGULATORS (ARRs) (Leibfried et al., 2005). The resulting high CK:auxin ratio and low GA levels promote indeterminate growth (Shani et al., 2006). While a high auxin concentration restricts STW and CUC expression (see section 3.4.2), it also downregulates CK biosynthesis and activity, thus yielding a high auxin:CK ratio and high levels of GA, which induce floral meristem formation. Raising GA levels or response, for example by crossing with the spindly (spy) mutant, is sufficient to suppress FM reversion to IM in lfy, ap1, ap2 and ag mutants. This demonstrates the importance of GA in the maintenance of FM identity (Okamuro et al., 1996; Okamuro et al., 1997).
Light signal transduction pathways are also involved in FM maintenance. Spontaneous floral reversion in wild-type Arabidopsis has only been observed at low frequencies in the first flowers of Landsberg erecta grown in short days. However, long hypocotyl (hy1-1), a mutant in which photochrome activity is blocked, suppresses floral reversion of both lfy and ag single mutants in short days (Okamuro et al., 1996). Floral reversion seems to be a developmental abnormality with no apparent adaptative significance, unless plant resources are somehow saved under certain conditions if flowering is reversed. Further ecological and evolutionary developmental studies of Arabidopsis ecotypes will continue to elucidate the genetic, epigenetic, physiological, and environmental mechanisms involved in the maintenance of the FMI.
3.3 Specification of Floral Organs: The ABC Genes
Very soon after FM specification (11–13 days after germination in Landsberg erecta ecotype), the flower meristem is subdivided into four regions. Each one will give rise to the primordia of the different floral whorls, which from the outside to the inside are: sepals, petals, stamens, and carpels. The genes responsible for floral whorl specification attain their spatio-temporal pattern as a result of regulatory interactions among themselves, interactions with meristem identity genes and with some other genes, such as WUS and UNUSUAL FLORAL ORGANS (UFO; Levin and Meyerowitz, 1995). The complexity of the interactions involved is shown in the ‘floral organ specification gene regulatory network’ (FOS-GRN) model, analyzed in Section 4.1. This model includes a set of interacting genes sufficient to pattern the IM and FM during the first stages of flower development.
One of the key FM identity genes is LFY. The protein encoded by this gene requires co-factors to set the spatial limits of expression of the floral organ identity genes AP3, Pl, and AG. For example, LFY participates with UFO in the regulation of AP1 and AP3 transcription (Lee et al., 1997; Chae et al., 2008), and with WUS co-regulates the expression of AG (Lenhard et al., 2001; Lohmann et al., 2001). LFY also regulates the expression of the SEPALLATA (SEP) genes SEP1, SEP2 and SEP3, additional MADS-box genes required for organ identity specification (Krizek and Fletcher, 2005).
UFO is expressed in the second and third whorls during floral stage 3, probably restricting the B-gene expression domain to these whorls, together with LFY (Lee et al., 1997; Traas and Doonan, 2003). The UFO gene encodes a protein containing an F-box domain, which is a characteristic of E3 ubiquitin ligases that are components of SCF (Skp Cullin F-box containing) complexes and mark proteins for proteosome-dependent degradation (Deshaies, 1999). It was recently shown that LFY interacts with UFO in order to directly bind the AP3 promoter. Furthermore, the proteosome activity mediated by UFO is required for the transcriptional activation of AP3 by LFY (Chae et al., 2008).
Key components of the GRN that underlies the early patterning of the flower meristem are the so-called ABC homeotic genes, AP1, AP2, AP3, Pl, and AG, which are all transcription factors belonging to the MADS-box gene family, except AP2 (Coen and Meyerowitz, 1991; Wagner et al., 1999; Ng and Yanofsky, 2001; Lamb et al., 2002).
The classic ABC model was inferred using Arabidopsis and Antirrhinum homeotic flower mutants (Coen and Meyerowitz, 1991). In these mutants two floral organ types are replaced by two other floral organ types as follows: A- class mutant flowers have carpels-stamens-stamens-carpels (from the outermost to the innermost whorl), B-class mutant flowers have sepals-sepals-carpels-carpels, and C-class mutant flowers have sepals-petalspetals-sepals (Coen and Meyerowitz, 1991). It was shown that mutations in all three functions lead to the transformation of all floral organs into leaf-like organs, suggesting that flowers are modified leaves (reviewed in Robles and Pelaz, 2005). The Arabidopsis ABC mutants are shown in Figure 11.
Hence, three different classes of homeotic genes with overlapping activities were proposed to be necessary for floral organ specification. The A function specifies sepals, the A and B functions specify petals, the B and C functions specify stamens and the C function specifies carpels (Figure 12; Bowman et al., 1991). The A and C functions negatively regulate each other and the B function is restricted to the second and third whorls. The latter was originally thought to be independent of A and C functions (Bowman et al., 1991; Drews et al., 1991), but it was later shown that the A function gene AP1 regulates the B genes. AP1 binds to the promoter of AP3 (Hill et al., 1998; Tilly et al., 1998). AP1 can also specify petals by regulating the spatial domain of B genes together with UFO in the first flowers to arise, and independently of UFO in later flowers (Ng and Yanoksky, 2001).
Once identified at the molecular level, the mRNA expression patterns of the ABC genes were shown to overlap with the floral regions where the corresponding mutants had a phenotype (Yanofsky et al., 1990; Mandel et al., 1992; Goto and Meyerowitz, 1994; Jack et al., 1994). AP1 and AP2 are A-function genes. AP1 is expressed in the two outer whorls of the floral meristem (Figures 10, 12, 13A; Mandel et al., 1992) and is important for the establishment of sepal and petal identity as well as the FM (section 3.2). AP1 expression is first up-regulated by LFY and FT/ FD (section 3.2), but later is maintained by the B class genes in a positive feedback loop (Sundström et al., 2006). Strong ap1 alleles (ap1-1) often lack petals in the second whorl, while weaker mutant alleles of this gene do not have a full homeotic conversion of floral organs (see section 3.2; Irish and Sussex, 1990).
In contrast to the MADS-box ABCs, the expression pattern of AP2 does not correlate with the site where it exerts its function in floral organ identity. AP2 mRNA is found throughout the flower meristem (Figures 10 and 12; Jofuku et al., 1994). Recent data has shown that AP2 is repressed at the translational level by a microRNA (miR172), which is active only in whorls 3 and 4 (Chen, 2004), thus explaining why the function of AP2 is restricted to the first two whorls of flower organs. In a recent experiment using double mutants of ag and an ap2 allele, which is insensitive to repression by miR172, it was shown that both AG and miR172 independently downregulate AP2, but miR172 is more important than AG (Zhao et al., 2007). ap2 mutants rarely develop petals and their sepals are transformed into carpelloid structures due to ectopic AG expression (Figure 11), which is negatively regulated by AP2 itself (Drews et al., 1991). AP2 is also implicated in the upregulation of the B genes, AP3 and Pl(Zhao et al., 2007).
The B class genes (AP3 and Pl) are expressed in the second and third whorls and mutant flowers of any or both of these two genes lack petals and stamens, as predicted in the ABC model (Figure 11,12 and 13; Coen and Meyerowitz 1991; Goto and Meyerowitz, 1994; Jack et al., 1994; Honma and Goto, 2000).The fact that both single mutants yield the same phenotype shows their interdependence. AP3 and Pl are regulated in two steps: they are first induced by LFY/UFO in response to flowering signals and they later maintain their expression in a self-regulatory loop (Honma and Goto, 2000). The proteins encoded by these two genes form heterodimers to exert their B function during petal and stamen development (Figure 14; Jack et al., 1992; Goto and Meyerowitz, 1994; Zik and Irish, 2003a) and this oligomerization is necessary for them to move into the nucleus (McGonigle et al., 1996).
Both genes are also regulated positively in a regulatory loop by AP1 and negatively by EARLY BOLTING IN SHORT DAYS (EBS), a gene that encodes a nuclear protein that participates in petal and stamen development and regulates flowering time by repressing FT (Gómez-Mena et al., 2001; Piñeiro et al., 2003) ANT, a member of the AP2 gene family, is another regulator of the B function, positively inducing AP3 (Klucher et al., 1996; Nole-Wilson and Krizek, 2006; see section 3.4.2).
The only C-type gene discovered up to now is the MADS-box gene AG (Bowman et al., 1989). ag mutant flowers lack stamens and carpels, and also bear indeterminate flowers with reiterating sepals and petals (Figure 11), suggesting that AG is important for floral meristem determinancy (see section 3.2), besides its role in stamen and carpel identity (Yanofsky et al., 1990; Mizukami and Ma, 1997). The regulation of AG has been much studied; at least ten proteins repress and five activate it to maintain its expression in the appropriate whorl (Figures 12 and 13A).
AG is repressed by a transcriptional co-repressor complex formed by LEUNIG (LUG) and SEUSS (SEU) (Figure 15; Franks et al., 2002). LUG encodes a transcription protein similar to TUP1 from yeast and interacts with SEU, which encodes a plant specific protein (see Table S1 (); Conner and Liu, 2000; Franks et al., 2002; Sridhar et al., 2004). Neither of these proteins are able to bind DNA sequences and AP1 and SEP3 recruit SEU/LUG to the second intron of AG to perform their inhibitory function and prevent the ectopic expression of AG (Sridhar et al., 2006). Recently, another transcriptional repressor of AG was identified, LEUNIG_HOMOLOG (LUH). This gene is the closest homolog of LUG and its inhibitory function on AG is completely dependent on SEU (Sitaraman et al., 2008).
Another repressor of AG is BELLRINGER (BLR), a homeodomain protein that binds to regions in the second intron of AG and prevents ectopic AG expression in the two outer whorls of the flower (see Table S1 (); Bao et al., 2004). AG is also negatively regulated epigenetically by a histone acetyltransferase GCN5 (Bertrand et al., 2003). Other genes that participate in floral organogenesis are repressors of AG, namely RABBIT EARS (RBE, see section 3.4.4), ANT and STERILE APETALA (SAP) (see Table S1 ()). AG is also positively regulated at the post-transcriptional level by several ENHANCER OF AG-4 (HUA) and HUA ENHANCER (HEN) genes. All of these genes play a major role in pre-mRNA processing of AG (Cheng et al., 2003).
The ABC proteins exert their regulatory function as multimers. In Antirrhinum majus, a ternary complex between A and B function proteins was found to bind CArG DNA boxes more efficiently than single proteins (Egea-Cortines et al., 1999). More specifically, a higher-order complex consisting of SQUAMOSA (SQUA, the AP1 ortholog), DEFICIENS, and GLOBOSA (DEF and, GLO are A. majus AP3 and Pl orthologues, respectively) bound DNA more efficiently than DEF/GLO or SQUA alone (Egea-Cortines et al., 1999). These results suggest that transcriptional complexes that combine A and B function proteins are more stable than those formed with proteins of any one function alone.
The fact that the ABC genes are necessary but not sufficient to determine floral organ identity was later confirmed in Arabidopsis. Honma and Goto (2001) used a yeast three-hybrid method to show that SEP3 and AP1 are able to interact with the heterodimer AP3/PI but not with AP3 or PI alone. Moreover, they described this interaction as essential, since the heterodimer AP3/PI lacks the activation domain necessary for a transcription factor to function, a domain which both SEP3 and AP1 possess (Honma and Goto, 2001). These findings suggest that the inclusion of SEP3 or AP1 together with AP3/PI could result in an active tetrameric transcriptional complex (Figure 14). It was also demonstrated that the ABC proteins on their own or combined according to the ABC model (A, AB, BC, or C) were not sufficient to determine floral organs when expressed in leaves under the action of the 35S constitutive promoter (Pelaz et al., 2001). However, floral organs could indeed be recovered in leaves once appropriate combinations of genes were expressed (Honma and Goto, 2001; Pelaz et al., 2001).
The SEP genes received their names because the floral organs that develop in all four whorls in triple sep mutants resemble sepals (Pelaz et al., 2000). This sep1 sep2 sep3 triple mutant phenotype is markedly similar to that of double mutants that lack both B and C class activity, such as pi ag and ap3 ag (Figure 11G; Bowman et al., 1989; Pelaz et al., 2000) in which the floral meristem becomes indeterminate as well. Single or double mutants for these SEP genes yield flowers indistinguishable from wild type, thus suggesting that the three SEP genes are functionally redundant and are important in determining three of the four floral organs: petals, stamens, and carpels (Honma and Goto, 2001; Pelaz et al., 2001; Robles and Pelaz, 2005).
Given that the triple sep1 sep2 sep3 mutant does not show alterations in sepal identity, an additional gene is likely to be involved in sepal specification. Indeed, another SEP-like MADSbox gene, SEP4 (previously AGL3), has now been characterized (Ditta et al., 2004), and the quadruple sep1 sep2 sep3 sep4 mutants produce flowers with leaf-like organs in all whorls, thus confirming the SEP genes contribute to each floral organ identity (Figure 14). Coincidently, SEP genes are expressed in the whole floral meristem during flower development (Figure 13B; Flanagan and Ma, 1994), are important in regulating B and C gene expression (Liu et al., 2009), and encode proteins that apparently interact with the ABC proteins (Figure 14; Robles and Pelaz, 2005).
Based on data from Antirrhinum and yeast two-hybrid and three-hybrid protein interactions, and on the phenotypes of the ABC mutants, three models have been proposed to explain how the MADS domain proteins interact to constitute functional transcriptional complexes and bind DNA. None of the models completely explains the experimental data available, but the quartet model seems the most plausible (Jack, 2001; de Folter et al., 2005). This model proposes that MADS domain proteins form tetrameric complexes during floral organ determination (Figure 14; Theissen, 2001; Theissen and Saedler, 2001; Becker and Theissen, 2003; Jack, 2004). Within each transcriptional complex, there would be two MADS dimers, each one binding to a single CArG-binding site causing the DNA of the promoter region to bend, enabling the MADS dimers to act cooperatively in a tetrameric complex to regulate the gene. For example, binding of one dirtier within the tetramer to DNA could increase the affinity of the second dirtier for local DNA binding (Melzer et al., 2009). Besides, one of the dimers could function as the activation domain of the tetramer allowing for efficient transcriptional activation (Honma and Goto, 2001). Interestingly, several dimers and potential tetramers have been documented in a complete Arabidopsis MADS-domain family protein-protein interactome via yeast two-hybrid interactions (de Folter et al., 2005). This data base has been updated with a yeast three-hybrid screen for MADS-domain proteins (Immink et al., 2009). Future studies should test which of the complexes inferred from the MADS interactomes are functional and what their roles are during Arabidopsis development.
3.3.1 Target genes of the ABCs
Target genes of the ABC genes link the floral organ specification - gene regulatory network (FOS-GRN) with processes in organ primordia establishment and development (for review of MADS target genes, see de Folter and Angenent, 2006). Among the direct targets of the ABC genes, transcriptional regulators and hormone-related genes are prominent (Sablowski, 2009). But the sets of target genes change as organ development progresses; at later stages of floral organ development, several components of what could be common modules have been found that are involved in generic developmental processes (see below) during sepal, petal, stamen and carpel development. Finally, multiple genes having cell-specific roles are turned on especially during stamen and carpel development that is much more complex than perianth development (Sablowski, 2009).
The first examples of genes regulated by the ABC genes were two MADS-box genes, AGL1 and AGL5 (renamed the SHATTERPROOF genes (SPH1 and 2, respectively). There is virtually no expression of either gene in ag mutants (Savidge et al., 1995). SHP2 is only expressed in carpels and AG can activate an SPH2:GUS reporter construct; furthermore, AG binds its promoter in vitro (Savidge et al., 1995).
The first non-ABC gene identified as a target of a MADS A, B or C protein was NAC-LIKE ACTIVATED BY AP3/PI (NAP), a target of the AP3/PI complex. It is important for the transition between cell division and cell expansion during petal and stamen development (see section 3.4.5 and Table S1 () ; Sablowski and Meyerowitz, 1998).
A recent study identified two genes negatively regulated by AP3/PI, GATA NITRATE INDUCIBLE, CARBON-METABOLISM-IN-VOLVED (GCN) and GCN-LIKE(GNL), a GCN paralog (Mara and Irish, 2008). Both genes regulate chlorophyll biosynthesis in plant cells. Thus, their downregulation could be important in preventing chlorophyll accumulation in petals and anthers. The same study shows that both GNC and GNL, together with the B class genes, regulate the expression of a number of other GATA-motif-containing target genes like HEXOKINASE1 (HXK1, Mara and Irish, 2008).
SUPERMAN (SUP; Bowman et al., 1992) is upregulated by AP3/PI and AG and by LFY (Riechmann et al., 1996; Sakai et al., 2000). SUP encodes a transcription factor with a C2H2-zinc finger motif and is involved in the maintenance of the stamen/carpel whorl boundary (Sakai et al., 2000; Dathan et al., 2002; see section 3.4.2). While the B genes and LFY seem to regulate early SUP expression, AG and the B genes are involved in maintaining its expression in flowers from stage 5 onward (Sakai et al., 2000).
Recent microarray experiments have proved useful in revealing new targets of the ABC MADS homeotic genes, as well as many putative components of the complex networks involved in floral organogenesis. For example, it was shown that the AP3/PI dimer regulates, directly or indirectly, 47 target genes. Only two of these are transcription factors, while most participate in basic cellular functions required for stamen and petal development (Zik and Irish, 2003a). By contrast, AG controls, directly or indirectly, the expression of 149 genes most of which are transcription factors, including other members of the MADS-box gene family. Ten of these were also shown to be direct targets of AG using chromatin immunoprecipitation (ChIP), including AG itself, AP3, CRC and ATH1, a gene that encodes a BELL-type homeodomain protein that participates in the development of the basal region of shoot organs (Gómez-Mena et al., 2005).
A more exhaustive experiment used four homeotic mutants (ap1/ap2, ap3, pi and ag) in two types of microarray assays: a whole genome microarray with approximately 26,090 gene-specific oligonucleotides and a flower specific-cDNA microarray with 5,000–6,000 genes. To summarize the assay results, transcription factors were neither over or underrepresented as being regulated by the ABC genes; on the contrary, genes involved in general cellular maintenance (DNA recombination and protein synthesis) were underrepresented. Genes specifically expressed in each of the four different whorls were identified: 13 genes for sepals, 18 for petals, 1162 for stamens, and 260 for carpels. As expected from their structural and cellular complexity, the reproductive floral organs had many more specific target genes than the perianth organs (Wellmer et al., 2004; Sablowski, 2009).
In another genomic study of early floral stages it was found that many genes were downregulated in incipient floral primordia while many of them were activated during the differentiation of floral organs (Wellmer et al., 2006). However, some genes were overrepresented during all stages analyzed (i.e. transcription factors including the family of MADS-box genes, PIN dependent auxin transport genes, as well as auxin and GA metabolism genes). Even though the MADS box genes were overrepresented, the promoter regions of the genes expressed during these different stages are not enriched in CArG-box sequences compared to random samples from the whole genome. This result suggests that MADS-domain transcription factors may be able to bind sequences other than CArG motifs, or that they have few direct targets during the developmental stages analyzed (Wellmer et al., 2006).
In a different approach, an inducible post-translational version of AG was used in gene expression profiling to detect AG target genes. One of the genes identified that is upregulated by AG is SPOROCYTELESS (SPL). AG is able to bind in vitro to the 3′ region (downstream of the stop codon) of the SPL gene (Ito et al., 2004). SPL has been described as a key regulator of sporogenesis later during stamen and carpel development (see sections 3.4.5, 3.4.6 and Table S1 (); Schiefthaler et al., 1999; Yang et al., 1999).
3.4 Floral Organogenesis
The challenge of inferring the topology of the gene regulatory network (GRN) underlying the establishment of floral organ primordia, and their development (cell differentiation, morphogenesis and growth) is still ahead. However, some key components and GRN functional modules characterized to date are summarized in Section 4. Such modules involve several functional feedback loops and underlie different generic developmental processes mainly: primordia type specification; delimitation; floral organ primordia positioning that depends on fundamentally on auxins; primordia number; inter-whorl and within-whorl boundaries; and primordia and organ adaxial-abaxial polarity (Figures 2C and 15; Irish, 2008). At later stages of floral organ development, subcellular differentiation and patterning, as well as overall organogenesis takes place and more specific regulatory modules are involved. The genes within such modules are treated separately for each organ type (Figures 16–17).
As a precursor to integrating GRN modules in the above categories, we now provide a synthesis of the molecular genetic studies of how such generic developmental processes are regulated. Several of these have also been identified as important regulators of leaf development, substantiating the proposal of Goethe that all plant organs are elaborations or modifications of a core leaf-like developmental program (for review of common pathways see Sablowski, 2009). ABC floral organ identity genes are also important in fine-tuning or coordinating the role of genes involved in some of the generic developmental modules during flower development (Figure 15; Sablowski, 2009). Some genes participate in more than one process or module and are important for making connections between different GRN modules. In such cases, they are considered in more than one category.
Regulatory modules controlling distinct components of floral organ development have been elucidated to different extents depending on available mutant phenotypes. In correlation with anatomical and morphological complexity, the size and complexity of the regulatory modules underlying stamen and carpel development are much greater than those that regulate sepal or petal development. Carpel development is covered in the “Fruit Development” chapter (Roeder and Yanofsky, 2006) in this book, and is only briefly considered here.
In the flower meristem, normal organogenesis depends upon a homeostatic equilibrium between stem cell specification and cellular differentiation (Green et al., 2005). Plant morphogenesis is influenced both by the orientation and rate of cell division, as well as by cell expansion and differentiation (see section 2 for a description of floral organ initiation and morphogenesis). How the molecular aspects of these processes are coordinated has been very difficult to elucidate. However, it is generally accepted that cells in meristematic regions respond to positional information important for inducing and controlling morphogenesis (Sussex, 1954; 1955; Meyerowitz 1997; Hauser et al., 1998). One of these positional signals is auxin (see Section 3.4.1; Reinhardt et al., 2000; Benková et al., 2003; Reinhardt et al., 2003; de Reuille et al., 2006). Several mutations that affect the number, size, and/or shape of one or several floral organs have also been characterized. Some of these phenotypes are pleiotropic consequences of mutations in genes acting from earlier steps of plant and flower development. Others are the result of alterations in organ specific genes (Figures 16–17). An extensive list of genes involved in flower organ morphogenesis with their inferred functions, mutant phenotypes and mRNA expression patterns is given in Table S1 ().
3.4.1. Floral meristem and organ primordia positioning: the role of auxin
The shoot apical meristem produces leaves and then flowers in a highly predictable and regular phyllotactic pattern (Tanaka et al., 2006). One of the key compounds that regulate this developmental process is the hormone auxin (Reinhardt et al., 2000). Increased auxin levels mark the initiation sites for organ primordia (including those of floral organs) and local application of auxin is sufficient to trigger leaf or flower formation in the shoot apex (Reinhardt et al., 2000; Tanaka et al., 2006). Once the primordium is established, there is a depletion of auxin around it and another peak of auxin is only able to form in cells at a specific distance from pre-existing primordia, generating a phyllotactic pattern (Reinhardt et al., 2000; Reinhardt et al., 2003; de Reuille et al., 2006; Tanaka et al., 2006; Berleth et al., 2007; Kuhlemeier, 2007). After Initiation, the primordium grows by cell proliferation and cell expansion, and the organ differentiates along the apical-basal and dorsal-ventral axes (Heisler et al., 2005; Golz, 2006).
The overall distribution of auxin depends on its biosynthesis, metabolism, and directional transport. Most auxin is synthesized in young tissues of the shoot and distributed throughout the plant by two physiologically distinct pathways. One of them is passive and occurs only by diffusion through the mature phloem. The other one is an active polar auxin transport (called PAT) that mediates cell-to-cell movement of auxin through two different types of proteins, efflux and influx carriers. Some of the genes that encode these transporters (or carriers) have been cloned: PIN-FORMED (PIN) and P-GLYCOPROTEINS (ABCB/PGP) for auxin efflux, and AUXIN1 (AUX1) and its paralogs LIKE-AUX1 (LAX1-3) for auxin uptake/influx (Figure 15; Bennett et al., 1996; Friml, 2003; Yang et al., 2006; Bandyopadhyay et al., 2007).
The PIN gene family encodes eight protein members in total; three of them (PIN5, 6, and 8) of unknown function. All of the PIN proteins characterized until now are asymmetrically distributed on the plasma membrane and some of them can be found in specific cell types with no pronounced polarity (Vieten et al., 2007). The direction of auxin flow is believed to be determined by the asymmetric cellular localization of PIN proteins (Friml, 2003). The first of these proteins to be characterized was PIN1, and its mutation (pin1) results in pin-shaped inflorescence meristems without flow- ers. PIN1 expression is induced by auxin and it encodes a protein with 10–12 putative transmembrane domains and shares some similarity with bacterial transporters (Gälweiler et al., 1998). pin1mutant plants accumulate high amounts of auxin in vegetative meristems and a deficiency in the apical inflorescence meristem, which results in a defective organ initiation of leaves and flowers, a phenotype that can be imitated in wild type using auxin efflux inhibitors (Okada et al., 1991; Reinhardt et al., 2000). Of the other PIN proteins, only pin3 and pin7 loss-of-function mutants have flowers, and these bear fused petals, no stamens, and occasionally no sepals (Benková et al., 2003). PIN3 is essentially involved in mediating differential shoot growth (Friml et al., 2002) and PIN7 is important during early embryo development (Friml et al., 2003).
Auxin movement mediated by PIN carrier proteins determines the growth axis of the developing organ by establishing an auxin gradient with its maximum at the tip (Benková et al., 2003; Tanaka et al., 2006). As the primordium rapidly expands, auxin is depleted from the tip. Two hypotheses have been proposed to explain this observation: either auxin is transported through the primordium interior into the vascular network (Benková et al., 2003; Tanaka et al., 2006) or it is depleted from primordial regions as a result of specific reversals in PIN1 polarity (Heisler et al., 2005).
The ABCB/PGPs are also transmembrane proteins that belong to the ATP-binding cassette (ABC) transporter superfamily. In Arabidopsis, three of their members, ABCB1, ABCB4, and ABCB9, are able to transport auxin away from apical tissues and are important in maintaining long-distance auxin transport (Titapiwatanakun et al., 2009). One of the PGP proteins (PGP19) co-localizes and interacts with PIN1 and the ABCB protein is apparently important in stabilizing plasma membrane microdomains necessary for enhanced PIN1 activity (Bandyopadhyay et al., 2007; Titapiwatanakun et al., 2009).
Auxin enters the cell passively by simple diffusion and also by the import activity of AUX1 and related LAX proteins. The AUX1 gene encodes a protein with 11 putative transmembrane domains (Hobbie, 2006) similar to plant amino acid permeases (Bennett et al., 1996). The mutant form (aux1) was identified in a screen for auxin resistant and agravitropic mutants (Bennett et al., 1996; Vieten et al., 2007). The AUX1 protein also has polar subcellular localization in some cells and co-localizes with PIN1 in the shoot apical meristem. AUX1/LAX function could be essential for stabilizing the phyllotactic pattern. The proposed model for AUX1/LAX function is that these proteins concentrate auxin in the cytoplasm of cells of the L1 layer, preventing auxin diffusion in the apoplast (Bainbridge et al., 2008).
PINOID (PID) encodes a Ser/Thr protein kinase (Christensen et al., 2000) which has been implicated to function in redirecting subcellular PIN polarities, because the loss of its activity causes a shift in apical-basal PIN polarity (Friml et al., 2004; Berleth et al., 2007; Michniewicz et al., 2007). pid mutants have a defect in organ formation similar to that of pin1, but they do produce a few flowers (Reinhardt et al., 2003) with altered floral organ numbers (more petals but fewer stamens) (Bennett et al., 1995). Recently, Michniewicz et al., (2007) reported that In vivo PIN1 phosphorylation is directly dependent on the kinase PID and a phosphatase PP2A, which may act directly by dephosphorylating PIN1 or indirectly through PID. This phosphorylation status determines the intracellular apical-basal localization of PIN1 and therefore auxin transport-dependent development. PIN1 is targeted to the apex when it is phosphorylated and to the base when it is dephosphorylated (Michniewicz et al., 2007; Vieten et al., 2007).
Accumulation of auxin activates downstream processes through specific receptors and the combinatorial action of members of two large families of transcription factors, AUXIN RESPONSE FACTORS (ARF) and IAA/AUX (Kuhlemeier, 2007). The Aux/IAA proteins are degraded when the levels of free auxin rise, resulting in derepression of ARFs. ETTIN (ETT)IARF3 has a dynamic role in patterning by acting in specific cells within floral meristems and reproductive organs. At early stages, ETT functions in determining the number of organ primordia, whereas later it is involved in the outgrowth and patterning of tissues within organ primordia (Figure 15; Sessions et al., 1997). err mutant plants show altered flower development; some flowers have missing petals and rudimentary radialized stamens, and others have normal fertile stamens, but radialized petals (Pekker et al., 2005). ETT is also involved in prepatterning apical and basal boundaries in the gynoecium primordium (see Table S1 (); Sessions and Zambryski, 1995; Sessions et al., 1997). MONOPTEROS (MP)/ARF5 mutants (mp) have inflorescences with smaller or absent flowers, similar to pin1 mutants (Przemeck et al., 1996).
3.4.2. Floral organ primordia number, size, and boundaries
In Arabidopsis, which is a self-fertilizing (autogamous) and partially cleistogamous (before flower bud opens) plant, floral organ size might not be under strong evolutionary pressure compared to allogamous species. However, it has been an important model to study genes that control size and architectural traits of flowers (Weiss et al., 2005).
Several mutations that affect meristem size and maintenance lead to alterations in flower organ number or size. Mutations in the CLV genes (Clark et al., 1993 and 1995; Kayes and Clark, 1998) cause an increase in meristem size, thus yielding additional whorls and a change in floral organ number with altered phyllotaxis (Clark et al., 1993; Clark et al., 1997; Fletcher et al., 1999; Brand et al., 2000; Doerner, 2000). Mutations in genes that control cell proliferation in the SAM, such as the CLV genes, are similar to ULT in that they have larger SAM and primordia (Fletcher, 2001; Carles et al., 2004) and WIGGUM (WIG; Running et al., 1998).
When WUS is repressed and the number of cells for floral primordia formation is reduced, organ architecture is compromised suggesting that there is a threshold number of cells required to form a normal organ (Weiss et al., 2005). In fact, the loss of organs observed in Á-function mutants, or any other AG repressor mutant could be explained as a result of premature repression of WUS by AG in these organs (Crone and Lord, 1994; Liu and Meyerowitz, 1995; Laux et al., 1996).
Other mutants that have altered floral organ numbers are pan (Running and Meyerowitz, 1996; Chuang et al., 1999), ett (Sessions et al., 1997) and sup (Jacobsen and Meyerowitz, 1997). Both pan and ett have more sepals and petals and fewer stamens, whereas sup produces more stamens at the expense of carpels (Weiss et al., 2005). Double pan sup mutants however have an attenuated sup phenotype in the fourth whorl, probably because in this mutant AG is downregulated and the domain of expression of WUS is expanded (Das et al., 2009).
The PAN gene mutation specifically alters floral organ number, yielding fertile plants with a pentamerous meristic pattern (Running and Meyerowitz, 1996). PAN encodes a member of the bZIP class of transcriptional regulators (Chuang et al., 1999) and is thought to act in the process by which cells assess their position within the developing floral meristem. This gene may affect the switch that commits floral organ primordia cells to enter an organ initiation program (Running and Meyerowitz, 1996). PAN and WUS expression overlaps and in clv mutants both genes are ectopically expressed (Chuang et al., 1999; Maier et al., 2009). WUS overexpression causes PAN overexpansion too suggesting that this gene is positively regulated by WUS (Maier et al., 2009).
Interestingly, pentameric symmetry is characteristic of flowers in early-diverging angiosperm lineages, thus suggesting that PAN may have been involved in changes to meristic patterns during angiosperm diversification; particularly the evolution from pentamerous to tetramerous flowers in the Brassicaceae lineage (Chuang et al., 1999).
Organ size is also regulated by the same components in all whorls. The ANT gene encodes a transcription factor of the AP2 family, which seems to be a general regulator of organ size during organogenesis (Elliott et al., 1996; Klucher et al., 1996; Krizek, 1999; Krizek et al., 2000; Mizukami and Fischer, 2000). The overexpression of ANT causes increased cell division in sepals and increased cell expansion in the inner three whorls, probably affecting both the rate and duration of cell divisions which are important determinants of the final size of lateral organs (Krizek, 1999; Mizukami and Fischer, 2000; Weiss et al., 2005). ARGOS participates in the same transduction pathway as ANT and acts downstream of AUXIN RESISTANT 1 (AXR1). Interestingly, increased organ size observed in ARGOS overexpression lines is due to an extended period of cell division rather than to an increase in growth rate (Hu et al., 2003; Weiss et al., 2005). So, it is plausible to assume that these two genes (and probably others) affect organ size by transducing signals from plant growth regulators, such as auxin, which is a key player in establishing SAM primordia and a general regulator of cell proliferation and expansion (Figure 15).
ANT also participates in defining abaxial-adaxial organ polarity in combination with FILAMENTOUS FLOWERIYABBY1 (FIU YAB1, Nole-Wilson and Krizek, 2006; see next section) and thus may be one of the links between the modules controlling primordia growth and the polarity establishment (Figure 15).
Ectopic expression of UFO (Levin and Meyerowitz, 1995) also causes increased floral organ size (Lee et al., 1997), due to increased cell division (Mizukami, 2001; Weiss et al., 2005). This pathway is regulated by UFO independently of its role in B gene expression, because ectopic expression of the B genes does not induce any increase in organ size, so missexpression of other unknown UFO-dependent factors may account for this phenotype (Ni et al., 2004). UFO and two gene enhancers of the ufo- phenotype, FUSED FLORAL ORGANS 1 and 3 (FFO1 and FFO3), could also participate in establishing and maintaining organ boundaries probably by affecting cell proliferation (Levin et al., 1998).
Morphological boundaries are established in the early stages of the formation of a primordium separating it from surrounding tissues, and later from adjacent organ primordia (Figure 2C; Aida and Tasaka, 2006a). Cells in the boundary are distinctly narrow and elongated with low proliferation rates (Aida and Tasaka, 2006b). Genes expressed in the boundary may affect both meristem and organ development by upregulating cell differentiation genes and downregulating meristematic genes (Borghi et al., 2007). CUC1, 2, and 3 encode NAC-domain transcription factors that promote morphological separation of lateral organs through growth repression (Aida et al., 1997; Vroemen et al., 2003; Taoka et al., 2004). cuc1 cuc2 double mutant seedlings have fused cotyledons with no shoots. However, when adventitious stems are induced in this genotype, flowers have fused sepals and stamens, fewer petals and stamens number, and reduced fertility (Aida et al., 1997). CUC genes are epigenetically regulated (Laufs et al., 2004; Kwon et al., 2006).
Other genes, such as LATERAL ORGAN BOUNDARY (LOB) and JAGGED LATERAL ORGANS (JLO), members of the LATERAL ORGAN BOUNDARY DOMAIN (LBD) gene family, encode putative transcription factors with a leucine-zipper motif that are also expressed in boundary cells (Shuai et al., 2002; Borghi et al., 2007). JLO along with the CUC genes probably coordinate auxin accumulation and loss of meristem-specific gene expression in organ anlagen (Takada et al., 2001 ; Borghi et al., 2007).
3.4.3. Floral organ polarity
Establishing organ polarity is an important aspect of morphogenesis and it is sometimes clearly associated with specific functions of plant organs. Both, adaxial-abaxial and proximal-distal polarities are regulated by genetic circuits that are similar for all lateral organs (Figure 2C; Feng and Dickinson, 2007), although each organ type has distinct cell types and morphogenesis in the abaxial versus adaxial surfaces, and in the proximal versus distal regions (Figures 2C and 8). Organ polarity is also linked to the establishment of hormone gradients.
Briefly, abaxial fate is conferred by members of the YABBY family (Sawa et al., 1999; Siegfried et al., 1999) and by some of the KANADI genes (Eshed et al., 2001; Kerstetter et al., 2001), whereas adaxial cell fate is determined by members of the PHAB family: REVOLUTA (REV), PHABULOSA (PHB), and PHAVOLUTA (PHV) (McConnell et al., 2001 ; Emery et al., 2003; reviewed in Bowman et al., 2002; Zik and Irish, 2003b; Golz, 2006) together with JAGGED (JAG) and NUBBIN (NUB) (Figure 15; Dinneny et ai., 2004; Dinneny et al., 2006).
YABBY proteins (YAB) are transcription factors with a Zn-finger and a helix-loop-helix (YABBY) domain that are promoters of abaxial cell fate in all lateral organs, among other functions (Bowman 1999; Sawa et al., 1999; Siegfried et al., 1999). During flower development they participate in establishing the primordium domain and meristem patterns, and later in maintaining abaxial polarity (Siegfried et al., 1999; Goldshmidt et al., 2008). FIL/YAB1, YAB2, and YAB3 are expressed in a polar manner in all lateral organs of the flower meristem, while CRABS CLAW(CRC/YAB63) is only expressed in carpels and nectaries, and INNER NO OUTER (INO/YAB4) is restricted to outer integuments (see section 3.4.6 and 3.4.7; Alvarez and Smyth, 1999; Bowman and Smyth, 1999; Villanueva et al.. 1999).
KANADI (KAN) genes encode transcription factors of the GARP family. KAN1, KAN2, and KAN3 have been implicated in promoting abaxial cell fates (Eshed et al., 1999; Eshed et al., 2001; Kerstetter et al., 2001). The kan1 mutant was selected as a genetic enhancer of crc gynoecium phenotype, producing a mirror-image of adaxial tissues in the kan1 crc double mutant, indicating that both genes participate in a redundant manner to promote abaxial identity (Eshed et al., 1999). In kan1 kan2 double mutants, all floral organs are also extremely adaxialized (Eshed et al., 2001; Kerstetter et al 2001). Although these KAN genes are not necessary for the activation of YAB genes, they are important in controling their proper abaxial localization (Eshed et al., 2001). Even though KAN and YAB genes may have common targets, they also have different ones, since the phenotype of the fil yab3 double mutant is not quite the same as the extreme phenotype of kan1 kan2 (Bowman et al., 2002).
It has been hypothesized that the “default” state of cells is the abaxial fate (Sussex 1954, 1955). Genes that belong to the PHAB family (class III homeodomain-leucine zipper, HD-ZIP III; Sessa et al., 1998; McConnell et al., 2001; Golz, 2006) of transcription factors, like PHB and PHV, might be activated by a proximal signal coming from the apical meristem. These cells that are programmed to yield the adaxial portion of the lateral organ, are predicted to in turn hava YAB and KAN genes repressed (Bowman et al., 2002). In this respect, semidominant gain-of-function mutants of PHB and PHV genes cause adaxialization of lateral organs (McConnell and Barton, 1998; McConnell et al., 2001). PHB, PHV, and REV have similar expression patterns. They are expressed in the SAM initiating lateral organs and later become adaxially restricted as the primordium emerges (McConnell et al., 2001; Otsuga et al., 2001; Prigge et al., 2005). Finally, phenotypes of the loss-of-function rev mutants could be interpreted as having a partial loss of adaxial identity (Talbert et al., 1995; Otsuga et al., 2001).
Besides the PHAB function in polarity, it is also interesting to note that a phb phv cna (corona, another member of the HDZIP III gene family) triple mutant has a very similar phenotype to those of clv mutants with a distinct increase in organ number in each whorl. This would suggest that HD-ZIP III genes and the CLV pathway regulate meristem function in a similar manner. The possible interrelation of these modules could contribute to homeostasis between stem cell maintenance and organ formation (Prigge et al., 2005).
NUB and JAG are similar genes which encode C2H2-zinc finger transcription factors that are proposed to play redundant functions in proliferation and differentiation of adaxial cells, particularly during anther and carpel development (Dinneny et al., 2004; Ohno et al., 2004; Dinneny et al., 2006; Xu et al., 2008). They specifically work together in determining the number of cell layers formed in floral organs, and like the PHAB family, they are not cell-fate genes. Hypothetically, JAG suppresses the premature differentiation of tissues by slowing down the cessation of cell division in distal regions of organs until it finally arrests after normal morphogenesis has occurred (Dinneny et al., 2004).
AS1 and AS2 have redundant functions in the establishment of adaxial identity (Ori et al., 2000; Sun et al., 2000; Semiarti et al., 2001). AS1 encodes a MYB-domain transcription factor, and AS2 is a member of the LBD gene family (Serrano-Cartagena et al., 1999; Byrne et al., 2000; Semiarti et al., 2001; Sun et al., 2002). AS1 protein is expressed in organ initials and physically interacts with AS2 to inhibit KNOX gene expression, thus guiding primordia toward differentiation (Figure 15; Ori et al., 2000; Byrne et al., 2002; Xu et al., 2003; Guo et al., 2008).
Other reviews on polarity determination in embryos and in leaves are found in other chapters in this series: “Embryogenesis: pattern formation from a single cell” (Berleth and Chatfield, 2002) and “Leaf development” (Tsukaya, 2002).
3.4.4 Sepals and petals
Sepals and petals constitute the sterile perianth in the first and second flower whorls, respectively. The sepal whorl or calyx protects the developing floral bud and in some plants, but not in Arabidopsis, it may be involved in fruit development (He et al., 2004). The petal whorl or corolla is generally thought to be important for attracting pollinators (Krizek and Fletcher, 2005), but in an autogamous plant such as Arabidopsis, the corolla is generally not showy.
According to the ABC model, sepal identity specification depends on the activity of both A and SEP genes (see section 3.3; Coen and Meyerowitz, 1991; Pelaz et al., 2000), and petal identity specification depends on the overlapping activities of A, B and SEP genes (see section 3.3; Coen and Meyerowitz, 1991; Pelaz et al., 2000). Also, it has been shown that sepal and petal identity specification depends, at least in part, on the correct downregulation of AG expression in the second whorl (see below).
Several molecular components known to influence development of sepals, influence petals too. But knowledge is still limited especially of sepal developmental gene networks. However, a basic GRN for petal development can be constructed based on available data (Figure 16). As stated earlier, organ identity determination, boundary establishment, and expression of polarity determinants are common features needed for the correct development of all the flower organs (Figure 15).There are several pieces of evidence that suggest that genes involved in these processes might be acting at the same time (for example, expression profiles and in situ hybridization assays), at least momentarily during flower development. However, we still do not understand fully how such functional modules interact with each other.
As it was said before sepal and petal boundary and organ number establishment are controlled by the CUC and FFO2 genes (see Figure 16 and section 3.4.2; Aida et al., 1997; Levin et al., 1998). CUC gene expression is regulated by the miR164c (encoded by EEP1) in an organ specific manner (Laufs et al., 2004; Baker et al., 2005).
Several genes are involved in establishing and maintaining the sepal and/or petal domain and, in a way, determining the boundaries between the organs. One of the main activities of these genes is to exclude AG expression from the first and second whorl. As stated in section 3.3, AG is repressed by RBE, LUG, SEU, ROXY1, AP2, BLR, ANT and SAP (for more information about each gene, see Table S1 () ; Figures 15 and 16).
Briefly, RBE is mainly involved in boundary and organ number determination of both sepals (non-autonomously) and petals, and in AG exclusion from the second whorl at early stages of flower organ development. But it is also important during late petal development as mutants may form filamentous organs in the second whorl. RBE expression is controlled by both PTL and UFO (Takeda et al., 2004; Krizek et al., 2006). PTL is a trihelix transcription factor that is expressed at early stages in four zones between the initiating sepal primordia and in lateral regions of stamen primordial. Later on, PTL expression can be detected at the margins of expanding sepals, petals, and stamens (Brewer et al., 2004). Thus PTL may delimit the AG expression region indirectly by activating RBE expression (Irish, 2008), and it may also be controlling lateral outgrowth of mature sepals, petals and stamens defining their final shape and orientation (Griffith et al, 1999; Brewer et al., 2004).
UFO is also an important regulator of petal development. Its action toward RBE may be indirect, as it may be degrading (as part of an SCF E3 ubiquitin ligase complex) an unknown repressor of RBE (Irish, 2008). But UFO is an important network link between the AG inactivation pathway and the B gene identity determination pathway, because UFO interacts with LFY to activate AP3 expression (See section 3.3; Lee et al., 1997; Samach et al., 1999; Chae et al., 2008). Importantly, UFO expression is also required for normal petal blade outgrowth after petal identity has been established (Laufs et al., 2003), as well as for determination of sepal shape and number in the first whorl (Levin and Meyerowitz, 1995; Samach et al., 1999).
SEU and LUG also repress AG expression in the first and second whorls by forming a protein complex with AP1 and SEP3 (see section 3.3; Sridhar et al., 2004; Sridhar et al., 2006). But these genes are also part of the adaxial/abaxial polarity establishment pathway in the petal GRN, as they are required for normal PHB and FIL expression (Figure 15). SEU and LUG participate in petal shape regulation by controlling blade cell number and petal vasculature development in an AG independent manner (Franks et al, 2006). Finally, SEU is also involved in auxin response pathways by directly interacting with ETT, and influencing the final shape, number and phyllotaxy of petals (Pfluger and Zambryski, 2004).
As part of the regulatory network that represses AG expression, AP2 is itself negatively regulated by miR172 when second whorl boundaries are determined (Chen, 2004; Zhao et al., 2007). Besides being a negative regulator of AG, ANT also affects organ number and morphology in the first three whorls (Elliott et al., 1996; Klucher et al., 1996). SAP, another regulator of the morphology of all organs, but mostly of petals, is unexpectedly more important in later flowers (Byzova et al., 1999).
Another important indirect repressor of AG is ROXY1. As a glutaredoxin, ROXY1 seems to be a postranslational modifier of AP2, LUG, UFO and RBE giving them the specificity to repress AG in the second whorl (Xing et al., 2005; Irish, 2008). ROXY1 is also important for repressing PAN expression and for activating other TGA factors at different stages of petal development (Li et al., 2009).
Genes that usually work in the establishment of lateral organ polarity (see section 3.4.3) are also important in determining the polarity of sepals and petals, e.g. PHB, JAG, FIL, YAB3, KAN, AS1 and AS2 (Figure 16). Experimental data suggest that AS1, AS2 and JAG are negative regulators of CUC1/2 and PLT(Xu et al., 2008). This links the expression of these genes with those important for boundary determination in the GRN of both sepals and petals. PHB and FIL expression are also part of the network and are regulated by SEU and LUG (Franks et al., 2006). Lateralaxis dependent development is determined by the PRESSED FLOWER (PRS) homeobox gene (Matsumoto and Okada, 2001). As with some other genes involved, its position in the GRN is unknown, but by analyzing the mutant phenotypes, it becomes clear that the same regulatory modules that underlie polarity determination are involved in organ shape regulation.
In Arabidopsis, as in other plants, several mutants featuring a foliose-sepal-syndrome (FSS) (leaf-like sepals) have been isolated. Ectopic expression of the MADS-box genes AGL24, SVP, and ZMM19 (from Zea mays), belonging to the STMADS11-clade (according to Theissen et al., 2000), result in FSS (He et al., 2004). The main feature of these leaf-like sepals is that they are large and have leaf-like stellate trichomes on their outer surface. One of the characteristics of ap1 mutant plants is that they also have large or foliose sepals. Thus, it has been proposed that, in addition to their roles in floral transition and/or organ determination, AP1, SVP, and AGL24 may also have a role regulating sepal size (He et al., 2004). But how they interact among themselves or with other sepal specific genes is still unknown.
Final sepal and/or petal morphology is also determined by FRL1 (Hase et al., 2000; Hase et al., 2005), TSO1 (Hauser et al., 1998), the AP3/PI regulated genes GNC, GNL, At4g30270, HXK1 (Mara and Irish, 2008), and NAP (Sablowski and Meyerowitz, 1998). Except for FRL1, which is involved in endoreduplication control, and TSO1, which is likely involved in chromatin remodelling, the position of these genes in the petal GRN has already been established (see Figure 16).
Using microarray approaches Wellmer et al. (2004), compared gene expression levels within different floral homeotic mutants (see section 3.3.1). Their first study of stage 2 flowers identified only 13 genes as being sepal-specific and only 18 genes expressed exclusively (or predominantly) in petals. However, a more recent study of flowers at stage 3, when sepal primordia have just formed, revealed that 199 genes are upregulated and 161 genes downregulated (Figures 3–4; Wellmer et al., 2006). One speculation is that sepals are relatively simple organs and not many specific genes are involved in their development. But more detailed studies are still required. Results also suggest that genes regulating sepal and petal development may have been recruited from leaf developmental pathways, and, hence, are not specific for the development of these organs.
Petals have been proposed as an excellent model system in which to study development because they have a simple organization and are not essential for survival or reproduction (Irish, 2008). Although much progress has been made, much work is still needed for an integrated and dynamical understanding of petal development.
Six stamens occupy the third whorl in the Arabidopsis flower. Stamen specification depends on the overlapping activities of B, C and SEP MADS-box genes (Coen and Meyerowitz, 1991; Pelaz et al., 2000). A complex network of gene regulatory modules is simultaneously activated in young stamen primordia, and these are also important for organ morphogenesis (Figure 15). These modules include those that regulate adaxial-abaxial primordium polarity (also affecting other vegetative and reproductive lateral organs) including genes from the PHAB (PHB, PHV, and REV), KANADI (KAN1-4), and YABBY (FIUYAB1, YAB2, and YAB3) families. At later stages of stamen development, genes involved in sporogenesis such as SPL and BAM1/2, and in anther development, such as JAG and NUB, are activated (see Figures 15 and 17 for regulatory modules and genes; Scott et al., 2004; Ma, 2005; Feng and Dickinson, 2007).
Among the most striking stamen development mutants is fil (also called antherless and undeveloped anther) which bears normal filaments with neither anthers nor pollen. The FIL gene is YABBY-like and the fil phenotype suggests that the developmental programs of the filament and anther are controlled by independent regulatory modules (Sanders et al., 1999).
As mentioned in section 3.3.1, SPL/NZZ is essential for male and female reproductive development and is probably the first reproductive gene to be activated in the anther or, at least, it is the only gene that remains active during most of early anther development. This transcription factor gene is expressed during micro- and megasporogenesis. AG directly induces SPL but AG is not necessary for maintaining its expression (Ito et al., 2004). spl mutants are not able to produce microsporogenous cells or tapetal tissue, and show several alterations in anther wall and nucellus development (Schiefthaler et al., 1999; Yang et al., 1999). Interestingly, BAM1 and BAM2, which participate in the first cell division of the archesporial cells and the subsequent periclinal divisions to produce the somatic cell layers, are proposed to form a regulatory loop with SPL (Figure 17; Hord et al., 2006; Feng and Dickinson, 2007). Since SPL maintains the sporogenous activity in the microsporogenous cells, and BAM1/2 maintain somatic differentiation, bam 1 bam2 anthers have cells interior to the epidermis with characteristics of pollen mother cells (Hord et al., 2006).
Although SPL is one of the genes expressed the earliest in stamen development, it is not the only one. Ectopic expression of SPL in all the whorls of an ag mutant, results in the formation of microsporangia only in the lateral parts of the staminoid ‘petals’, suggesting that microsporangial localization is established independently of AG, and that there is at least one other SPL inducer that is expressed in the second whorl, and not in other whorls (Ito et al., 2004; Feng and Dickinson, 2007). Two other genes, JAG and NUB play a crucial role in the formation of the four-locular anther architecture, independent of SPL induction, jag nub double mutants do not have a proper microsporangium. Instead, they form a finger-like structure that expresses SPL in its tips (Dinneny et al., 2006; Feng and Dickinson, 2007).
The correct number of microsporangial initials and the subsequent production of the tapetal cell and middle cell layer identities are properties specified by a putative LRR receptor kinase, EXCESS MICROSPOROCYTES1 (EMS1)/EXTRA SPOROGENOUS CELLS (EXS) (Canales et al., 2002). Until recently, the ligand for EMS1 was unknown, though it was hypothesized that it could be involved in the same signaling pathway as the TAPETAL DETERMINANT (TPD1) gene. Both tpd1 and tpd1 ems1 mutants are similar to the single ems1 mutant with arrested meiotic cytokinesis and degenerated microsporocytes (Yang et al., 2003). TPD1, is a small putatively secreted protein that interacts with EMS1 and induces its phosphorylation suggesting that TPDI is the ligand of the EMS1 receptor that signals cell fate determination during sexual cell morphogenesis (Jia et al., 2008).
ROXY1 and ROXY2 redundantly regulate anther development in Arabidopsis (Xing and Zachgo, 2008). Lateral and medial stamens of roxy1 mutants might be fused and the former are sometimes missing (Xing et al., 2005). In these mutants, the adaxial anther lobes are affected in sporogenous cell formation during early differentiation steps, abaxial lobes develop normally but pollen mother cells degenerate, while the tapetum overgrows and occupies most of the locule space. Eventually, the tapetum degenerates too. R0XY1 and ROXY2 function downstream of SPL and upstream of DYSFUNCTIONAL TAPETUM1 (DVT1). As with other glutaredoxins, they may need an interaction with glutathione to catalyze biosynthetic reactions, suggesting that they may have a role in redox regulation and/or plant stress defense mechanisms involved in the control of male gametogenesis (Xing and Zachgo, 2008).
After tapetal cells are specified, a range of genes are essential for subsequent development. DYT1 encodes a putative bHLH transcription factor which functions downstream of SPL and EMS1. However DYT1 is not able to complement the spl or ems1 mutant phenotypes when it is overexpressed, indicating that it is required but not sufficient for normal tapetum development, dyt1 exhibits abnormal anther morphology with largely vacuolated tapetal cells that eventually collapse. Several tapetum-expressed genes, such as MALE STERILE 1 (MS1) and ABORTED MICROSPORES (AMS) are upregulated by DYT1 (Zhang et al., 2006). In ms1 mutants for example, tapetal cell abnormalities can be seen and pollen development is arrested just after microspores are released from the tetrads (Bowman, 1994; Wilson et al., 2001; Yang et al., 2007a). Other genes that participate in tapetum development include RECEPTOR-LIKE PROTEIN KINASE2 (RPK2), FAT TAPETUM and GUS-NEGATIVE1 and 2 (GNE1, GNE2). RPK2 regulates tapetal function and middle layer differentiation (Mizuno et al., 2007). FAT TAPETUM, when mutated, has a middle layer that fails to collapse after meiosis and shows tapetal-like behavior (Sanders et al., 1999; Ma, 2005). In gne1 and gne2 mutants the sporogenous cells enter meiosis, but cytokinesis is frequently arrested. The few highly aberrant tetrads formed degenerate early and microsporangia of mature anthers end up empty (Sorensen et al., 2002).
Several mutants affecting pollen development have been described: pollenless3; three division mutant (tdm1); ms5, ms3 and ms15; determinate infertile1 (dif1); switch1 (swi1); defectivepollen 1, and 6492 among others (Bhatt et al., 1999; Sanders et al., 1999; Sorensen 2002). Meiotic cells in pollenless3 anthers undergo a third division without DNA replication generating some cells with unbalanced chromosome numbers (Sanders et al., 1999) or “tetrads” with more than four microspores, dif1 and swi1 mutants have micro- and megaspores of uneven sizes because the encoded proteins are essential for sister chromatid cohesion in male and female meiosis and so mutants are totally infertile (Bhatt et al., 1999; Parisi et al., 1999; Mercier et al., 2001; Ma, 2005). Finally, other pollen mutants exhibit abnormal callose deposition (ms32, ms31, ms37, 7219, and 7593).
There are late-developmental anther mutants that affect anther dehiscence. In non-dehiscence1 mutant plants, anthers contain apparently wild-type pollen but do not dehisce. It has been hypothesized that a cell death suppression program, which is normally responsible for dehiscence, might be inactive in this mutant (Sanders et al., 1999). ms35 is also affected in anther dehiscence, because endothecial cells fail to develop the lignified secondary walls that after desiccation shrink differentially leading to the retraction of the anther wall and full opening of the stomium (Dawson et al., 1999; Scott et al., 2004). MS35, now MYB26 (Steiner-Lange, 2003), is expressed during early anther development and may be a regulator of NAC SECONDARY WALL-PROMOTING FACTOR 1 and 2 (NST1, NST2), which have also been linked to secondary thickening in the anther endothecium (Yang et al., 2007b). In delayed-dehiscence mutants (dd1, dd2, dd3, dd4, dd5) anther dehiscence and pollen release occurs after the stigma is no longer receptive preventing successful pollination, but stamens look wild-type and pollen is viable (Goldberg et al., 1993). On the contrary, in defective-pollenl, 2, and 3, anthers are able to dehisce, but the pollen is aberrant and unviable.
Recent publications have established that gibberellic acid (GA), jasmonic acid (JA), and auxins are involved during stamen development (Figure 18; Fleet and Sun, 2005; Nagpal et al., 2005; Wu et al., 2006; Cecchetti et al., 2008). The GA-deficient mutant, ga1–3, produces an abortive anther where microsporogenesis is arrested prior to pollen mitosis (Cheng et al., 2004). Mutations in two GA receptors, GA-INSENSITIVE DWARF1a and b (AtGID1a, b), affect the elongation of stamens, suggesting that these receptors have specific roles during stamen development (Iuchi et al., 2007). GA induces the degradation of the DELLA protein REPRESSOR OF GA1–3 (RGA) upon ubiquitination. Microarray analysis shows that 38% of the RGA downregulated genes are expressed in the male gametophyte at various stages of microsporogenesis (Hou et al., 2008).
Temporal coordination of the elongation of filaments, pollen maturation, and dehiscence of anthers is important for efficient fertilization. The expression overlap of RGA-regulated genes and jasmonate-responsive genes during stamen development suggest a crosstalk between GA and JA signaling pathways in these processes (Hou et al., 2008).
JA has been shown to be involved in at least three androecial developmental pathways: filament elongation, anther dehiscence and pollen production (Mandaokar et al., 2006). Different male sterile mutants have been found to be JA biosynthetic mutants (McConn and Browse, 1996; Sanders et al., 2000) including: the triple fed mutant (fad3–2 fad7–2 fad8), which lacks the fatty acid precursors of JA; defective in anther dehiscence 1 (dad1), which encodes a phospholipase A1 that catalyzes the initial step of JA biosynthesis; and dd1, a member of the 12-OXOPHYTODIENOATE REDUCTASE (OPR3) gene family (Stintzi and Browse, 2000; Ishiguro et al., 2001). OPR3/DD1 is expressed in the stomium and in the septum cells of the anther that are involved in pollen release. All these mutant phenotypes can be rescued by exogenous application of JA, suggesting that this hormone plays an important role in controlling the timing of anther dehiscence. Interestingly, DAD1 is a direct target of AG (Ito et al., 2007).
Similarly, the coronatine insensitive 1 (coi1; JA receptor) mutant is defective in both pollen development and anther dehiscence. Stamens of coi1 flowers have shorter filaments than those of wild-type flowers and anthers are indehiscent containing pollen grains with conspicuous vacuoles (Feys et al., 1994; Xie et al., 1998).
Three related polygalacturonases, enzymes involved in pectin degradation that promotes cell separation, are also involved in JA-regulated anther dehiscence. ARABIDOPSIS DEHISCENCE ZONE POLYGALACTURONASE 1 (ADPGI) and 2 (ADPG2), and QUARTET2 (QRT2) gene expression are distinctly regulated by JA (Ogawa et al., 2009).
To determine the jasmonate-regulated stamen-specific transcriptome the expression profiles of JA-treated and untreated opr3 mutants were compared (Mandaokar et al., 2006). It was found that 821 genes were induced (70% of them expressed in the sporophytic tissue) and 480 genes were repressed by JA and 13 transcription factors were identified that could be important for stamen maturation pathway(s). Of these, MYB21, MYB24, and MYB28 are JA-responsive genes (Mandaokar et al., 2006). myb21 mutants have short filaments, are late to dehisce and have reduced fertility. Though myb24 mutants look like wild type, myb21 myb24 double mutants have a more severe phenotype than myb21, suggesting that these two genes might be redundantly involved in important aspects of JA-dependent stamen development. MYB28 is involved in amino acid metabolism and it is downregulated by both JA and RGA. This study also uncovered several other biochemical pathways that could be important during stamen and pollen maturation. Other results indicate that JA coordinates pollen maturation, anther dehiscence, and flower opening (Ishiguro et al., 2001). Auxins have also been proved to participate in these processes arf6 arf8 double mutants are defective in anther dehiscence probably because they produce too little JA. Accordingly, this phenotype can be rescued by application of JA (Nagpal et al., 2005). However, auxins trigger filament elongation and prevent premature anther dehiscence and pollen maturation at earlier stages of stamen development. While JA production peaks at stages 11–12 of flower development (see Figure 6 and 18; Nagpal et al., 2005) auxin receptors (TIR1 and AFBs) are already expressed at the end of meiosis. Mutants in these genes cause the release of mature pollen grains before filaments elongate. At later stages, the amount of JA decreases allowing these processes to continue (Figure 18; Cecchetti et al., 2008).
Additional stamen or pollen microarray analyses have been performed recently. For example, a clear difference was found between the genes that are expressed in the sporophyte and in pollen with 39% of the expressed genes being pollen specific (Honys and Twell, 2003; Pina et al., 2005). The global gene expression profiles of wild-type reproductive axes have been compared to those of the floral mutants ap3, spl/nzz, and ms1 in order to study gene expression during stamen development and microspore formation (Alves-Ferreira et al., 2007). The data suggest that different interconnected regulatory modules may control specific stages of anther and microspore development (for further details see: Amagai et al., 2003; Cnudde et al., 2003; Honys and Twell, 2003; Zik and Irish, 2003a; Wellmer et al., 2004; Pina et al., 2005; Alves-Ferreira et al., 2007; Verelst et al., 2007).
3.4.6 Carpels and ovules
Carpels are specified by the C gene AG, and the SHP1, SHP2, and STK genes (in an AG independent manner) together with the SEP genes (Bowman et al., 1989; Coen and Meyerowitz, 1991; Favaro et al., 2003; Pelaz et al., 2000; Pinyopich et al., 2003). They arise in the center of the flower meristem and when carpels are fully developed the floral meristematic cells are completely consumed. Carpels are the most complex structures within flowers and a GRN underlies their development (Figure 19; Table S1 ()), Comprehensive reviews on carpel and fruit development can be found in Bowman et al., (1999), Ferrándiz et al., (1999), Balanzá et al., (2006) and in Roeder and Yanofsky (2006) in this book.
Little is known about the molecular genetics of nectary development. It is clear that nectaries are ABC-independent because ap2-2 pi-1 ag-1 triple mutant flowers develop nectaries, although in these mutants nectaries are clearly smaller. However, ABC genes may play a role in nectary patterning as pi-1 ag-1 and ap3-3 ag-3 double mutants lack them (Baum et al., 2001). Also, single mutant lfy and ufo plants show reduced nectary formation (Lee et al., 2005a).
Several genes have been found to be expressed in the nectaries (e.g., SHP1, ALC, SPL, MS35 and XAL1), but no detectable defect is observed in their respective mutants (Figure 5G; Schiefthaler et al., 1999; Roeder and Yanofsky, 2006; Yang et al., 2007b; Tapia-López et al., 2008). The only gene that has been clearly related to nectary development is CRC, which is also important for gynoecial development (Alvarez and Smyth, 1999; Bowman and Smyth, 1999). The regulation of CRC in the nectaries seems to be independent of its expression in the gynoecium. Expression of this gene is already detectable at stage 6 of flower development in the area where the nectaries will be formed. Thus, CRC may be important for the early specification of nectary cells (Bowman and Smyth, 1999). CRC may also be necessary for maturation or maintenance of the nectaries, because it is expressed at high levels when they develop (Bowman and Smyth, 1999) and crc mutants have defects in nectary development. But CRC is not sufficient for nectary development, because its ectopic expression does not yield ectopic nectaries (Lee et al., 2005b). Bioinformatic and functional molecular genetic approaches have been taken to discover components of the regulatory network in which CRC participates during nectary and carpel development. A combination of floral homeotic gene activities acting redundantly with each other, involving AP3, PI and, AG and in combination with SEP proteins, activate CRC in both organs (Lee et al, 2005a). Interestingly in another study, CRC was also found to be a direct target gene of AG (Gómez-Mena et al, 2005) and to be indirectly regulated by LFY and UFO (Lee et al., 2005a). A model has been proposed in which LFY and UFO activate downstream MADS-box genes which could be working in conjunction with region-specific factors to activate CRC during nectary and carpel development (Lee et al., 2005a).
Evolutionary studies have indicated that the CRC gene may have been recruited as a regulator of nectary development in the core eudicot plant lineage, but its ancestral function could have been related to carpel development (Lee et al., 2005b).
4. THEORETICAL MODELS: INTEGRATIVE AND DYNAMIC TOOLS FOR UNDERSTANDING FLOWER DEVELOPMENT
As shown throughout this chapter, morphogenetic patterns underlying flower development arise from complex, often non-additive, interactions among several molecular and other kinds of components (e.g., cells) and factors (e.g., morphogen gradients, physical and geometrical constraints) at different levels of organization. Dynamical models can be used to study the concerted action of many elements at different spatio-temporal scales and levels of organization; an approach which is becoming both necessary and possible for understanding how morphogenetic patterns emerge and are maintained during development in general, and in particular, in flower development (for reviews Alvarez-Buylla et al., 2007; Grieneisen and Schieres, 2009). At the level of GRN, mathematical and computational models provide useful tools for integrating and interpreting molecular genetic information, or for detecting gaps and contradictions in the evidence for particular functional regulatory modules. At other levels, two or three-dimensional morphogenetic models of coupled GRNs within cells or among cells are useful for understanding spatiotemporal cell patterning in individual organs and overall plant architecture; and this enables novel insights into the mechanisms underlying developmental processes to be made. Such morphogenetic models are also a way of posing informed non-trivial predictions, testing hypotheses, uncovering potentially generic mechanisms underlying conserved features, and performing in silico investigations that guide novel experiments in biological development.
As the amount of experimental evidence increases and novel theoretical approaches and techniques develop, there continue to arise experimentally-grounded models of development and theoretical tools useful in posing predictions amenable to further experimental testing. These advances contribute to discussions of central issues in developmental and evolutionary biology (e.g., Kauffman, 1969; Berg et al., 2004; Milo et al., 2004; Wagner, 2005; Alvarez-Buylla et al., 2007; Balleza et al., 2008). In Arabidopsis, early flower development has been studied using dynamic gene regulatory network (GRN) models. Such models have helped capture the logic of gene regulation, mostly at the transcriptional level, during cell-type specification in various systems (e.g. von Dassow et al., 2000; Espinosa-Soto et al., 2004; Huang and lngber, 2006; Li et al., 2006; Benítez et al., 2008). In this section we focus on this modeling approach and present some of the main results derived from network modeling in flower development.
4.1 Gene Regulatory Network Models
In this section we review some central notions in GRN theory and the main assumptions that are made and present some of the main results derived from network modeling in flower development. GRN models are composed of nodes, which stand for genes or proteins, and edges or connections, which represent the interactions among nodes (arrows for upregulation and bars for downregulation; for an example see Figure 20). Genes in the GRN model may take different activation states, depending on the activation states of their inputs. Given the architecture of the network and the sign of the interactions, it is possible to define a set of rules or kinetic functions that govern the GRN dynamics, that is, the way activation states of the genes change over time. These rules or kinetic functions may be defined and studied in the context of different mathematical frameworks, some of which have been thoroughly reviewed elsewhere (Gibson and Mjolsness, 2004; Alvarez-Buylla et al., 2007). In experimentallybased GRNs, the dynamic rules may be obtained from reported molecular genetics data as well as from functional genomics datasets.
The kinetic functions of gene activation depend on the states of the input nodes and are multivariate. These may be modeled with discrete or continuous functions. In the former, Boolean functions that allow only “0” (OFF; not expressed) or “1” (ON; expressed) values for the nodes have been successfully used to recover the key qualitative aspects of GRNs (e.g., Albert and Othmer, 2003; Espinosa-Soto et al., 2004). In Boolean networks, parameters of specific kinetic functions are not required. It is appropriate to assume that the GRN nodes are Boolean variables given that: (1) transcriptional regulation may be discrete and take place in the form of pulses, rather being continuous (Ross et al. 1994, Fiering et al., 2000, Ozbudak et al., 2002); (2) the experimental data at hand can be readily formalized as logical rules (see detailed discussions in Albert and Othmer, 2003; Espinosa-Soto et al., 2004; Chaos et al., 2006), while there are no or very few available data on parameters required to postulate continuous functions; and (3) in complex GRNs with many components interacting in non-linear manners, the overall topology of the GRN and the form of the logical rules of gene interaction, rather than the details of the kinetic functions, are what determine the qualitative network dynamics.
Independently of the mathematical formalism used, dynamical analyses of GRNs mostly focus on finding the steady gene activation profiles, that is, the configurations of the network that, once reached, remain in that configuration. These configurations are called attractors. The GRN model may be initialized on a particular gene-activation configuration known as an initial condition and then the elements of the GRN change their activation state according to the dynamic rules until they reach an attractor. Kauffman (1969) proposed that Boolean GRN attractors correspond to the activation profiles typical of different cell types and therefore that exploring the GRN architecture and dynamics is fundamental to understanding cell-type determination processes. This idea has now been verified experimentally and explored further (e.g. Albert and Othmer, 2003; Huang and lngber, 2006; Alvarez-Buylla et al., 2007).
Another helpful notion in GRN dynamical studies is that of basins of attraction. Given the dynamic rules of the network, the set of initial conditions that lead to each of the attractors is known as its basin of attraction. As we discuss below, the concepts related to a GRN - attractor, initial condition and basin of attraction - may be useful in addressing some pertinent aspects of flower development.
4.1.2 Functional Modules in Flower Development
The functional data on genes involved in flower development reviewed in this Chapter suggest that several regulatory modules act at different stages and in different structures (Figures 9, 15–17 and 19). We define a regulatory module as a set of interacting genes that have more interactions among themselves than with other genes. These modules are semi-autonomous, meaning that their dynamic outcomes are fairly independent of other modules. In Figure 15 we have presented the best-studied modules associated with flower development. The approach described here for the functional module that includes the ABC genes could in principle be used for all of these modules as sufficient nodes have been identified and their regulatory interactions characterized. Eventually, models of coupled GRN that consider several such models together, both within and among cells, will be possible. For now, we have focused in just one such regulatory module.
In previous studies, we have proposed and analyzed the regulatory module, which includes the ABC genes as well as other components, that is sufficient to regulate the partitioning of the inflorescence and floral meristems into subregions of primordial cells. In the case of the flower meristem, each one of the four subregions is composed of the primordial cells that eventually give rise to each of the four types of floral organs: sepals, petals, stamens and carpels.
We use this functional module as a working example of the protocol that has been used in order to assemble an experimentally grounded gene regulatory network (GRN) model corresponding to a functional module. Then we demonstrate how once such a GRN model is postulated, it is possible to follow its dynamics, and explore how the concerted action of multiple interconnected molecular components eventually lead to stable gene expression profiles that may be compared to those characterizing different cell types. Then we delineate some theoretical approaches put forward to model coupled GRN dynamics that may underlie pattern formation and morphogenesis during the early stages of flower development, when the floral meristem is partitioned into four concentric rings of primordial cells. Finally, we review other modeling approaches that are useful to study signal transduction pathways.
4.2 Arabidopsis Flower Organ Specification GRN (FOS-GRN)
Soon after flowering is induced, the flower meristem is partitioned into four concentric regions of primordial cells from which floral organs will later form. During the last decade, an experimentallygrounded GRN model for flower organ specification (FOS-GRN) has been built and investigated (Figure 20; Mendoza and Alvarez-Buylla 1998; Espinosa-Soto et al., 2004; Chaos et al., 2006; Alvarez-Buylla et al., 2008). This model incorporates the intricate regulatory interactions among ABC genes themselves and among ABC and non-ABC genes that are key to this process. This functional module includes: some key regulators underlying the transition from IM to FM (FT, TFL, EMF, LFY, AP1, FUL); the ABCs and some of their interacting genes (AP1, AP3, PI, AP2, AG, SEP); some genes that link floral organ specification to other modules regulating primordia formation and homeostasis (AG, CLF and WUS); and some regulators of organ boundaries (UFO and LUG; Figures 9, 15 and 20).
The main result obtained from analyzing this GRN is that the postulated network converges to only ten attractors—even though it can be initialized in more than 130,000 different configurations of gene activation. Furthermore, the attractors—the stable configurations recovered—match gene activation profiles typical of the four inflorescence meristem regions (i.e., a region lacking WUS and UFO, two regions with either one of these two genes turned on, and a fourth region with both genes turned on; see Espinosa-Soto et al., 2004), as well as those of primordial sepal, petal, stamen and carpel cells (Figure 21 ). This shows that the FOS-GRN is sufficient to recover the gene activation profiles required to specify primordial cells during the first stages of flower organ development. Therefore the GRN itself constitutes a functional module that robustly leads to the gene configurations that characterize different regions of inflorescence and flower meristems during early flower development; and this independently of the activation states of additional genes that are connected to this elucidated regulatory module. Furthermore, various robustness analyses have been performed showing that the recovered attractors are also robust in response to permanent alterations in the logical functions of gene interactions and the inclusion of gene duplications. Therefore, these results (Espinosa-Soto, et al., 2004; Chaos et al., 2006) suggest that FOS dynamically and robustly emerges from complex networks of molecular components, rather than from a series of linear or hierarchical gene interactions or from the action of particular genes. The FOS-GRN model not only recovers the ABC gene combinations that are necessary for FOS, but it also provides a dynamic explanation for the formation of such gene combinations, and postulates a set of gene interactions with the ABC genes, that are also sufficient for FOS. The functions and interactions of the genes included are reviewed earlier in this chapter.
The FOS-GRN was validated by using this model to simulate the effect of loss-of-function mutations or overexpression, and comparing the results recovered from the model with the gene activation profiles determined experimentally in mutant or overexpressor lines. The mutants were simulated by fixing the state of the gene to 0 for loss of function, and to 1 for gain of function or overexpression (Figure 20; Table 1 and Table S2 ()). In all cases tested, the simulated and empirically-reported profiles matched (Espinosa-Soto et al., 2004).
In addition, this GRN model has enabled investigations to be made into the sufficiency and necessity of particular gene regulatory interactions, which have led to novel predictions. For example, these analyses predicted that AG upregulated itself (EspinosaSoto el al., 2004), which seemed somewhat counterintuitive at the time, but which was then verified by independent experiments (Gómez-Mena et al., 2005). Also, computer simulations of the FOS-GRN that show that its attractors are robust to different types of perturbation and to duplications (Espinosa-Soto et al., 2004; Chaos et al., 2006) can account for the overall conservation of the flower structure throughout angiosperm (particularly eudicot) evolution (Rudall, 2007; Whipple et al., 2004; Adam et al., 2007).
Since the FOS-GRN model was based on thorough molecular data and is one of the few well-characterized regulatory modules, it has been used as a “model GRN” for further methodological, theoretical and conceptual developments in GRN and systems biology research (Table 2). The main conclusions obtained from the first versions of this GRN have been confirmed. New data regarding FOS are continuously being generated (novel data are also summarized in Table 1) and the FOS-GRN constitutes a basic theoretical framework in which to integrate it alongside previous data. Here, we have updated the FOS-GRN taking these novel data into account and conclude that the basic module originally put forward (Espinosa-Soto et al., 2004; Chaos et al., 2006) is robust to the addition of these newly discovered interactions. We consider, for instance, that EMF1 downregulates AG (Calonje et al., 2008), and AP3/PI downregulate AP1 (Sundström et al., 2006), so the postulated module seems to be robust to the addition of intermediary components or previously missing interactions.
Simulations of the updated FOS-GRN have been performed with the new software, ATALIA ( http://www.ecologia.unam.mx/~achaos/Atalia/atalia.htm) developed in the Alvarez-Buylla laboratory by A. Chaos-Cador. This tool can be used to readily update this and other GRN models and explore their dynamics. We illustrate the use of this software with a visualization of the attractors' basins (Figure 22) and a simulation of the updated wildtype and certain mutant FOS-GRN dynamics (Figure 23).
In the simulated FOS-GRN, genes can take only two activation states: 0 for no expression and 1 for expression. Hence, by using combinations of Os and 1 s, we can describe all the possible initial conditions of the GRN. Figure 22 presents the so-called fan diagrams that show all the GRN configurations leading to each of the attractors. Knowing the relative sizes of the basins of attraction of each steady state is the key to exploring the robustness of each attractor in the face of perturbations.
ATALIA can also calculate the attractor that every possible initial condition will eventually reach and show this information in a tapestry of destinies. In such tapestries, each possible configuration of the GRN is represented by a square in a lattice and is colored according to the attractor it reaches. Moreover, ATALIA can draw a tapestry that represents the difference between the original wild-type tapestry and a mutant one (Figure 23). For example, if we want to know whether an ap2 mutation has a more or less drastic effect in terms of the GRN dynamics than a pi mutation, we can analyze the tapestries of ap2 and pi shown in Figure 23 and conclude that ap2 mutation has stronger dynamic effects than pi given the GRN postulated up to now. Given the complexity of the network involved, such predictions are impossible to make without a tool like ATALIA. As the regulatory interactions in other modules that participate in flower development are gradually uncovered, for each one the experimental data can be exhaustively mined and formalized in the form of a GRN topology and logical rules governing its components' interactions. ATALIA can then be used to explore their dynamics, validate the proposed GRN models by simulating experimental reports of mutants or overexpressing lines, and to postulate novel interactions. Eventually, two or more functional modules may be interconnected via common components to postulate GRN aggregates. Such an approach will be useful in beginning to uncover the types of microtopological trait that characterize the nodes connecting different functional modules, for example.
We have illustrated the potential of using dynamic GRN models to understand cell differentiation using a relatively small and well-characterized module. Approaches used for small regulatory modules that are well-characterized in terms of molecular genetics, should feedback from functional genomic efforts that span the dynamics of a larger number of genes or proteins under diverse conditions and developmental stages or tissues.
4.2. Temporal and Spatial Patterns of Cell-fate Attainment During Early Flower Development
In real biological systems, populations of meristematic cells differentiate into different cell types in stereotyped temporal sequences and spatial patterns. The first primordial cells to be determined in the flower meristem are those of sepals, then those of petals, stamens and carpels going from the periphery to the center of the floral meristem. This suggests that in the population of meristematic cells the most probable temporal order in which each attractor is visited follows the same sequence (Alvarez-Buylla et al., 2008). Recent results from another theoretical approach show that the sequence of floral organ determination can be recovered by introducing some level of stochasticity (random noise) in the GRN dynamics, namely, a degree of error in the updating dynamical rules of the GRN (Alvarez-Buylla et al., 2008). These results are consistent with a handful of other recent studies showing that stochasticity at the molecular scale may contribute to the formation of spatiotemporal patterns in development (see review in Raj and van Oudenaarden, 2008). Studies with the stochastic version of the FOS-GRN also concluded that the relative position of the basins is important in determining the most probable temporal sequence of cell-fate attainment referred to above (Alvarez-Buylla et al., 2008). This fascinating result certainly suggests that the stereotypical temporal pattern of cell fate specification at early stages of flower development may be an emergent and robust consequence of the complex GRN underlying cell-fate determination and that, in principle, it could take place in the absence of inducing signals, emerging only as a result of the stochastic fluctuations that occur during transcriptional regulation (Alvarez-Buylla et al., 2008). Ongoing modeling efforts are explicitly focusing on spatial domains, and exploring the need and sufficiency of different cell-cell communication mechanisms or physical fields (e.g., created by curvature or tension forces) that could provide positional information for spatio-temporal cell patterning during early stages of flower development.
It is important to mention that the FOS-GRN modeled up to now is an abstraction of the qualitative regulatory logic underlying the IM and FM subregionalization during early stages of flower development when the ABC patterns are established. However, other regulatory modules for meristem positioning, growth and polarity, among others, still need to be considered in order to fully understand spatiotemporal cell patterning and morphogenesis of IM and FM. Some genes interacting with FOS-GRN components (e.g. AGL24, BEL, RBE and those described in the last section of Table 1) that do not seem to directly affect cell-type determination in the floral meristem, could link the FOS-GRN with: a) signaling pathways (e.g. Díaz and Alvarez-Buylla, 2006); b) genes involved in cell growth and proliferation both before and after the partitioning of the floral meristem into the four concentric regions; and c) other types of downstream genes or modules that are important during cell sub-differentiation and organogenesis at later stages of flower development.
A complete understanding of flower morphogenesis will continue to require multidisciplinary approaches and modeling tools that help unravel how such single-cell GRNs are coupled in explicit cellularized spatial domains and physicochemical fields (e.g. Jönsson et al., 2005, Savage et al., 2008; Benítez et al., 2008), including metabolism, signaling, and emergent gradients of morphogens (e.g., auxin), cell growth and proliferation, mechanical forces and cell-cell communication mechanisms. All of these are likely to feedback in non-linear ways from and to the GRNs underlying cell differentiation or proliferation (for example see Hamant et al., 2008).
It is important to keep in mind, for example, that plant cell growth in meristems is symplastic. This implies that the contacts between cells are preserved because there is no displacement or sliding at middle lamellas that join neighboring cells (Priestley, 1930 and Erickson, 1986; cited in Kwiatkowska, 2008). Therefore, overall plant growth could be modeled using the principles of solid body mechanics (see review in Kwiatkowska, 2008). However plant cells also grow anisotropically which implies a variation in the directional growth rates at a given point (Baskin, 2005). Hence, meristem growth has rather been modeled using the principles of continuum mechanics, computing variables that characterize plastic strain (Goodall and Green, 1986; for review see Green, 1999).
Summary of evidence for the FOS-GRN gene interactions shown in Figures 20–23 (ChIP, chromosome immunoprecipitation; EMSA, electrophoretic mobility shift assays; arrows indicate gene induction and bars repression; Espinosa-Soto et al., 2004; Chaos et al., 2006).
Some of the contributions that have used the flower organ specification GRN model in order to test, advance or discuss novel conceptual or methodological approaches.
Some quantitative mesoscopic models for flower development and growth in Arabidopsis and other angiosperms have been put forward (e.g., Rolland-Lagan et al., 2003; Lee et al., 2004; Skryabin et al., 2004; Mündermann et al., 2005). A finite element model of the SAM has also shown, for example, that lateral bulging of the meristem surface leading to the formation of a primordium results in a gradient of shear stresses with high shear stress at the point where the future primordium emerges (Selker et al., 1992; reviewed in Kwiatkowska 2008). More recently, it was shown that cells in the Arabidopsis SAM orient their cortical microtubules along lines of mechanical stress generated during tissue formation, and this then affects the mechanical properties of the cell, thus establishing a feedback loop (Hamant et al., 2008). This seems to be particularly relevant during the formation of the groove between the apical meristem and the primordium of lateral organs, but less so during growth and differentiation, because the lateral organ primordia are not affected when the microtubular network is disintegrated by a drug (Hamant et al., 2008). This implies that the mechanical feedback loop described is likely to act in parallel with the previously described auxin-mediated patterning mechanism (Laufs et al., 2009). Similar morphogenetic mechanisms are likely to be at work in flower meristem and floral organ development, and both morphogenetic mechanisms connected to the functional regulatory modules, including FOS-GRN and others that have been partly elucidated and reviewed in this Chapter.
5. CONCLUSIONS AND PERSPECTIVES
Arabidopsis has been indispensable in unraveling the molecular genetic bases of the stereotypical and most conserved aspects of flower development. It has also been used to resolve some basic mechanisms of floral meristem determination, as well as floral organ cell differentiation and morphogenesis. The challenge ahead will be to understand how modules regulating each aspect of flower development are interconnected among themselves and with signal transduction pathways that respond to environmental and internal cues to yield coupled GRN spatiotemporal dynamics during flower development. Such dynamics likely involve feedback from physical or mechanical forces, structural and geometric characteristics of domains of activity and from cell dynamics (cell growth and division) in complex ways still requiring multiple theoretical multilevel models and coordinated experimental research. Different functional modules are now being characterized (Figure 24 and Table S1 ()) and shown to regulate some of the main processes involved in flower development. Some of these modules or their components may participate in one or more flower developmental process and data on the functions and interactions of genes are becoming available to enable new dynamic computational models of GRN and signaling pathways during flower development (Figure 24 and Table S1 ()).
Computational models for the gene regulatory module that underlies patterning of the inflorescence meristem and determination of the primordial cell types during early stages of flower organ specification, have demonstrated the potential and need of theoretical dynamic approaches in understanding complex GRN underlying flower development. But information on each regulatory module and the interconnections between modules and with signal transduction pathways is still scanty.
It would be fascinating to unravel which molecular components, circuits, or sub-networks underlie the development and evolution of the diversity of flower forms and the variations around the overall conserved “theme” of floral structure among angiosperms. This will be possible with integrated multidisciplinary approaches addressing pending questions. For example, in order to understand how a flower meristem forms will require knowledge of the regulatory mechanisms underlying mechanoreception and cell wall, microfibril and microtubule behaviour. How are such mechanisms interconnected or coordinated with the cell differentiation GRNs as well as with the morphogen-mediated patterning mechanisms? The challenge ahead consists in integrating mesoscopic mechanical and morphogen-gradient models with experimentally grounded models of the GRNs underlying cell behaviour, dynamics and differentiation. The aim is to build multilevel computational modeling frameworks that can be used to test the sufficiency and necessity of contrasting mechanisms, which scale from the biochemical and GRN level to the physical factors constraining plant growth (Hogeweg, 2002). Ideally, joint efforts in modeling, bioinformatics and experimentation continually feeding back on each other should give a better understanding of flower, and more generally, plant development and evolution.
Notwithstanding the usefulness of Arabidopsis, such a grand challenge will surely benefit or require comparative experimental and evolutionary studies of other angiosperms with divergent floral structures such as the monocots, other eudicots and basal angiosperms. Such an approach has been successful in understanding and interpreting morphological traits of plants (Kaplan, 2001). Recently, studies in non-model monocots such as orchids (Tsai et al., 2004; Xu et al., 2006) and commelinids. (Ochiai et al., 2004), in maize and rice (Whipple et al., 2004; Xu and Kong, 2007), in members of the Solanaceae, such as tomato (Hileman et al., 2006; de Martino et al., 2006), and in basai angiosperms (Soltis et al., 2007) among others, have started to demonstrate the power of coupling functional and evolutionary questions of a comparative approach with detailed molecular experimentation in several species.
Findings from diverse groups of angiosperms, mostly comparative analyses of ABC gene expression data among diverse angiosperm groups (especially basal angiosperm taxa), with emphasis on the A and B class genes, have already been used to account for the underlying genetic differences in the diversity of petal and stamen morphology among extant flowering plants (Kim et al., 2005; Rudall, 2007). Figure 25 shows a diagrammatic and very simplified angiosperm phylogeny and the variations observed in the domains of expression of the ABC class genes in selected species, representative of the morphological diversity present in their respective angiosperm lineages. Overall, these approaches are helping refine our knowledge of flower development, and will be instrumental in understanding the canonical GRN modules involved in flower formation and discovering variations.
We greatly acknowledge Diana Romo for her help formatting this chapter. We thank Silvia Espinosa-Matias from the Laboratorio de Microscopia Electronica de Barrido Facultad de Ciencias UNAM for most of the SEMs presented. Many thanks to John L. Bowman, who provided SEM pictures in Figures 4A, 4B, 4C, 4I, 5A (with permission of the American Society of Plant Biology), Figure 11 (with permission of the International Journal Developmental Biology) and Figure 12. Work in Álvarez-Buylla/Garay-Arroyo/ García-Ponce, Gamboa de Buen and de Folter laboratories is financed by: Mexican Science Council (CONACyT 41848; 81542; 81433; 90565; 82826) and UNAM (PAPIIT 221406; 210408; 223607). The de Folter laboratory is also financed by CONCyTEG. We also thank the students in our laboratory for continued feedback and useful discussions that have enriched this work. Particular thanks also go to M. Aldana, C. Espinosa-Soto, D. Hartasánchez, P. Padilla, and G. Santos for ideas in Section 4.
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