developmental biology 8e ch20

THE DEVELOPMENTAL STRATEGIES OF PLANTS have evolved separately from thoseof the animals over millions of years. The two kingdoms have many common-alities (and the land plants are sometimes referred to as “embryophytes,” call-ing attention to the significance of the embryo in their life histories), but someof the challenges and solutions found in plants are sufficiently unique to war-rant separate discussion in this chapter. What are the fundamental differencesbetween development in animals and development in the land plants?

• Plant cells do not migrate. Plant cells are trapped within rigid cellulosewalls that generally prevent cell and tissue migration. Plants, like mostmetazoan animals, develop three basic tissue systems (dermal, ground, andvascular), but do not rely on gastrulation to establish this layered system oftissues. Plant development is highly regulated by the environment, a strate-gy that is adaptive for a stationary organism.

• Plants have sporic meiosis rather than gametic meiosis. That is, meiosis inplants produces spores, not gametes. Plant gametes are produced by mitoticdivisions following meiosis.

• The life cycle of land plants (as well as many other plants) includes both

diploid and haploid multicellular stages. This type of life cycle is referredto as alternation of generations and results in two different multicellular bodyplans over the life cycle of an individual.

• Plant germ cells are not set aside early in development. While this is alsothe case in several animal phyla, it is the case for all plants.

• Plants undergo extended morphogenesis. Clusters of actively dividing cellscalled meristems, which are similar to stem cells in animals, persist longafter maturity. Meristems allow for iterative development and the formationof new structures throughout the life of the plant.

• Plants have tremendous developmental plasticity. Many plant cells arehighly plastic. While cloning in animals also demonstrates plasticity, plantsdepend far more heavily on this developmental strategy. For example, if ashoot is grazed by herbivores, meristems in the leaf often grow out to replacethe lost part. (This strategy has similarities to the regeneration seen in someanimals.) Whole plants can be regenerated from some single cells. In addition,a plant’s form (including branching, height, and relative amounts of vegeta-tive and reproductive structures) is greatly influenced by environmental fac-tors such as light and temperature, and a wide range of morphologies canresult from the same genotype (see Figure 2.15). The amazing level of plastici-ty found among the plants may help compensate for their lack of mobility.

20An Overview of PlantDevelopmentSusan R. SingerLaurence McKinley Gould Professor of the Natural Sciences, Carleton College

“The search for differences or fun-damental contrasts … has occupiedmany men’s minds, while the searchfor commonality of principle oressential similarities, has been pur-sued by few; the contrasts are apt toloom too large, great though theymay be.” D’ARCY THOMPSON (1942)

“Do not quench your inspirationand your imagination; do notbecome the slave of your model.”VINCENT VAN GOGH

[From Developmental Biology, Eighth Edition, by Scott F. Gilbert, published by Sinauer Associates, Inc.]

Chapter 20 pagestest�.qxd 8/5/10 10:35 AM Page 627

© 2010 Sinauer Associates, Inc. This material cannot be copied, reproduced, manufactured, or disseminated in any form without the express written permission of the publisher.

• Developmental mechanisms evolved independently

in plants and animals. The last common ancestor ofplants and animals was a single-celled eukaryote.Genome-level comparisons indicate that there is mini-mal homology between the genes and proteins used toestablish body plans in plants and animals (Meyerowitz2002). While both homeobox and MADS box genes werepresent in the last common ancestor of plants and ani-mals, the MADS box family controls major developmen-tal regulatory processes in plants, but not in animals.

Despite the major differences among many plants and ani-mals, developmental genetic studies are revealing somecommonalities in the logic of their pattern formation, alongwith evolutionarily distinct solutions to the problem of cre-ating three-dimensional form from a single cell.

The green plants include many organisms, from algaeto flowering plants (angiosperms). Recent phylogeneticstudies show a common lineage for all green plants, dis-tinct from the red and brown plants. While comparisonsof developmental strategies among diverse plants is bothfascinating and informative, this chapter focuses primari-ly on the flowering plants (angiosperms). The goal is toexamine plant development within the larger context ofdevelopmental biology.

Gamete Production in Angiosperms

Plants have both multicellular haploid and multicellulardiploid stages in their life cycles, and embryonic develop-ment is seen only in the diploid generation. The embryo,however, is produced by the fusion of gametes, which areformed only by the haploid generation. Understanding therelationship between the two generations is important inthe study of plant development.

GametophytesUnlike animals, plants have multicellular haploid and mul-ticellular diploid stages in their life cycles (see Chapter 2).Gametes develop in the multicellular haploid gametophyte

(from the Greek phyton, “plant”). Fertilization gives rise toa multicellular diploid sporophyte, which produces hap-loid spores via meiosis. This type of life cycle is called ahaplodiplontic life cycle (Figure 20.1). It differs from thediplontic life cycle of animals, in which only the gametesare in the haploid state.

In a haplodiplontic life cycle, gametes are not the directresult of a meiotic division. Diploid sporophyte cells under-go meiosis to produce haploid spores. Each spore goesthrough mitotic divisions to yield a multicellular, haploidgametophyte. There are two types of spores in angio-sperms. Megaspores produce female gametophytes, whilemicrospores produce male gametophytes. Male and femalegametophytes have distinct morphologies. Wind or mem-

bers of the animal kingdom deliver the male gameto-phyte—pollen—to the female gametophyte. Mitotic divi-sions within the gametophytes are required to producegametes. The diploid sporophyte results from the fusionof two gametes. Among land plants, the gametophytes andsporophytes of a species have distinct morphologies, andhow a single genome can be used to create two uniquemorphologies is an intriguing puzzle.

At first glance, angiosperms may appear to have diplon-tic life cycles because the gametophyte generation has beenreduced to just a few cells (Figure 20.2). However, mitoticdivision follows meiosis in the sporophyte, resulting in amulticellular gametophyte, which produces eggs or sperm.All of this takes place in the organ that is characteristic ofthe angiosperms: the flower.

POLLEN The pollen grain is an extremely simple multicel-lular structure (Figure 20.3). The outer wall of the pollengrain, the exine, is composed of resistant material provid-ed by both the tapetum (sporophyte generation that pro-vides nourishment for developing pollen) and themicrospore (gametophyte generation). The inner wall, the

628 CHAPTER 20

Diploid generation Haploid generation

Path C Path B

GametesFERTILIZATION

MEIOSIS

Mitosis

Mitosis

Path A1n

organism

Path D2n

organism

2n

1n

1n

1n

20.1 Plant life cycles. In plants there is anevolutionary trend from sporophytes that arenutritionally dependent on autotrophic gameto-phytes to the opposite—gametophytes that aredependent on autotrophic sporophytes. Thistrend is exemplified by comparing the life cycles of mosses and ferns to that of angiosperms. [Note: This Web topic is included at the end of this chapter.]

W E B S I T E

FIGURE 20.1 Plants have haplodiplontic life cycles thatinvolve mitotic divisions (resulting in multicellularity) in boththe haploid and diploid generations (paths A and D). Most ani-mals are diplontic and undergo mitosis only in the diploid gen-eration (paths B and D). Multicellular organisms with haplonticlife cycles follow paths A and C.



intine, is produced by the microspore. A mature pollengrain consists of two cells: a tube cell, and a generative

cell within the tube cell. The nucleus of the tube cell guidespollen germination and the growth of the pollen tube after

the pollen lands on the stigma of a female gametophyte.The generative cell divides to produce two sperm. One ofthe two sperm will fuse with the egg cell to produce thenext sporophyte generation. The second sperm will par-

AN OVERVIEW OF PLANT DEVELOPMENT 629

Diploid sporophyte generation Haploid gametophyte generation

Filament

Microsporangium(2n)

Embryo

Megasporangium(2n)

Microspores (1n)

Pollen

Gametophyte

GametophyteEmbryo sac

Megaspores (1n)

Sporophyte (2n)

Flower

Ovary

Ovule

Petals

Pollen tube

Embryo sacEndosperm

Seed coat

CarpelAntherStamen

FERTILIZATION

MEIOSIS Mitosis

Seedgermination

FIGURE 20.2 Life cycle of an angiosperm, represented hereby a pea plant (genus Pisum). The sporophyte is the dominantgeneration, but multicellular male and female gametophytesare produced within the flowers of the sporophyte. Cells of themicrosporangium within the anther undergo meiosis to pro-duce microspores. Subsequent mitotic divisions are limited, butthe end result is a multicellular pollen grain. Integuments andthe ovary wall protect the megasporangium. Within the mega-

sporangium, meiosis yields four megaspores—three small andone large. Only the large megaspore survives to produce thefemale gametophtye (the embryo sac). Fertilization occurswhen the male gametophyte (pollen) germinates and thepollen tube grows toward the embryo sac. The sporophyte gen-eration may be maintained in a dormant state, protected by theseed coat.

(A) (B)

Exine

Intine

Tube cell

Tube cell nucleus

Generativecell

1n 1n

FIGURE 20.3 (A) Pollen grainshave intricate surface patterns, asseen in this scanning electronmicrograph of aster pollen. (B) Apollen grain consists of a cellwithin a cell. The generative cellwill undergo division to producetwo sperm cells. One will fertilizethe egg, and the other will joinwith the polar nuclei, yielding theendosperm.



ticipate in the formation of the endosperm, a structure thatprovides nourishment for the embryo.

THE OVARY The angiosperm ovary is part of the carpel ofa flower, which gives rise to the female gametophyte (Fig-ure 20.4). The carpel consists of the stigma (where thepollen lands), the style, and the ovary. Following fertiliza-tion, the ovary wall will develop into the fruit. This uniqueangiosperm structure provides further protection for thedeveloping embryo and also enhances seed dispersal byfrugivores (fruit-eating animals). Within the ovary are oneor more ovules attached by a placenta to the ovary wall.Fully developed ovules are called seeds.

The ovule has one or two outer layers of cells, called theinteguments. The integuments enclose the megaspo-

rangium, which contains sporophyte cells that undergomeiosis to produce megaspores (see Figure 20.2). There isa small opening in the integuments called the micropyle,through which the pollen tube will grow. The integumentsdevelop into the seed coat, a waterproof physical barrierthat protects the embryo. When the mature embryo dis-perses from the parent plant, diploid sporophyte tissueaccompanies the embryo in the form of the seed coat andthe fruit.

Within the ovule, meiosis and unequal cytokinesis yieldfour megaspores. The largest of these megaspores under-goes three mitotic divisions to produce the female game-tophyte, a seven-celled embryo sac with eight nuclei (Fig-ure 20.5). One of these cells is the egg. Two synergid cells

surround the egg and the pollen tube enters the embryosac by penetrating one of the synergids. The central cell

contains two or more polar nuclei, which will fuse with thesecond sperm nucleus and develop into the polyploidendosperm. Three antipodal cells form at the opposite endof the embryo sac from the synergids and degeneratebefore or during embryonic development. There is noknown function for the antipodals. Genetic analyses offemale gametophyte development in maize and Arabidop-sis* are providing insight into the regulation of the specif-ic steps in this process (Pagnussat et al. 2005).

Pollination

Pollination refers to the landing and subsequent germina-tion of the pollen on the stigma. Hence it involves an inter-action between the gametophytic generation of the male(the pollen) and the sporophytic generation of the female(the stigmatic surface of the carpel). Pollination can occurwithin a single perfect flower that contains both male andfemale gametophytes (self-fertilization), or pollen can landon a different flower on the same or a different plant.About 96 percent of flowering plant species produce maleand female gametophytes on the same plant. However,about 25 percent of these produce two different types offlowers on the same plant, rather than perfect flowers.

Staminate flowers lack carpels, while carpellate flow-ers lack stamens. Maize plants, for example, have stami-nate (tassel) and carpellate (ear) flowers on the same plant.Such species, which include the majority of the angio-sperms, are considered to be monoecious (Greek mono,

630 CHAPTER 20

Stigma

Megasporangium

Integuments

Megaspores

Micropyle

1n

1n

1n1n

Placenta

Style

Ovule

Ovary

Sepals

Ovary wall(eventual fruit)

Placenta

FIGURE 20.4 The carpel consists of the stigma, the style, andan ovary containing one or more ovules. Each ovule containsmegasporangia protected by two layers of integument cells.The megasporangia divide meiotically to produce haploidmegaspores. All of the carpel is diploid except for the mega-spores, which divide mitotically to produce the embryo sac (the female gametophyte).

Antipodal cells

Integuments

Micropyle(pollen entrypoint)

Polar nucleiSynergid

Egg

FIGURE 20.5 The embryo sac is the product of three mitoticdivisions of the haploid megaspore; it comprises seven cells andeight haploid nuclei. The two polar nuclei in the central cell willfuse with the second sperm nucleus and produce the endo-sperm that will nourish the egg. The other six cells, includingthe egg, contain one haploid nucleus each.

*A small weed in the mustard family, Arabidopsis is used as a model

organism because of its very small genome.



“one”; oecos, “house”). The remaining 4 percent of species(e.g., willows, maples, and date palms*) produce stami-nate and carpellate flowers on separate plants. Thesespecies are considered to be dioecious (“two houses”).Only a few plant species have true sex chromosomes, yetthey arose several times in flowering plant evolution(Charlesworth 2002). The terms “male” and “female” aremost correctly applied only to the gametophyte genera-tion, not to the sporophyte (Cruden and Lloyd 1995).

SELF-INCOMPATIBILITY The arrival of a viable pollen grainon a receptive stigma does not guarantee fertilization.Interspecific incompatibility refers to the failure of pollenfrom one species to germinate and/or grow on the stigmaof another species. Intraspecific incompatibility is incom-patibility that occurs within a species. Self-incompatibil-

ity—incompatibility between the pollen and the stigmasof the same individual—is an example of intraspecificincompatibility (see Kao and Tsuikamoto 2004). Self-incom-patibility blocks fertilization between two genetically sim-ilar gametes, increasing the probability of new gene com-binations by promoting outcrossing (pollination by adifferent individual of the same species). Groups of close-ly related plants can contain a mix of self-compatible andself-incompatible species.

Several different self-incompatibility systems haveevolved. Recognition of self depends on the multiallelicself-incompatibility (S) locus (Nasrallah 2002). Gametophyt-ic self-incompatibility occurs when the S allele of the pollengrain matches either of the S alleles of the stigma (remem-ber that the stigma is part of the diploid sporophyte gen-eration, which has two S alleles, while a single pollen graincarries one S allele). In this case, the pollen tube begins

developing but stops before reaching the micropyle (Fig-ure 20.6A). Sporophytic self-incompatibility occurs when oneof the two S alleles of the pollen-producing sporophyte(not the gametophyte) matches one of the S alleles of thestigma (Figure 20.6B). Most likely, sporophyte contribu-tions to the pollen exine are responsible for this type of self-incompatibility.

The S locus consists of several physically linked genesthat regulate recognition and rejection of pollen. An S genehas been cloned that codes for an RNase (called S RNase)that is sufficient, in the gametophytically self-incompati-ble petunia pistil, to recognize and reject self-pollen (Leeet al. 1994). The pollen component of gametophytic self-incompatibility in the petunia, SLF (S-locus, F-box), is anF-box gene† within the S locus (Sijacic et al. 2004).

A different, more rapid gametophytic response to self-incompatibility has been investigated in poppies, a rela-tive of the more basal flowering plants. Calcium ions accu-mulate in the tip of the pollen tubes, where open calciumchannels are concentrated (Jaffe et al. 1975; Trewavas andMalho 1998). There is direct evidence that pollen tubegrowth in the poppy is regulated by a slow-moving Ca2+

wave controlled by the phosphoinositide signaling path-way (Figure 20.7; Franklin-Tong et al. 1996). Ca2+ influxoccurs at both the tip of the pollen tube and on the shanks.Altered calcium influx is observed when the pollen tube isself-incompatible with the style, which leads to F-actindepolymerization, destabilization of the pollen cytoskele-ton, and cessation of pollen tube growth (Franklin-Tong etal. 2002; Franklin-Tong and Franklin 2003). The incompat-ible pollen tube then undergoes programmed cell death(Thomas and Franklin-Tong 2004).

In sporophytic self-incompatibility, a ligand on thepollen is thought to bind to a membrane-bound kinasereceptor in the stigma, starting a signaling process that


Pollen Pollen

(A)–Gametophytic self-incompatibility (B)–Sporophytic self-incompatibility

S1/S2Stamen

S2/S3Carpel

S2/S3Carpel

S1/S2Stamen

S2/S3Carpel

S2/S3Carpel

S1 S2 S2S1 FIGURE 20.6 Self-incompatibility.S1, S2, and S3 are different alleles ofthe self-incompatibility (S) locus. (A)Plants with gametophytic self-incom-patibility reject pollen only when thegenotype of the pollen (i.e., thegametophyte) matches either one ofthe carpel’s two alleles. (B) In sporo-phytic self-incompatibility, the geno-type of the pollen parent (i.e., thesporophyte), not just that of the hap-loid pollen grain, can trigger anincompatibility response.

*The discovery that plants had sexes was important to the economy

of date palms in the ancient Near East over two thousand years

ago. Since only the female trees bore fruit, date farmers planted just

a few male trees, then hand-pollinated the many female trees. This

practice greatly increased the fruit yield per acre, and such pollina-

tion events became associated with spring fertility festivals.

†Members of the F-box family of genes share a common “F-box

domain” for binding transcription factors. Although some F-box

genes have been found in other eukaryote groups, most of these

genes are unique to the plants.



leads to pollen rejection. In Brassica, one of the genes of theS locus encodes a transmembrane serine-threonine kinase(SRK) that functions in the epidermis of the stigma andbinds a cysteine-rich peptide (SCR) from the pollen (Fig-ure 20.8; Kachroo et al. 2001).

There are numerous examples of plant populations thathave switched from self-incompatible to self-fertilizing sys-tems. Changes in the S locus, specifically the SKR and SCRgenes, could account for these evolutionary changes. TheNasrallahs (2002) created self-incompatible Arabidopsisthaliana plants (which are normally self-compatible) byintroducing the SKR and SCR genes that encode self-recog-nizing proteins from A. lyrata (a self-incompatible species).This experiment demonstrates that A. thaliana still has allof the downstream components of the signal cascade thatcan lead to pollen degradation. The mechanism of pollendegradation is unclear, but appears to be highly specific.

POLLEN GERMINATION If the pollen and the stigma are com-patible, the pollen takes up water (hydrates) and the pollentube emerges. The pollen tube grows down the style of thecarpel toward the micropyle (Figure 20.9). The tube nucle-us and the sperm cells are kept at the growing tip by bandsof callose (a complex carbohydrate). It is possible that thismay be an exception to the “plant cells do not move” rule,as the generative cell(s) appear to move forward via adhe-sive molecules (Lord 2000). Pollen tube growth is quiteslow in gymnosperms (up to a year), while in someangiosperms the tube can grow as rapidly as 1 cm per hour.

Genetic approaches have been useful in investigatinghow the growing pollen tube is guided toward unfertil-ized ovules. In Arabidopsis, the pollen tube appears to beguided by a long-distance signal from the ovule (Hul-skamp et al. 1995; Wilhelmi and Preuss 1999). Analysis ofpollen tube growth in ovule mutants of Arabidopsis indi-cates that the haploid embryo sac is particularly importantin the long-range guidance of pollen tube growth. Mutants

632 CHAPTER 20

Dir

ectio

n of

Ca

2+ w

ave

TubenucleusCa2+ channels

Embryosac Ovule

Calloseplug

Pollentube

Spermcells

× ×

SRK-SCRbinding

Pollen coat

Pollen tube inhibition Pollen tube growth

Stigma cell wall

å

(A) Self-pollination (self-self)

(B) Cross-pollination (self-nonself)

S1

S2

S1

S2

ç

åS1

S2

S3

S4

SRKactivation

NOSRK-SCRbinding

NO SRKactivation

ç

FIGURE 20.7 Calcium and pollen tube tip growth. After com-patible pollen germinates, the pollen tube grows toward themicropyle. Waves of calcium ions play a key role in this growthof the tube. (After Franklin-Tong et al. 1996.)

FIGURE 20.8 Receptor-ligand self-recognition is the key toself-incompatibility in Brassicas. Allelic variability in both the SRKand SRC genes leads to a variety of possible combinations of lig-and and receptor proteins. Unlike the common self-recognitionsystems of animals, including immunity and mating, self-incom-patibility results from the binding of SRK and SRC proteins ofself (from allelic S loci) rather than nonself. (After Nasrallah 2002;photographs courtesy of J. Nasrallah.)

Chapter 20 pagestest�.qxd 8/5/10 10:35 AM 32


with defective sporophyte tissue in the ovule but a normalhaploid embryo sac appear to stimulate normal pollen tubedevelopment.

While the evidence points primarily to the role of thegametophyte generation in pollen tube guidance, diploidcells may make some contribution. The Arabidopsis genePOP2 encodes an enzyme that degrades γ-amino butyricacid (GABA) and establishes a gradient of GABA in thestyle up to the micropyle (Palanivelu et al. 2003). The pop2mutant has misguided pollen tube growth, presumablybecause there is no GABA gradient. POP2 is expressed inboth the pollen and the style, which may explain whywild-type pollen tubes find their way to the micropylewhen the style has a pop2 genotype. The wild-type enzymein the pollen tube may degrade GABA and create a suffi-cient gradient to guide itself to the micropyle.

As the final step in pollen guidance, the two synergidcells in the embryo sac may attract the pollen tube. In Tore-nia fournieri (wishbone flower), the embryo sac protrudesfrom the micropyle and can be cultured. In vitro, it canattract a pollen tube. Higashiyama and colleagues (2001)used a laser beam to destroy individual cells in the embryosac and then tested whether or not pollen tubes were stillattracted to the embryo sac. A single synergid was suffi-cient to guide pollen tubes; however, when both synergidswere destroyed, pollen tubes were not attracted to the sac.

Fertilization

The growing pollen tube enters the embryo sac throughthe micropyle and grows through one of the synergids. Thetwo sperm cells are released, and a double fertilization

event occurs (see Southworth 1996). One sperm cell fuseswith the egg, producing the zygote that will develop intothe sporophyte. The second sperm cell fuses with the bi-or multinucleate central cell, giving rise to the endosperm,

which nourishes the developing embryo. This second eventis not true fertilization in the sense of male and femalegametes undergoing syngamy (fusion)—that is, it does notresult in a zygote, but in nutritionally supportive endo-derm. When you eat popcorn, you are actually eating“popped” endosperm. The other accessory cells in theembryo sac degenerate after fertilization.

The zygote of the angiosperm produces only a singleembryo.* Double fertilization, first identified a century ago,is generally restricted to the angiosperms, but it has alsobeen found in the gymnosperm genera Ephedra and Gne-tum, although no endosperm forms. Friedman (1998) hassuggested that endosperm may have evolved from a sec-ond zygote “sacrificed” as a food supply in an early gym-nosperm lineage with double fertilization.

Investigations of Amborella, the most closely relatedextant relative of the basal angiosperm (Figure 20.10A), isproviding information on the evolutionary origin of theendosperm (Brown 1999). It is probable that the firstangiosperm had a four-nucleus embryo sac (Williams andFriedman 2002, 2004). The critical cell to consider is thecentral cell, which is fertilized by the second sperm to cre-ate the endosperm. In eight-nuclei embryo sacs, there areseven cells. The central cell contains two nuclei and, whenfertilized, produces a triploid endosperm. In Nuphar, abasal angiosperm, the embryo sac consists of four nuclei,and the central cell has a single nucleus that, when fertil-ized, develops into a 2n endosperm (Figure 20.10B). The2n endosperm provides convincing evidence that the four-celled embryo sac in Nuphar does not result from the degra-dation of four nuclei. If other cells had degraded, a 3nendosperm would be predicted.


(A) (B)

Pollen tubes

Sperm nuclei

Style cells

Pollen tube

Tube cellnucleus

FIGURE 20.9 Pollen tube germination. (A) Scanning electron micrograph of an Arabidopsis pollen tube en route to the ovule for fertilization. (B) Lily pollen tubesgrown in vivo and removed from the ovary. Each green strand is an individual pollentube and contains two sperm nuclei (bright blue stain) and a fainter (lighter blue) tubecell nucleus. Note the huge number of pollen tubes, all “racing” to fertilize a singleegg. (Photographs courtesy of E. Lord.)

*The gymnosperm zygote, on the other hand, produces two or

more embryos after cell division begins, by a process known as

cleavage embryogenesis.



Fertilization is not an absolute prerequisite for angi-osperm embryonic development (Mogie 1992). Embryoscan form within embryo sacs from haploid eggs and fromcells that did not divide meiotically. This phenomenon iscalled apomixis (Greek, “without mixing”), and results inviable seeds. The viability of the resulting haploid sporo-phytes indicates that ploidy alone does not account for themorphological distinctions between the gametophyte andthe sporophyte. Embryos can also develop from culturedsporophytic tissue. These embryos develop with no associ-ated endosperm, and they lack a seed coat.

Embryonic Development

Embryogenesis

In plants, the term embryogenesis covers development fromthe time of fertilization until dormancy occurs. The basicbody plan of the sporophyte is established during embryo-genesis; however, this plan is reiterated and elaboratedafter dormancy is broken. The major challenges of plantembryogenesis are:

•To establish the basic body plan. Radial patterning pro-duces three tissue systems (dermal, ground, and vascu-lar), and axial patterning establishes the apical-basal(shoot-root) axis.

•To set aside meristematic tissue for postembryonic elab-oration of the body structure (leaves, roots, flowers, etc.).

•To establish an accessible food reserve for the germinatingembryo until the embryo becomes autotrophic.

Embryogenesis is similar in all angiosperms in terms of theestablishment of the basic body plan. There are differencesin pattern elaboration, however, including differences inthe precision of cell division patterns, the extent ofendosperm development, cotyledon development, and theextent of shoot meristem development (Esau 1977; Steevesand Sussex 1989; Johri et al. 1992).

MATERNAL EFFECTS IN EARLY EMBRYOGENESIS Maternal effectgenes play a key role in establishing embryonic patternsin animals (see, for example, the discussion of Drosophilain Chapter 9). The extent of extrazygotic gene involvementin plant embryogenesis is an open question, complicatedby at least three potential sources of influence: sporophytictissue, gametophytic tissue, and the polyploid endosperm.All of these tissues are in close association with the egg/zygote (Ray 1998). Endosperm development could also beaffected by maternal genes. Sporophytic and gametophyt-ic maternal effect genes have been identified in Arabidop-sis, and it is probable that the endosperm genome influ-ences the zygote as well.

The first maternal effect gene identified, SHORTINTEGUMENTS 1 (SIN1), must be expressed in the sporo-phyte for normal embryonic development to occur (Ray etal. 1996). Two transcription factors (FBP7 and FBP11) areneeded in the petunia sporophyte for normal endospermdevelopment (Columbo et al. 1997). A female gametophyt-ic maternal effect gene, MEDEA,* has protein domains sim-ilar to those of a Drosophila maternal effect gene (Gross-niklaus et al. 1998). Curiously, MEDEA is in the Polycombgene group (see Chapter 9), whose products alter chro-matin, directly or indirectly, and affect transcription.MEDEA affects an imprinted gene (see Chapter 5) that is

634 CHAPTER 20

Synergids

Egg

Central cell

(B)(A)

FIGURE 20.10 Ancestral angiosperms.(A) Amborella trichopoda. This plant ismore closely related to the firstangiosperm than any other extantspecies. (B) Ancestral angiosperms prob-ably had 2n endosperms. The embryosac of the basal angiosperm Nuphar (yel-low water lily) has a single nucleus in itscentral cell , which when fertilized willproduce a 2n endosperm. DAPI stainingshows that the DNA content is 1n , not n.Because this is a section of tissue, theegg cell is hidden behind the two syn-ergids and is shown in the insert. (A photograph courtesy of Sandra K.Floyd; B photograph courtesy of WilliamFreidman.)

*Another name from Greek mythology, after Euripides’ Medea,

who killed her own children.



expressed by the female gametophyte and by maternallyinherited alleles in the zygote, but not by paternally inher-ited alleles (Vielle-Calzada et al. 1999). The significance ofmaternal effect genes in establishing the sporophyte bodyplan has been highlighted by Pagnussat and co-workers’(2005) screen of 130 female gametophytic mutants. Near-ly half the mutations were in a maternal gene, furtherimplicating the female gametophyte or maternal genomein embryo development.

FIRST ASYMMETRIC DIVISION: BROWN ALGAE Polarity is estab-lished in the first cell division following fertilization.Because angiosperm embryos are deeply embedded inmultiple layers of tissue, the establishment of polarity isalso investigated in brown algae, a model system withexternal fertilization (Belanger and Quatrano 2000; Brown-lee 2004). The zygotes of these plants are independent ofother tissues and are amenable to manipulation. The ini-tial cell division results in one smaller cell, which will form

the rhizoid (root homologue) and anchor the rest of theplant, and one larger cell, which gives rise to the thallus(the main body of the sporophyte). The point of spermentry fixes the position of the rhizoid end of the apical-basal axis. This axis is perpendicular to the plane of thefirst cell division. F-actin accumulates at the rhizoid pole(Figure 20.11A; Kropf et al. 1999). However, light or grav-ity can override this fixing of the axis and establish a newposition for cell division (Figure 20.11B; Alessa and Kropf1999). Once the apical-basal axis is established, secretoryvesicles are targeted to the rhizoid pole of the zygote (Fig-ure 20.12). These vesicles contain material for rhizoid out-growth, with a cell wall of distinct macromolecular com-


(A) (B)

Direction

of light

FIGURE 20.11 Axis formation in thebrown alga Pelvetia compressa. (A) An F-actin patch (orange) is first formed at thepoint of sperm entry; the blue spot marksthe sperm pronucleus. (B) Later, light wasshone in the direction of the arrow. Thesperm-induced axis was overridden, andan F-actin patch formed on the dark side,where the rhizoid will later form. (Pho-tographs courtesy of W. Hables.)

Secretoryvesicles

Cellplate

25 hours after fertilization

8 hours after fertilization

FIGURE 20.12 Asymmetrical cell division in brown algae.Time course from 8 to 25 hours after fertilization, showing algalcells stained with a vital membrane dye to visualize secretoryvesicles, which appear first, and the cell plate, which begins toappear about halfway through this sequence. (Photographscourtesy of K. Belanger.)



position. Targeted secretion may also help orient the firstplane of cell division. Maintenance of rhizoid versus thal-lus fate early in development depends on information inthe cell walls (Brownlee and Berger 1995). Such cell wallinformation also appears to be important in angiosperms(see Scheres and Benfey 1999).

FIRST ASYMMETRIC DIVISION: ANGIOSPERMS The basic body planof the angiosperm laid down during embryogenesis alsobegins with an asymmetrical* cell division, giving rise toa terminal cell and a basal cell (Figure 20.13). The terminalcell gives rise to the embryo proper. The basal cell formsclosest to the micropyle and gives rise to the suspensor.The hypophysis is found at the interface between the sus-pensor and the embryo proper. In some species it gives riseto a portion of the root cells. (The suspensor cells divide toform a filamentous or spherical organ that degenerateslater in embryogenesis.) In both gymnosperms andangiosperms, the suspensor orients the absorptive surfaceof the embryo toward its food source; in angiosperms, italso appears to serve as a nutrient conduit for the devel-oping embryo.

The study of embryo mutants in maize and Arabidopsishas been particularly helpful in sorting out the differentdevelopmental pathways of embryos and suspensors.Investigations of suspensor mutants (sus1, sus2, and rasp-berry1) of Arabidopsis have provided genetic evidence thatthe suspensor has the capacity to develop embryo-likestructures (Figure 20.14A,B; Schwartz et al. 1994; Yadegari

et al. 1994). In these mutants, abnormalities in the embryoproper appear prior to suspensor abnormalities.† Earlierexperiments in which the embryo proper was removedalso demonstrated that suspensors could develop likeembryos (Haccius 1963). A signal from the embryo prop-er to the suspensor may be important in maintaining sus-pensor identity and blocking the development of the sus-pensor as an embryo. Molecular analyses of these andother genes are providing insight into the mechanisms ofcommunication between the suspensor and the embryoproper (Figure 20.14C).

The SUS1 gene has been renamed DCL1 (DICER-LIKE1)because its predicted protein sequence is structurally likethat of Dicer in Drosophila melanogaster and DCR-1 inCaenorhabditis elegans (Schauer et al. 2002). These proteinsmay control the translation of developmentally importantmRNAs. This is an exciting discovery that will lead to a bet-ter understanding of the regulation of development beyondthe level of transcriptional control. Intriguingly, DCL1 hasseveral alleles that were originally assumed to be complete-ly different genes regulating very different developmentalprocesses in plants. DCL1 alleles include sin1 alleles. Thesemutants affect ovule development (discussed in the nextsection) and the transition from vegetative to reproductivedevelopment (discussed later in the chapter). The carpel fac-tory (caf1) allele of DCL1 causes indeterminancy in floralmeristems leading to extra whorls of carpels. Extrapolat-ing from Drosophila Dicer function, DCL1 protein may beinvolved in cleaving small, noncoding RNAs into evensmaller, 21- to 25-nucleotide, single-stranded RNA prod-ucts that could cleave to mRNAs and affect translation.

Many questions about the role of microRNAs as possi-ble developmental signals are arising from the work beingdone on DCL1 alleles and on leaf asymmetry genes, whichwill be discussed later (Kidner and Martienssen 2005).

636 CHAPTER 20

Zygote Basal cell

Terminal cell

2-Cell embryo

Suspensor

Embryo

Suspensor Hypophysis

Rootmeristem

Shootmeristem

Cotyledons

Globular stageembryo

Heart stageembryo

Torpedo stageembryo

FIGURE 20.13 Angiosperm embryogenesis. A representativedicot is shown; a monocot would develop only a single cotyle-don. The embryo proper forms from the terminal cell; the basalcell divides to form the suspensor, which will degenerate asdevelopment progresses. The point of interface between thesuspensor and the embryo is the hypophysis. While there arebasic patterns of embryogenesis in angiosperms, there istremendous morphological variation among species.

*Asymmetrical cell division is also important in later angiosperm

development, including the formation of guard cells of leaf stomata

and of different cell types in the ground and vascular tissue systems.

†Another intriguing characteristic of these mutants is that cell differ-

entiation occurs in the absence of morphogenesis. Thus, cell differen-

tiation and morphogenesis can be uncoupled in plant development.



RADIAL AND AXIAL PATTERNING Radial and axial patternsdevelop as cell division and differentiation continue (Fig-ure 20.15). The cells of the embryo proper divide in trans-verse and longitudinal planes to form a globular stageembryo with several tiers of cells. Superficially, this stagebears some resemblance to cleavage in animals, but thenucleus/cytoplasm ratio does not necessarily increase. Theemerging shape of the embryo depends on regulation ofthe planes of cell division and expansion, since the cellsare not able to move and reshape the embryo. Cell divisionplanes in the outer layer of cells become restricted, and thislayer, called the protoderm, becomes distinct. Radial pat-terning emerges at the globular stage as the three tissuesystems (dermal, ground, and vascular) of the plant areinitiated (Figure 20.15A). The dermal tissue (epidermis)will form from the protoderm and contribute to the outerprotective layers of the plant. Ground tissue (cortex andpith) forms from the ground meristem, which lies beneaththe protoderm. The procambium, which forms at the coreof the embryo, gives rise to the vascular tissue (xylem andphloem), which will function in support and transport. Thedifferentiation of each tissue system is at least partiallyindependent. For example, in the keule mutant of Arabidop-sis, the dermal system is defective while the inner tissuesystems develop normally (Mayer et al. 1991).

The globular shape of the embryo is lost as cotyledons

(“first leaves”) begin to form. Dicots have two cotyledons,which give the embryo a heart-shaped appearance as theyform. In monocots, such as maize, only a single cotyledonemerges.The axial body plan is evident by this heart stage

of development (Figure 20.15B). Hormones (specifically,auxins) may mediate the transition from radial to bilateralsymmetry (Liu et al. 1993).

AUXIN AND THE APICAL-BASAL AXIS The hormone auxin playsa key role in establishing the apical-basal axis of theembryo. Very early in embryogenesis, the family of pin-formed (PIN) auxin efflux carriers are asymmetrically dis-tributed (Figure 20.16A; Friml et al. 2003, 2004). The pinoid(PID) protein kinase aids in the asymmetric localization ofPIN proteins. At the 4-cell stage, auxin moves apically, butby the globular stage, the efflux carriers direct the flow ofauxin to the hypophysis, which organizes as the root meris-tem (Figure 20.16B). Disruption of the auxin morphogenet-ic gradient, either through mutation or overexpression of


(A) (B) (C)

Signals fromembryosuppressembryonicdevelopmentin suspensorcells

Signals fromsuspensor cells promoteembryonicdevelopment

FIGURE 20.14 The SUS gene (a DCL1 allele) suppressesembryonic development in the suspensor. (A) Wild-typeembryo and suspensor. (B) The suspensor of a sus mutant devel-ops like an embryo (arrow). (C) Model showing how the embryoproper suppresses embryonic development in the suspensorand the suspensor provides feedback information to theembryo. (Photographs courtesy of D. Meinke.)

(A) Radial patterning

(B) Axial patterning

Globular stage

Heart stage

Procambium(vascularlayer) Root–shoot

axis

Cotyledon

Shoot meristem

Procambium

Root meristem

Protodermlayer

Ground layer

FIGURE 20.15 Radial and axial patterning. (A) Radial patterning in angiosperms begins inthe globular stage and results in the establishment of three tissue systems. (B) The axial pat-tern (shoot-root axis) is established by the heart stage.



PID allows auxin to accumulate more apically and rootdevelopment is abnormal or inhibited (Figure 20.16C).Although auxin accumulation is necessary for apical-basalpolarity, an extensive study of the PIN proteins reveals thatauxin alone may not be sufficient to establish polarity inthe embryo (Weijers and Jürgens 2005).

The discovery of the auxin receptor provided a linkbetween the asymmetric distribution of auxin and theexpression of auxin-induced genes (Dharmasiri et al. 2005;Kipinski and Leyser 2005). In the absence of auxin, theauxin response factors (ARF) are in a repressed state (Fig-ure 20.17A). These genes form a heterodimer withAux/IAA protein, which inhibits the transcription ofauxin-induced genes. In the presence of auxin, the enzymeSCF binds Aux/IAA, catalyzing Aux/IAA ubiquitination(Figure 20.17B). Ubiquitination targets Aux/IAA for degra-dation by the 26S proteosome. Freed from the Aux/IAArepressor, ARF can act on its own to stimulate transcrip-tion or form a homo/heterodimer with another ARF to fur-ther modulate gene expression.

COTYLEDONS In many plants, the cotyledons aid in nour-ishing the plant by becoming photosynthetic after germina-

tion (although those of some species never emerge fromthe ground). In some cases—peas, for example—the foodreserve in the endosperm is used up before germination,and the cotyledons serve as the nutrient source for the ger-minating seedling.* Even in the presence of a persistentendosperm (as in maize), the cotyledons store foodreserves such as starch, lipids, and proteins. In manymonocots, the cotyledon grows into a large organ pressedagainst the endosperm and aids in nutrient transfer to theseedling. Upright cotyledons can give the embryo a torpe-do shape. In some plants, the cotyledons grow sufficient-ly long that they must bend to fit within the confines of theseed coat. The embryo then looks like a walking stick. Bythis point, the suspensor is degenerating.

The Arabidopsis LEAFY COTYLEDON1 (LEC1) gene, firstidentified by a mutant with leaflike cotyledons (Meinke1994), is necessary to maintain the suspensor early in devel-opment, to specify cotyledeon identity, to initiate matura-tion, and to prevent early germination of the seed. LEC1belongs to a gene class that is unique among the embryo-genesis genes in acting throughout the course of embryodevelopment (Harada 2001; Kwong et al. 2003).

MERISTEM ESTABLISHMENT The shoot apical meristem androot apical meristem are clusters of stem cells that will per-

638 CHAPTER 20

Concentrationof PIN1 auxinefflux carriers

Auxin-inducedgene expression

Auxin-inducedgene expression

(A) (B) Wild type (C) PID overexpression

Cells accumulating auxin

Auxin transport via PIN familyauxin efflux carriers

Cells producing auxin

Embryo

Suspensor

Embryo

No PIN 1auxin effluxcarriers

Suspensor

FIGURE 20.16 An auxin gradient specifies the shoot-root axis.(A) A family of PIN auxin efflux carriers are responsible for theearly apical flow of auxin in the embryo and the shift to basalauxin flow as the apical end of the globular stage embryobegins to produce auxin. (B,C) The PID protein kinase localizesthe PIN auxin efflux carriers. (B) In wild-type globular embryos,PID expression leads to the accumulation of PIN1 at the basalend of the embryo. When an auxin-response gene is fused withGFP and expressed, basal accumulation and normal embryonicdevelopment occur. (C) Ectopic overexpression of the PID gene(induced by fusion to a viral promoter) eliminates basal accumu-lation of PIN1. The GFP-labeled auxin response gene is thenexpressed throughout the embryo proper rather than in thebasal portion of the embryo.

*Mendel’s famous wrinkled-seed mutant (the rugosus or r allele)

has a defect in a starch branching enzyme that affects starch, lipid,

and protein biosynthesis in the seed and leads to defective cotyle-

dons (Bhattacharyya et al. 1990).



sist in the postembryonic plant and give rise to most of thesporophyte body (see Jurgens 2001 for a review of apical-basal pattern formation). The root meristem is partiallyderived from the hypophysis in some species (see Figure20.15B). All other parts of the sporophyte body are derivedfrom the embryo proper.

Genetic evidence indicates that the formation of theshoot and root meristems is regulated independently. Froman evolutionary perspective, this is not surprising. One ofthe major adaptations to terrestrial life involved the evo-lution of a root system.* This independence is demonstrat-ed by the dek23 maize mutant and the shootmeristemless(STM) mutant of Arabidopsis, both of which form a rootmeristem but fail to initiate a shoot meristem (Clark andSheridan 1986; Barton and Poethig 1993). The STM gene,which has a homeodomain, is expressed in the late globu-lar stage, before cotyledons form. Genes have also beenidentified that specifically affect the development of theroot axis during embryogenesis. Mutations of the HOBBITgene in Arabidopsis, for example, affect the hypophysisderivatives and eliminate root meristem function (Willem-son et al. 1998). While it is clear that root and shoot devel-opmental programs are different, what triggers root or

shoot development is less tractable. The TOPLESS gene inArabidopsis may provide some clues. A single gene muta-tion has been identified that converts a shoot into a root,but how the wild-type gene functions is still a puzzle (Longet al. 2002).

The shoot apical meristem will initiate leaves after ger-mination and, ultimately, the transition to reproductivedevelopment. In Arabidopsis, the cotyledons are producedfrom general embryonic tissue, not from the shoot meris-tem (Barton and Poethig 1993). In many angiosperms, afew leaves are initiated during embryogenesis. In the caseof Arabidopsis, clonal analysis points to the presence ofleaves in the mature embryo, even though they are notmorphologically well developed (Irish and Sussex 1992).Clonal analysis has demonstrated that the cotyledons andthe first two true leaves of cotton plants are derived fromembryonic tissue rather than an organized meristem(Christianson 1986).

Clonal analysis experiments provide information on cellfates, but do not necessarily indicate whether or not cellsare determined for a particular fate. Clonal analysis hasdemonstrated that cells that divide in the wrong plane and“move” to a different tissue layer often differentiate accord-ing to their new position. Position, rather than clonal ori-gin, appears to be the critical factor in embryo pattern for-mation, suggesting some type of cell-cell communication(Laux and Jürgens 1994).

Dormancy

From the earliest stages of embryogenesis, there is a highlevel of zygotic gene expression. As the embryo reachesmaturity, there is a shift from constructing the basic bodyplan to creating a food reserve by accumulating storagecarbohydrates, proteins, and lipids. Genes coding for seedstorage proteins were among the first to be characterizedby plant molecular biologists because of the high levels ofspecific storage protein mRNAs that are present at differ-ent times in embryonic development. The high level of


N

CH2COOH

H

(A) Auxin (Indole-3-acetic acid)

(B) No Auxin present

No transcription

Aux/IAA (inhibitor)

Aux/IAA

Ubiquitin+

SCF (enzyme)

SCF

Auxin

26S proteosome

Proteindegradation

ARF (transcriptionfactor)

Promoter Auxin-induced gene

(C) Auxin present

TranscriptionARF

Promoter Auxin-induced gene

Auxin-inducedmRNA

FIGURE 20.17 Mechanism of auxin action. (A) In the absenceof auxin, auxin response factors (ARF) are in a repressed state.These genes form a heterodimer with Aux/IAA protein, whichinhibits the transcription of auxin-induced genes. (B) In the pres-ence of auxin, the enzyme SCF binds Aux/IAA, catalyzingAux/IAA ubiquitination. Ubiquitination targets Aux/IAA fordegradation by the 26S proteosome. Freed from the Aux/IAArepressor, ARF can act on its own to stimulate transcription orform a homo or heterodimer with another ARF to further modu-late gene expression.

*It should be noted, however, that the nonvascular land plants,

including the mosses, did not develop root systems.



metabolic activity in the developing embryo is fueled bycontinuous input from the parent plant into the ovule.Eventually metabolism slows, and the connection of theseed to the ovary is severed by the degeneration of theadjacent supporting sporophyte cells. The seed desiccates(loses water), and the integuments harden to form a toughseed coat. The seed has entered dormancy, officially endingembryogenesis. The embryo can persist in a dormant statefor weeks or years, a phenomenon that affords tremendoussurvival value. There are even cases where seeds foundstored in ancient archaeological sites have germinated afterthousands of years of dormancy.

Maturation leading to dormancy is the result of a pre-cisely regulated program. The viviparous mutation in maize,for example, produces genetic lesions that block dorman-cy (Steeves and Sussex 1989). The apical meristems of vivip-arous mutants behave like those of ferns, with no pausebefore producing postembryonic structures. The embryocontinues to develop, and seedlings emerge from the ker-nels on the ear attached to the parent plant (Figure 20.18).A group of plant genes have been identified that belong tothe Polycomb group, which regulates early developmentin mammals, nematodes, and insects (Preuss 1999). Thesegenes encode chromatin silencing factors, which may playan important role in seed formation.

Plant hormones are critical in dormancy, and linkingthem to genetic mechanisms is an active area of research.The hormone abscisic acid is important in maintainingdormancy in many species. Gibberellins, another class ofhormones, are important in breaking dormancy.

Germination

The postembryonic phase of plant development beginswith germination. Some dormant seeds require a periodof after-ripening, during which low-level metabolic activ-ities continue to prepare the embryo for germination. High-ly evolved interactions between the seed and its environ-ment increase the odds that the germinating seedling willsurvive to produce another generation.

Temperature, water, light, and oxygen are all key indetermining the success of germination. Stratification isthe requirement for chilling (5°C) to break dormancy insome seeds. In temperate climates, this adaptation ensuresthat germination takes place only after the winter monthshave passed. In addition, seeds have maximum germina-tion rates at moderate temperatures of 25–30°C and oftenwill not germinate at extreme temperatures. Seeds such aslettuce require light (specifically, the red light wavelengths)for germination; thus seeds will not germinate so far below

ground that they use up their food reserves before photo-synthesis is possible.

Desiccated seeds may be only 5–20 percent water. Imbi-

bition is the process by which the seed rehydrates, soak-ing up large volumes of water and swelling to many timesits original size. The radicle (primary embryonic root)emerges from the seed first to enhance water uptake; it isprotected by a root cap produced by the root apical meris-tem. Water is essential for metabolic activity, but so is oxy-gen; a seed sitting in a glass of water will not survive. Somespecies have such hard protective seed coats that they mustbe scarified (scratched or etched) before water and oxy-gen can cross the barrier. Scarification can occur when theseed is exposed to the weather and other natural elementsover time, or by its exposure to acid as the seed passesthrough the gut of a frugivore (fruit eater). The frugivorethus prepares the seed for germination, as well as dispers-ing it to a site where germination can take place.

During germination, the plant draws on the nutrientreserves in the endosperm or cotyledons. Interactionsbetween the embryo and endosperm in monocots use gib-berellin as a signal to trigger the breakdown of starch intosugar. As the shoot reaches the surface, the differentiationof chloroplasts is triggered by light. Seedlings that germi-nate in the dark have long, spindly stems and do not pro-duce chlorophyll. This environmental response allowsplants to use their limited resources to reach the soil sur-face, where photosynthesis will be productive.

640 CHAPTER 20

FIGURE 20.18 Viviparous maize mutant. Each kernel on thismaize ear contains an embryo. Viviparous embryos do not gothrough a dormant phase, but begin germinating while still onthe ear.



Vegetative Growth

When the shoot emerges from the soil, most of the sporo-phyte body plan remains to be elaborated. Figure 20.19shows the basic parts of the mature sporophyte plant,which will emerge from meristems.

Meristems

As has been mentioned, meristems are clusters of cells thatallow the basic body pattern established during embryoge-nesis to be reiterated and extended after germination.Meristematic cells are similar to stem cells in animals.* Theydivide to give rise to one daughter cell that continues to bemeristematic and another that differentiates. Meristems fallinto three categories: apical, lateral, and intercalary.

Apical meristems occur at the growing shoot and roottips. Root apical meristems produce the root cap, whichconsists of lubricated cells that are sloughed off as themeristem is pushed through the soil by cell division and

elongation in more proximal cells. The root apical meris-tem also gives rise to daughter cells that produce the threetissue systems of the root (Figure 20.20A). New root api-cal meristems are initiated from tissue within the core ofthe root and emerge through the ground tissue and der-mal tissue. Root meristems can also be derived secondari-ly from the stem of the plant; in the case of maize, this isthe major source of root mass.

The shoot apical meristem produces stems, leaves, andreproductive structures. In addition to the shoot apicalmeristem initiated during embryogenesis, axillary shootapical meristems (axillary buds; Figure 20.20B) derivedfrom the original one form in the axils (the angles betweenleaf and stem). Unlike new root meristems, these arise fromthe surface layers of the meristem.


Root apicalmeristem

Epicotyl

CotyledonsHypocotyl

Internode

Vegetativeaxillary bud

Leafblade

Axillaryflower

Terminalflower

Inflorescence

Node

*The similarities between plant meristem cells and animal stem

cells may extend to the molecular level, indicating that stem cells

existed before plants and animals pursued separate phylogenetic

pathways. Homology has been found between genes required for

plant meristems to persist and genes expressed in Drosophila germ

line stem cells (Cox et al. 1998).

FIGURE 20.19 Morphology of a generalized angiospermsporophyte.

(A) Root meristems

Root hairs

Protoderm

Groundtissue

Procambium

Lateral root

(B) Shoot meristems

Leafprimordium

Leafprimordium

Protoderm

Groundtissue

Apical meristem

Procambium

Pericycle

Apicalmeristem

Root cap

Axillarymeristem

FIGURE 20.20 Both shoots and roots develop from apicalmeristems, with undifferentiated cells clustered at their tips. (A) The root meristem is protected by the root cap as it pushesthrough the soil. Lateral roots are derived from pericycle cellsdeep within the root. (B) The lateral organs of the shoot (leavesand axillary branches) have a superficial origin in the shoot api-cal meristem.



Angiosperm shoot apical meristems are composed ofup to three layers of cells (labeled L1, L2, and L3) on theplant surface (Figure 20.21A). One way of investigating thecontributions of different layers to plant structure is by con-structing chimeras (Figure 20.21B). Plant chimeras are com-posed of layers having distinct genotypes with discerniblemarkers. When L2, for example, has a different genotypethan L1 or L3, all pollen will have the L2 genotype, indi-cating that pollen is derived from L2. Chimeras have alsobeen used to demonstrate classical induction in plants, inwhich (as in animal development) one layer influences thedevelopmental pathway of an adjacent layer.

The size of the shoot apical meristem is precisely con-trolled by intercellular signals, most likely between layersof the meristem (see Bäurle and Laux 2003). Mutations inthe Arabidopsis CLAVATA (CLV) genes, for example, lead toincreased meristem size and the production of extraorgans. CLV1, CLV2, and CLV3 all limit the number ofundifferentiated, stem cells in both vegetative and floralmeristems (Figure 20.22). CLV1 is a serenine-threoninekinase that, along with the receptor-like transmembraneCLV2 protein, form a receptor for CLV3, which is localizedin the extracellular space between cell layers of the meri-stem (Clark et al. 1997; Jeong et al. 1999; Rojo et al. 2002).Reddy and Meyerowitz (2005) have demonstrated thatCLV3 restricts its own domain of expression to the centralzone by preventing the differentiation of surrounding cells.POLTERGEIST (POL) and WUSCHEL (WUS) are redun-dant in function and appear to keep genes in an undiffer-entiatied state (Yu et al. 2000). WUS specifies stem cell iden-

tity in the cells positioned above,and the CLV signaling pathway isproposed to negatively feed backon WUS expression to control thesize of the meristem. It is possiblethat POL expression is a target inCLV1 signal transduction. STM, likePOL and WUS, plays an importantrole in maintaining an undifferenti-ated population of meristematiccells, and STM may positively reg-ulate WUS (Clark and Schiefelbein1997). The interactions of these geneproducts balance the rate of celldivision (which enlarges the meri-stem) and the rate of cell differenti-ation in the periphery of the meri-stem (which decreases meristemsize) (Meyerowitz 1997).

Lateral meristems are cylindri-cal meristems found in shoots androots that result in secondarygrowth (an increase in stem androot girth by the production of vas-cular tissues). Monocots do nothave lateral meristems, but oftenhave intercalary meristems insert-

ed in their stems between mature tissues. The poppingsound you can hear in a cornfield on a summer night iscaused by the rapid increase in stem length due to inter-calary meristems.

642 CHAPTER 20

(B)

L1

Apical meristem

L2L3

(A)

FIGURE 20.21 Organization of the shoot apical meristem. (A) Angiosperm meristemshave two or three outer layers of cells that are histologically distinct (L1, L2, and L3). Whilecells in certain layers tend to have certain fates, they are not necessarily committed tothose fates. If a cell is shifted to a new layer, it generally develops like the other cells inthat layer. (B) The fates of the cell layers can be seen in a chimeric tobacco plant. One por-tion of the meristem contains three layers of wild-type cells, while the other portion hasan L2 that lacks chlorophyll. This section of the meristem has given rise to the variegatedleaves. In wild-type plants, the L1 layer always lacks chlorophyll (except in guard cells),but in this plant the L2, too, is genetically unable to produce chlorophyll; the L3 remainsgreen. The L3 does not contribute to the outer edges of leaves, which is why they appearwhite in this plant. (Photograph courtesy of M. Marcotrigiano.)

STM STM

Stem cellpopulation

Celldifferentiation

Celldifferentiation

CLV1/2

WUS

L1 L2 L3

CLV3CLV3

FIGURE 20.22 The WUS and STM proteins act to keep meris-tem cells in an undifferentiated state, while the products of theCLAVATA genes CLV1, CLV2, and CLV3 all limit the number ofundifferentiated meristem cells. The presence of WUS indirectlyinduces expression (upward arrow) of CLV3 in L1 and L2. CLV3 isa ligand that binds to the CLV1/CLV2 receptor in L3 cells. Thisbinding triggers a negative feedback signal cascade thatinhibits WUS expression and regulates the number of undiffer-entiated cells in the meristem, thus counterbalancing the rolesof STM and WUS in keeping cells undifferentiated and dividing.



Root development

Radial and axial patterning in roots begins during embryo-genesis and continues throughout development as the pri-mary root grows and lateral roots emerge from the pericy-cle cells deep within the root. Both clonal analyses and laserablation experiments that elilminate single cells havedemonstrated that cells are plastic, and that position is theprimary determinant of fate in early root development.Analyses of mutations in root radial organization haverevealed genes with layer-specific activity (Scheres et al.1995; Scheres and Heidstra 1999). We will illustrate thesefindings by looking at two Arabidopsis genes that regulateground tissue fate.

In wild-type Arabidopsis, there are two layers of rootground tissue. The outer layer becomes the cortex while theinner layer becomes the endodermis, which forms a tubearound the vascular tissue core. The SCARECROW (SCR)and SHORT-ROOT (SHR) genes have mutant phenotypeswith only a single layer of root ground tissue (Benfey et al.1993). The SCR gene is necessary for an asymmetrical celldivision in the initial layer of cells, yielding a smaller endo-dermal cell and a larger cortex cell (Figure 20.23). The scrmutant expresses markers for both cortex and endodermalcells, indicating that differentiation progresses in theabsence of cell division (Di Laurenzio et al. 1996). SHR isresponsible for endodermal cell specification. Cells in theshr mutant do not develop endodermal features.

Axial patterning in roots may be morphogen-depend-ent, paralleling some aspects of animal development. Avariety of experiments have established that the distribu-tion of the plant hormone auxin organizes the axial pat-tern. A peak in auxin concentration at the root tip must beperceived for normal axial patterning (Sabatini et al. 1999;Ueda et al. 2005).

As discussed earlier, distinct genes specifying root andshoot meristem formation have been identified; however,root and shoot development may share common groups ofgenes that regulate cell fate and patterning (Benfey 1999).This appears to be the case for the SCR and SHR genes. Inthe shoot, these genes are necessary for the normal gravit-ropic response, which is dependent on normal endodermisformation (a defect in mutants of both genes; see Figure20.29C). It is important to keep in mind that there are anumber of steps between establishment of the basic patternand elaboration of that pattern into anatomical and mor-phological structure. Uncovering the underlying controlmechanisms is likely to be the most productive strategy inunderstanding how roots and shoots develop.

Shoot development

The unique aboveground architectures of different plantspecies have their origins in shoot meristems. Shoot archi-tecture is affected by the amount of axillary bud outgrowth.Branching patterns are regulated by the shoot tip—a phe-nomenon called apical dominance—and plant hormones


(C) Shoot(B) Root

(D) Wild type (E) scr mutant

(F) shr mutant

EnEn

EpEpC

V

P

M V

EpEp

P

M

V

EpEp

P

En

EpC

V

P

M V

Ep

P

M

V

Ep

P

(A)

Cortex/endodermal

initial

Cortex/endodermal

initial

Cortex/endodermal

initial

Daughtercell

Cortexcell

Endo-dermal

cell

Positional information

FIGURE 20.23 SCR and SHRregulate endodermal differ-entiation in root radial devel-opment. (A) Diagram of nor-mal cell division yieldingcortical and endodermal cells.SCR regulates this asymmetri-cal cell division. (B,C) SCRexpression in root and shoot.The SCR promoter is linked tothe gene for GFP (green fluo-rescent protein). (D–F) Cross sections of primary roots of (D)wild-type Arabidopsis, (E) scr mutant, and (F) shr mutant. Ep, epi-dermis; C, cortex; En, endodermis; M, mutant layer; P, pericycle;V, vascular tissue; St, stele. (A after Scheres and Heidstra 1999;B–F photographs courtesy of P. Benfey.)



appear to be the regulating factors. Auxin is produced byyoung leaves and transported toward the base of the leaf.It can suppress the outgrowth of axillary buds. Grazingand flowering often release buds from apical dominance,at which time branching occurs. Cytokinins can alsorelease buds from apical dominance. Axillary buds can ini-tiate their own axillary buds, so branching patterns can getquite complex. Branching patterns can be regulated byenvironmental signals so that an expansive canopy in anopen area maximizes light capture. Asymmetrical treecrowns form when two trees grow very close to each other.In addition to its environmental plasticity, shoot architec-ture is genetically regulated. In several species, genes havenow been identified that regulate branching patterns.

Leaf primordia (clusters of cells that will form leaves)are initiated at the periphery of the shoot meristem (seeFigure 20.20). The union of a leaf and the stem is called anode, and stem tissue between nodes is called an intern-

ode (see Figure 20.19). In a simplistic sense, the maturesporophyte is created by stacking node/internode unitstogether. Phyllotaxy, the positioning of leaves on the stem,involves communication among existing and newly form-ing leaf primordia. Leaves may be arranged in various pat-terns, including a spiral, 180º alternation of single leaves,pairs, and whorls of three or more leaves at a node (Jeanand Barabé 1998). Experimentation has revealed a numberof mechanisms for maintaining geometrically regular spac-ing of leaves on a plant, including chemical and physicalinteractions of new leaf primordia with the shoot apex andwith existing primordia (Steeves and Sussex 1989).

It is not clear how a specific pattern of phyllotaxy getsstarted. Descriptive mathematical models can replicate theobserved patterns, but reveal nothing about the mecha-nism. Biophysical models (e.g., of the effects of stress orstrain on deposition of cell wall material, which affects celldivision and elongation) attempt to bridge this gap. Devel-opmental genetics approaches are promising, but few phyl-lotactic mutants have been identified.* Currently there ismuch interest in the role of local auxin maxima and mini-ma in determining where the next leaf primordium willform on a meristem. The PIN gene family that plays animportant role in embryo axis formation has also beenimplicated in phyllotaxy because of the correlation betweenlocalization of PIN auxin efflux carriers and primordia sit-ing (Fleming 2005).

Leaf development

Leaf development includes the cells’ commitment tobecome a leaf; establishment of the leaf axes; and morpho-genesis, which gives rise to a tremendous diversity of leaf

shapes. Culture experiments have assessed when leaf pri-mordia become determined for leaf development. Researchon ferns and angiosperms indicates that the youngest vis-ible leaf primordia are not determined to make a leaf;rather, these young primordia can develop as shoots in cul-ture (Steeves 1966; Smith 1984). The programming for leafdevelopment occurs later. The radial symmetry of the leafprimordium becomes dorsal-ventral, or flattened, in allleaves. Two other axes, the proximal-distal and lateral axes,are also established.

The unique shapes of leaves result from regulation ofcell division and cell expansion as the leaf blade develops.There are some cases in which selective cell death (apop-tosis) is involved in the shaping of a leaf, but differentialcell growth appears to be a more common mechanism (Gif-ford and Foster 1989).

DORSAL-VENTRAL PATTERNING IN LEAVES Plant biologists referto the surface of the leaf that is closest to the stem as theadaxial side and the more distant surface is the abaxial side.As the leaf begins to form, the Arabidopsis genes PHABU-LOSA (PHB) and PHAVOLUTA (PHV) initially have uni-formly expressed RNA throughout the primordium (Fig-ure 20.24A). The PHB and PHV proteins are postulated tobe receptors for an adaxial signal, which leads to the accu-mulation of PHB and PHV on the adaxial leaf surface(McConnell et al. 2001; Byrne 2005). Exclusion of PHB andPHV from the abaxial side is caused by microRNA bindingto the transcripts of the two genes, which leads to theirdegradation (Kidner and Martienssen 2005). In addition,the PHB- and PHV-specific microRNAs appear to increasemethylation of their complementary PHB and PHV DNAsequences, suppressing transcription (Bao et al. 2004). Dom-inant gain-of-function mutants have disrupted microRNAbinding sites such that these genes are expressed on bothsides of the primordium, leading to a leaf with two adaxialsides. The sequences of PHB and PHV make it likely thatthey are activated by a lipid ligand and that this activationis followed by the development of adaxial cells.

KANADI (KAN) genes initiate abaxial cell differentia-tion in Arabidopsis (Kerstetter et al. 2001). In situ hybridiza-tion shows that KAN is transiently expressed on the abax-ial side of cotyledons, leaves, and initiating floral organprimordia (Figure 20.24B). KAN contains a GARP domainfound in transcription factors. KAN and PHB/PHV mutual-ly suppress each other to maintain abaxial and adaxialfates, respectively.

Abaxial fate also appears to be specified by three mem-bers of the YABBY gene family (Siegfried et al. 1999). Theexact mechanism of YABBY genes in leaf polarity isunknown, although promoter deletion experiments revealthat FILAMENTOUS, a gene in the YABBY family, is active-ly excluded from the adaxial side of the primordium. KANand YABBY likely have redundant functions. KAN geneexpression is restricted to the abaxial side by PHB andPHV, while KAN restricts PHB and PHV expression to theadaxial side (Figure 20.24C).

644 CHAPTER 20

*One candidate phyllotactic mutant is the terminal ear mutant

in maize, which has irregular phyllotaxy. The wild-type gene is

expressed in a horseshoe-shaped region, with a gap where the leaf

will be initiated (Veit et al. 1998). The plane of the horseshoe is per-

pendicular to the axis of the stem.



Leaves fall into two categories, simple and compound(Figure 20.25; see review by Sinha 1999). There is muchvariety in simple leaf shape, from smooth-edged leaves todeeply lobed oak leaves. Compound leaves are composedof individual leaflets (and sometimes tendrils) rather thana single leaf blade. Whether simple and compound leavesdevelop by the same mechanism is an open question. Oneperspective is that compound leaves are highly lobed sim-ple leaves. An alternative perspective is that compoundleaves are modified shoots. The ancestral state for seedplants is believed to be compound, but for angiosperms itis simple. Compound leaves have arisen multiple times inthe angiosperms, and it is not clear if these are reversions tothe ancestral state.

Developmental genetic approaches are being applied toleaf morphogenesis. The Class I KNOX genes are home-obox genes that include STM and the KNOTTED 1 (KN1)

gene in maize. Gain-of-function mutations of KN1 causemeristem-like bumps to form on maize leaves. In wild-typeplants, this gene is expressed in meristems. KNOX genesstimulate meristem initiation and growth. Although someYABBY genes specify abaxial leaf identity (see Figure20.24), others function by downregulating KNOX genes,


Leafprimordium

(A) Adaxialsurface

Abaxialsurface

Adaxialdevelopment

Adaxial signal

PHB, PHV

Micro RNAs bind PHB and PHV RNA

KAN

YABBY

(B)

Abaxialdevelopment

(C)

Normaldevelopment

Abaxial surfa

ce

Adaxial surface

FIGURE 20.24 Patterning of adaxial and abaxial leaf surfacesof Arabidopsis. (A) PHB and PHV proteins are uniformly distrib-uted throughout the leaf primordium. MicroRNAs expressed onthe abaxial side specifically degrade PHB and PHV, leading todegradation by DICER and blocking translation. In addition,microRNAs increase methylation of the complementary regionsin PHB and PHV DNA, suppressing transcription. (B) Transientexpression of KAN on the abaxial side of the leaf leads to theexpression of YABBY genes, which in turn lead to the transcrip-tion of other abaxial genes. (C) PHB and PHV block KAN adaxialactivity. Expression of PHB, PHV, and KAN is transient and occursearly in leaf development.

Tendril

LeafletRachisPetiole

Petiole Blade

Vein

Midrib

Stipule

COMPOUND LEAF

SIMPLE LEAF

FIGURE 20.25 Simple and compound leaves.



thereby restricting where meristems form (Kumaran et al.2002). If too much KN1 is present, YABBY is insufficient tocontrol KN1 expression.

When KN1, or the tomato homologue LeT6, has its pro-moter replaced with a promoter from cauliflower mosaicvirus and is inserted into the genome of tomato, the geneis expressed at high levels throughout the plant, and theleaves become “super compound” (Figure 20.26; Harevenet al. 1996; Janssen et al. 1998). Simple leaves become morelobed (but not compound) in response to overexpressionof KN1, consistent with the hypothesis that compoundleaves may be an extreme case of lobing in simple leaves(Jackson 1996). The role of KN1 in shoot meristem and leaf

development, however, is consistent with the hypothesisthat compound leaves are modified shoots. Looking at pat-terns of KN1 expression over a broad range of angiospermtaxa, Bharathan et al. (2002) demonstrated that KN1 expres-sion is associated with compound leaf primordia, but is notpresent in the developing leaf primordia of simple leaves(Figure 20.27). These data support the conclusion that com-pound leaves maintain shootlike activity, at least for sometime. How KN1 works in meristems and in leaf primordiais an area of active research. In tomato leaf, KN1 causes a

646 CHAPTER 20

(A) (B)

(C)

FIGURE 20.26 Overexpression of Class 1 KNOX genes intomato. The photograph shows the single leaves of (A) a wild-type plant, (B) a mouse ears mutant, with increased leaf com-plexity, and (C) a transgenic plant that uses a viral promoter tooverexpress the tomato homologue (LeT6) of the KN1 genefrom maize. (Photographs courtesy of N. Sinha.)

(C) (C)

(F) (F)

(I) (I)

(C)

(F)

(I)

(A) (B)

(D) (E)

(G) (H)

LPLPLP

LPLPLPMM

LP

FIGURE 20.27 KNOX is known to occur in leaf primordia inspecies making complex leaves. Here we show that species withsimple leaves can also express KNOX in leaf primordia that startout complex but undergo secondary alteration to become sim-ple. (A) Coffee simple leaf. (B) SEM of primordia (arrow) showingearly complexity. (C) Expression of KNOX genes in these earlyleaves. (D) Anise simple leaf. (E). SEM showing early complexity.(F) KNOX expression in these primordia. (G) Amborella showing asimple leaf that is simple throughout development (H) and hasno KNOX expression (I). LP, leaf primordia; M, meristem. (FromBharathan et al. 2002; photographs courtesy of N. Sinha.)



decrease in the expression of a gene needed for the produc-tion of the plant hormone gibberellic acid (Hay et al. 2002).

A second gene, LEAFY, is essential for the transitionfrom vegetative to reproductive development and alsoappears to play a role in compound leaf development.LEAFY was identified in Arabidopsis and snapdragons (inwhich it is called FLORICAULA) and has homologues inother angiosperms. The pea homologue, UNIFOLIATA, hasa mutant phenotype in which compound leaves arereduced to simple leaves (Hofer and Ellis 1998). This find-ing is also indicative of a regulatory relationship betweenshoots and compound leaves. In an intriguing evolution-ary twist, UNI appears to have taken on the role of KN1genes in leaf development in peas (Sinha 2002). It appearsto have been co-opted from its role in flowering in thishighly derived legume.

In some compound leaves, developmental decisionsabout leaf versus tendril formation are also made. Muta-tions of two leaf-shape genes can individually and in sumdramatically alter the morphology of the compound pealeaf. The acacia (tl) mutant converts tendrils to leaflets; afil-ia (af) converts leaflet to tendrils (Marx 1987). The af tl dou-ble mutant has a complex architecture and resembles aparsley leaf (Figure 20.28).

At a more microscopic level, the patterning of stomata(openings for gas and water exchange) and trichomes(hairs) across the leaf is also being investigated. In mono-cots, the stomata form in parallel files, while in dicots thedistribution appears more random. In both cases, the pat-terns appear to maximize the evenness of stomata distri-bution. Genetic analysis is providing insight into the mech-anisms regulating this distribution. A common gene groupappears to be working in both shoots and roots, affectingthe distribution pattern of both trichomes and root hairs(Benfey 1999).

The Vegetative-to-Reproductive Transition

Unlike most animal systems, in which the germ line is setaside during early embryogenesis, the germ line in plantsis established only after the transition from vegetative toreproductive development (flowering). The vegetative andreproductive structures of the shoot are all derived fromthe shoot meristem formed during embryogenesis. Clon-al analysis indicates that no cells are set aside in the shootmeristem of the embryo to be used solely in the creationof reproductive structures (McDaniel and Poethig 1988).In maize, irradiating seeds causes changes in the pigmen-tation of some cells. These seeds give rise to plants thathave visually distinguishable sectors descended from themutant cells. Such sectors may extend from the vegetativeportion of the plant into the reproductive regions (Figure20.29), indicating that maize embryos do not have distinctreproductive compartments.

Maximal reproductive success in angiosperms dependson the timing of flowering and on balancing the numberof seeds produced with the resources allocated to individ-ual seeds. As in animals, different strategies work best fordifferent organisms in different environments. There is agreat diversity of flowering patterns among the over300,000 angiosperm species, yet there appears to be anunderlying evolutionary conservation of flowering genesand common patterns of flowering regulation.

A simplistic explanation of the flowering process is thata signal from the leaves moves to the shoot apex andinduces flowering. In some species, this flowering signalis a response to environmental conditions. The develop-mental pathways leading to flowering are regulated atnumerous control points in different plant organs (roots,cotyledons, leaves, and shoot apices) in various species,


(A) (B)

(C) (D)

FIGURE 20.28 Leaf morphology mutantsin peas. (A) Wild-type pea plant. (B) The tlmutant, in which tendrils are converted toleaflets. (C) The af mutant, in which leafletsare converted to tendrils. (D) An af tl doublemutant, which results in a “parsley leaf” phe-notype. (Photographs courtesy of S. Singer.)



resulting in a diversity of flowering times and reproduc-tive architectures. The nature of the flowering signal, how-ever, remains unknown. There is now evidence that RNAwith developmental functions can move through thephloem, and the role of very small pieces of RNA in signaling developmental mechanisms has recently beenrecognized.

Juvenility

Some plants, especially woody perennials, go through ajuvenile phase during which the plant cannot producereproductive structures even if all the appropriate environ-mental signals are present (Lawson and Poethig 1995). Thetransition from the juvenile to the adult stage may requirethe acquisition of competence by the leaves or meristem to

respond to an internal or external signal (McDaniel et al.1992; Singer et al. 1992; Huala and Sussex 1993). Even inherbaceous plants, there is a phase change from juvenile toadult vegetative growth. For example, the EARLY PHASECHANGE (EPC) gene in maize is required to maintain thejuvenile state (Vega et al. 2002). Mutant epc plants flowerearly because they have fewer juvenile leaves which are dis-tinguished from adult leaves by the presence of wax and theabsence of hairs. These epc mutants, however, have the samenumber of adult leaves as wild-type plants. Juvenile traitsare not expressed in the absence of EPC. The mechanismsof phase change genes are just beginning to be understood.The Arabidopsis juvenility gene HASTY (HST), is necessaryfor microRNA processing and export from the nucleus. Loss-of-function mutants undergo early phase change becausemicroRNAs are trapped in the nucleus (Park et al. 2005).

648 CHAPTER 20

Tassel(male flowers)

SECTOR ALength =4 internodes

Width =1/8 stemcircumference

SECTOR BLength =2 internodes

Width =1/24 stemcircumference

Plant A Plant B

FIGURE 20.29 Clonal analysis canbe used to construct a fate map of ashoot apical meristem in maize.Seeds that are heterozygous for cer-tain pigment genes (anthocyanins)are irradiated so that the dominantallele is lost in a few cells (a chanceoccurrence). All cells derived fromthe somatic mutant will be visuallydistinct from the nonmutant cells.Plants A and B have mutant sectorsthat reveal the fate of cells in theshoot meristem of the seed. Themutant sector in A includes bothvegetative and reproductive (tassel) internodes.Thus there is no distinct developmental compart-ment that forms the tassel. The mutant sector in Ais longer and wider than the mutant sector in B.This indicates that more cells were set aside tocontribute to the lower than to the upper intern-odes in the shoot meristem in the seed. The actualnumber of cells can be calculated by taking thereciprocal of the fraction of the stem circumfer-ence the sector occupies. Sector A contributes to1/8 of the circumference of the stem; thus 8 cellswere fated to contribute to these internodes in theseed meristem. Sector B is only 1/24 of the stemcircumference; thus 24 cells were fated to con-tribute to these internodes. In this example, onlycells derived from the L1 are being analyzed. It isalso important to consider the possible contribu-tions of the L2 and L3 cell layers of the shootmeristem. (Data from McDaniel and Poethig 1988;photographs courtesy of C. McDaniel.)



Floral signals

Grafting and organ culture experiments, mutant analyses,and molecular analyses give us a framework for describ-ing the reproductive transition in plants. Grafting experi-ments have identified the sources of signals that promoteor inhibit flowering and have provided information on thedevelopmental acquisition of meristem competence torespond to these signals (Lang et al. 1977; Singer et al. 1992;Reid et al. 1996). Analyses of mutants and molecular char-acterization of genes are yielding information on themechanics of these signal-response mechanisms (Levy andDean 1998; Hempel et al. 2000; Hecht et al. 2005).

Leaves produce a graft-transmissible substance thatinduces flowering. In some species, this signal is producedonly under specific photoperiods (day lengths), whileother species are day-neutral and will flower under anyphotoperiod (Zeevaart 1984). Not all leaves may be com-petent to perceive or pass on photoperiodic signals. Thephytochrome pigments transduce these signals from theexternal environment. The structure of phytochrome ismodified by red and far-red light, and these changes caninitiate a cascade of events leading to the production ofeither floral promoter or floral inhibitor (Deng and Quail

1999). Leaves, cotyledons, and roots have been identifiedas sources of floral inhibitors in some species (McDaniel etal. 1992; Reid et al. 1996). A critical balance betweeninhibitor and promoter is needed for the reproductive tran-sition. In addition, vernalization, a period of chilling, canenhance the competence of shoots and leaves to perceiveor produce a flowering signal.

The “black box” between environmental signals and theproduction of a flower is vanishing rapidly, especially inthe model plant Arabidopsis (Searle and Coupland 2004;Achard et al. 2006). The signaling pathways from light viadifferent phytochromes to key flowering genes are beingelucidated (Figure 20.30A). CONSTANS (CO) responds today length, promoting flowering under long-day condi-tions. CO activates transcription of a gene called FLOW-ERING LOCUS T (FT). FT is transcribed only in the leaves,but the transcript travels through the phloem (transporttissue) from the leaf to the shoot (see Blázquez 2005). FT islikely translated when it arrives at the shoot meristem. TheFT protein interacts with the transcription factor FD, whichis expressed only in the shoot tip. Together these two pro-teins are responsible for the expression of regulatory genesthat lead to the production of flowers.


Light

(A)

(B)

CO FT

FTtranscript

Phase change(HST, EPC)

Floral organ identitygenes (ABC genes)

TFL1

FT+

FD

FT+

FD

LFY/API

Flowers

TFL1

Flowers

Floral meristemidentity genes

Phase changegenes

Floral organidentity genes

Juvenilevegetative

growth

Adultvegetative

growth

Inflorescencegrowth

Flowerformation

Photoperiodicpathway

Autonomouspathway

Vernalizationpathway

Giberellinpathway

Possible florigen?

Floral meristemidentity genes

FIGURE 20.30 The vegetative-to-reproductivetransition. (A) Signals promoting or inhibitingflowering can move from the roots, cotyledons,or leaves to the shoot apex, where meristemcompetence determines whether or not theplant will respond to the signals. Leaves may alsoneed to develop competence to respond to envi-ronmental signals before they can produce floralpromoters. Leaves and meristems that are com-petent to respond to flowering signals haveundergone a juvenile-to-adult phase change. (B)Internal and external factors regulate whether ameristem produces vegetative or reproductivestructures. Not all of the regulatory mechanismsshown are used in all species, and some speciesflower independently of external environmentalsignals.



Is FT the long-sought florigen? Possibly, but we don’tknow if other proteins aid in the movement of FT, or ifother transcripts must also move through the phloembefore flowering can occur.

Some of the genes that CONSTANS protein activates inaddition to FT are also activated by transcription factorsin other, non-light-dependent, flowering pathways. Mol-ecular explanations are revealing redundant pathways thatensure that flowering will occur. In Arabidopsis, four sepa-rable pathways leading to flowering are being elucidated.Light-dependent, vernalization-dependent, gibberellin,and autonomous pathways that regulate the floral transi-tion have been genetically dissected (Figure 20.30B). Theavailability of the Arabidopsis genome sequence now makesit possible to search on a broader scale for classes of genesthat regulate critical events such as time of flowering (Rat-cliffe and Riechmann 2002). Schmid and colleagues (2005)created a developmental gene expression profile of almostall the Arabidopsis genes using microarrays. The currentchallenge is to integrate the vast amount of expression datawith functional analysis.

One promising approach is the search for quantitativetrait loci (QTL) that control responses to environmentaland hormonal factors. In Arabidopsis, two lines, including awild accession with natural variation in light and hormoneresponses, were used to identify new genes (Borevitz et al.2002). Once QTL are mapped, the Arabidopsis genome mapmakes it more likely that researchers will be able to asso-ciate function with a specific gene.

Inflorescence development

The ancestral angiosperm is believed tohave formed a terminal flower directlyfrom the terminal shoot apex (Stebbins1974). In modern angiosperms, a vari-ety of flowering patterns exist in whichthe terminal shoot apex is indetermi-nate, but axillary buds produce flowers.This observation introduces an interme-diate step into the reproductive process:the transition of a vegetative meristemto an inflorescence meristem, whichinitiates axillary meristems that can pro-duce floral organs, but does not directlyproduce floral parts itself. The inflores-cence is the reproductive backbone(stem) that displays the flowers (see Fig-ure 20.19).

The inflorescence meristem probably arises through theaction of a gene that suppresses terminal flower formation.The CENTRORADIALUS (CEN) gene in snapdragons sup-presses terminal flower formation by suppressing expres-sion of FLORICAULA (FLO), which specifies floral meris-tem identity (Bradley et al. 1996). Curiously, the expressionof FLO is necessary for CEN to be turned on. The Arabidop-sis homolog of CEN (TERMINAL FLOWER 1 or TFL1) isexpressed during the vegetative phase of development aswell, and has the additional function of delaying the com-mitment to inflorescence development (Bradley et al. 1997).Overexpression of TFL1 in transgenic Arabidopsis extendsthe time before a terminal flower forms (Ratcliffe et al.1998). TFL1 must delay the reproductive transition.

Floral meristem identity

The next step in the reproductive process is the specifica-tion of floral meristems—those meristems that will actu-ally produce flowers (Weigel 1995). In Arabidopsis, LEAFY(LFY), APETALA 1 (AP1), and CAULIFLOWER (CAL) arefloral meristem identity genes (Figure 20.31). LFY is thehomologue of FLO in snapdragons, and its upregulationduring development is key to the transition to reproduc-tive development (Blázquez et al. 1997; Blázquez andWeigel 2000). The LEAFY transcription factor can actuallymove between cell layers in the meristem before activat-ing other flowering genes (Sessions et al. 2000). Analysisof the promoter of LFY reveals that there are separate sitesfor activation by the photoperiod pathway and theautonomous pathway. The autonomous pathway worksthrough the binding of giberellin to the LFY promoter.

650 CHAPTER 20

(A) (B)

(C) (D)

FIGURE 20.31 Floral meristem identitymutants. (A) Wild-type Arabidopsis. (B) Theleafy mutant. (C) The apetala1 mutant. (D)The leafy apetala1 double mutant. (Pho-tographs courtesy of J. Bowman.)



Expression of floral meristem identity genes is necessaryfor the transition from an inflorescence meristem to a floralmeristem. Strong lfy mutants tend to form leafy shoots inthe axils where flowers form in wild-type plants; they areunable to make the transition to floral development. If LFYis overexpressed, flowering occurs early. For example,when aspen was transformed with anLFY gene that was expressed through-out the plant, the time to floweringwas dramatically shortened from yearsto months (Weigel and Nilsson 1995).

AP1 and CAL are closely relatedand redundant genes. The cal mutantlooks like the wild-type plant, but ap1cal double mutants produce inflores-cences that look like cauliflower heads(Figure 20.32). More recently anothergene, FRUITFUL (FUL), with closesequence similarity to AP1 and CAL,has been characterized. The FRUITFULgene family is closely related to the

AP1 gene family and most likely arose as a result of geneduplication and divergence. The FUL gene is partiallyredundant to AP1 and CAL (Ferrandiz et al. 2000). CAL,however, arose only within the brassicas through an AP1gene duplication event, and appears to have accompaniedthe domestication of cauliflower and broccoli (Puruggananet al. 2000).

Floral meristem identity genes initiate a cascade of geneexpression that turns on region-specifying (cadastral)genes, which further specify pattern by initiating transcrip-tion of floral organ identity genes (Weigel 1995). SUPER-MAN (SUP) is an example of a cadastral gene in Arabidop-sis that plays a role in specifying boundaries for theexpression of organ identity genes. Three classes (A, B,and C) of organ identity genes are necessary to specify thefour whorls of floral organs (Figure 20.33; Coen andMeyerowitz 1991). They are homeotic genes (but not Hoxgenes; rather, most are members of the MADS box genefamily that had its origins before the divergence of animalsand plants) and include AP2, AGAMOUS (AG), AP3, andPISTILLATA (PI) in Arabidopsis. Class A genes (AP2) alonespecify sepal development. Class A genes and class B genes(AP3 and PI) together specify petals. Class B and class C(AG) genes are necessary for stamen formation; class C


FIGURE 20.32 Arabidopsis double mutant of ap1 and cal.Since cal alone gives a wild-type phenotype, the double mutantdemonstrates the redundancy of these two genes in the flower-ing pathway. (Photograph courtesy of J. Bowman.)

Wild-type

b

c

abc

ac

bc

ab

a

Se

Ca

Ca

Se

Se*

Pe

Ca

Ca

Ca

Se

Se

Se

Lf

Lf

Lf

Lf

Ca St Pe Se

B + C

A + B

A

Pe

Ca

Ca

St

St

Pe/St

Pe/St

Pe/St

Pe/St

C

Ca

Se

Se

FIGURE 20.33 The ABC model for floralorgan specification. Three classes ofgenes—A, B, and C—regulate organ iden-tity in flowers. The central diagram repre-sents the wild-type flower; surroundingdiagrams represent mutants that are miss-ing one or more of these gene functions(indicated by the lowercase a, b, or c). Se,sepal; Pe, petal; St, stamen; Ca, carpel;Pe/St, a hybrid petal/stamen; Lf, leaf; Se*, amodified sepal indicating that other genes(possibly ovule genes) also regulate floralorgan specification. (Model of Coen andMeyerowitz 1991.)



genes alone specify carpel formation. When all of thesehomeotic genes are not expressed in a developing flower,floral parts become leaflike (Figure 20.34). The ABC genescode for transcription factors that initiate a cascade ofevents leading to the actual production of floral parts.

While the ABC model is compelling, it is not sufficientto support the hypothesis that flowers evolved from leaves.Overexpressing the ABC genes in leaves does not producepetals or other flower parts. About a decade after the ABCmodel was proposed, a fourth class of floral organ identi-ty genes, SEPALLATA (SEP), was identified (see Jack 2001).These MADS box genes can convert a leaf into a petal whenectopically expressed in the leaf. In the absence of SEP func-tion, flowers become whorls of sepals (Figure 20.35). SEPtranscription factors form dimers with ABC or other SEPtranscription factors to initiate floral development.

In addition to the ABC and SEP genes, class D genes arenow being investigated that specifically regulate ovuledevelopment. The ovule evolved long before the otherangiosperm floral parts, and while its development is coor-

dinated with that of the carpel, one wouldexpect more ancient, independent pathwaysto exist.

Transcription of floral organ identitygenes is actually the beginning rather thanthe end of flower development. One of theamazing attributes of angiosperms is thetremendous diversity of flower phenotypes,many of which attract specific pollinators.Adaxial/abaxial asymmetry in flowers is

one contributing factor to diverse floral morphologies thatattract pollinators. Phylogenetic evidence indicates thatthis trait arose independently many times, as well as beinglost many times. The cloning of the CYCLOIDEA (CYC)gene in snapdragons has led to extensive discussion of themolecular mechanisms involved in the evolution of asym-metry (Donoghue et al. 1998; Cubas et al. 2001). In snap-dragons, CYC expression is observed on the adaxial sideof the floral primordium early in development. In cycmutants, flowers have a more radial symmetry. Did CYCevolve independently numerous times, or are there manyways to make an asymmetrical flower? Putative CYCorthologues have been identified in other species, and oneplausible explanation for the multiple origins of asymme-try is that the CYC gene was recruited for the same func-tion multiple times. The combination of developmentaland phylogenetic approaches to the study of patterning inplants promises to provide insight into the origins of themyriad morphological novelties that are found among theangiosperms.

652 CHAPTER 20

Sepal SepalPetal Petal

A A A A

B BGenes

ag (C–)

Petal StamenStamen CarpelCarpel

C C C C

B BGenes

ap2 (A–)

(A)

(B)

(C)

(D)

Sepal Petal Stamen Carpel

A A C C

B B

Flower structure

Genes

Wild-type

Sepal Sepal CarpelCarpel

A A C CGenes

ap3 and pi (B–)

Flower structure

Flower structure

Flower structure

FIGURE 20.34 Wild-type and mutant pheno-types of the Arabidopsis class A (ap2), class B(ap3, pi), and class C (ag) floral organ identitygenes. (Photographs courtesy of J. Bowman.)



Senescence

Senescence, a developmental program leading to death,is closely linked to flowering in many angiosperms. Insome species, individual flower petals senesce followingpollination; orchids, which can stay fresh for long periodsof time unless they are pollinated, are a good example.Fruit ripening (and ultimately overripening) is an exam-ple of organ senescence. Whole-plant senescence leads tothe death of the entire sporophyte generation. Monocarpic

plants flower once and then senesce. Polycarpic plants,such as the bristlecone pine, can live thousands of years(4,900 years is the current record) and flower repeatedly.In polycarpic plants, death is accidental; in monocarpicplants, it appears to be genetically programmed.

Flowers and fruits play a key role in senescence, andtheir removal can sometimes delay the process. In somelegumes, senescence can be delayed by removing thedeveloping seed—in other words, the embryo may triggersenescence in the parent plant. During flowering and fruitdevelopment, nutrients are reallocated from other parts ofthe plant to support the development of the next genera-tion. The reproductive structures become a nutrient sink,and this can lead to whole-plant senescence.


Sepal Petal Stamen Carpel

A

SEP1 SEP2

SEP3

A C C

B B

Flower structure

Genes

Wild-type(A)

Sepal

Normal Arabidopsisleaf

Ectopic expression ofSEP in Arabidopsis leaf

Sepal Sepal Sepal

A

SEP1 SEP1

SEP1

A C C

B B

Flower structure

Genes

Wild-type(B)

(C)

FIGURE 20.35 SEPALLATA (SEP) genes are a fourth class oforgan identity genes. Arabidopsis plants with the triple mutationsep1–sep 2–sep 3– produce whorls of sepals and lack the otherthree floral organs. Ectopic expression of all three SEP genes inleaves causes them to convert to petals.

1. Plants are characterized by alternation of genera-tions; that is, their life cycle includes both diploidand haploid multicellular generations.

2. Land plants have evolved mechanisms to protectembryos. Angiosperm embryos develop deeplyembedded in parent tissue. The parent tissue pro-vides nutrients and some patterning information.This evolutionary theme of an increasingly protectedembryo is shared by both plants and animals.

3. A multicellular diploid sporophyte produces hap-loid spores via meiosis. These spores divide mitoti-cally to produce a haploid gametophyte. Mitoticdivisions within the gametophyte produce thegametes. The diploid sporophyte results from thefusion of two gametes.

4. In angiosperms, the male gamete, pollen, arrives atthe style of the female gametophyte and effects fer-tilization through the pollen tube. Two sperm cellsmove through the pollen tube: one joins with theovum to form the zygote, and the other is involvedin the formation of the endosperm.

5. Early embryogenesis is characterized by the estab-lishment of the shoot-root axis and by radial pattern-ing yielding three tissue systems. Pattern emerges byregulation of planes of cell division and the direc-tions of cell expansion, since plant cells do not moveduring development.

6. As the angiosperm embryo matures, a food reserveis established. Only the rudiments of the basic bodyplan are established by the time embryogenesis ceas-es and the seed enters dormancy.

7. Pattern is elaborated during postembryonic devel-opment, when meristems construct the reiterativestructures of the plant.

8. Unlike most animals, the germ line is not set asideearly in plant development. Coordination of signal-ing among leaves, roots, and shoot meristems regu-lates the transition to the reproductive state.

9. Floral meristem identity genes and organ identitygenes enable the angiosperm flower to display atremendous amount of morphological diversity.

10. Reproduction may be followed by genetically pro-grammed senescence of the parent plant.

Plant DevelopmentSnapshotSummary



654 CHAPTER 20

Bao, N., K.-W. Lye and M. K. Barton.2004. MicroRNA binding sites in Ara-bidopsis class III HD-ZIP mRNAs arerequired for methylation of the tem-plate chromosome. Dev. Cell 7: 653–662.

Blázquez, M. A. 2005. The right timeand place for making flowers. Science309: 1024–1025.

Brownlee, C. 2004. From polarity to pat-tern: Early development of in fucoidalgae. Ann. Plant Rev. 12: 138–156.

Byrne, M. E. 2005. Networks in leafdevelopment. Curr. Opin. Plant Biol. 8: 59–66.

Dharmasiri, N., S. Dharmasiri, and M.Estelle. 2005. The F-box protein TIR1 isan auxin receptor. Nature 435: 441–445.

Fleming, A. J. 2005. Formation of pri-mordia and phyllotaxy. Curr. Opin.Plant Biol. 8: 53–58.

Friml, J., X. Yang, M. Michniewicz, D. Weijers, A. Quint, O. Tietz, R. Ben-jamins, et al. 2004. A PINOID-depend-ent binary switch in apical-basal PINpolar targeting directs auxin efflux. Science 306: 862–865.

Jack, T. 2001. Relearning our ABCs:New twists on an old model. TrendsPlant Sci. 6: 310–316.

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Website Topic 20.1: Plant Life Cycles (Originally included in the Developmental Biology, Eighth Edition Companion Website)

Plants have both multicellular haploid and multicellular diploid stages in their life cycles. Embryonic development is seen only in the diploid generation. The embryo, however, is produced by the fusion of gametes, which are formed only by the haploid generation. Understanding the relationship between the two generations is important in the study of plant development.

In plants, gametes develop in the multicellular haploid gametophyte (Greek phyton, “plant”). Fertilization gives rise to a multicellular diploid sporophyte, which produces haploid spores via meiosis. This type of life cycle is called a haplodiplontic life cycle (Figure 1). It differs from the diplontic life cycle of most animals, in which only the gametes are in the haploid state. In haplodiplontic life cycles, gametes are not the direct result of a meiotic division. Diploid sporophyte cells undergo meiosis to produce haploid spores. Each spore then goes through mitotic divisions to yield a multicellular, haploid gametophyte. Mitotic divisions within the gametophyte are required to produce the gametes. The diploid sporophyte results from the fusion of two gametes. Among the Plantae, the gametophytes and sporophytes of a species have distinct morphologies (in some algae they look alike). How a single genome can be used to create two unique morphologies is an intriguing puzzle.

Figure 1 Plants have haplodiplontic life cycles that involve mitotic divisions (resulting in multicellularity) in both the haploid and diploid generations (paths A and D). Most animals are diplontic and undergo mitosis only in the diploid generation (paths B and D). Multicellular organisms with haplontic life cycles follow paths A and C.

All plants alternate generations. There is an evolutionary trend from sporophytes that are nutritionally dependent on autotrophic (self-feeding) gametophytes to the opposite—gametophytes that are dependent on autotrophic sporophytes. This trend is exemplified by comparing the life cycles of a moss, a fern, and an angiosperm (see Figures 2-4). (Gymnosperm


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life cycles bear many similarities to those of angiosperms; the distinctions will be explored in the context of angiosperm development.)

The “leafy” moss you walk on in the woods is the gametophyte generation of that plant (Figure 2). Mosses are heterosporous, which means they make two distinct types of spores; these develop into male and female gametophytes. Male gametophytes develop reproductive structures called antheridia (singular, antheridium) that produce sperm by mitosis. Female gametophytes develop archegonia (singular, archegonium) that produce eggs by mitosis. Sperm travel to a neighboring plant via a water droplet, are chemically attracted to the entrance of the archegonium, and fertilization results.1 The embryonic sporophyte develops within the archegonium, and the mature sporophyte stays attached to the gametophyte. The sporophyte is not photosynthetic. Thus both the embryo and the mature sporophyte are nourished by the gametophyte. Meiosis within the capsule of the sporophyte yields haploid spores that are released and eventually germinate to form a male or female gametophyte.

Figure 2 Life cycle of a moss (genus Polytrichum). The sporophyte generation is dependent on the photosynthetic gametophyte for nutrition. Cells within the sporangium of the sporophyte undergo meiosis to produce male and female spores, respectively. These spores divide mitotically to produce multicellular male and female gametophytes. Differentiation of the growing tip of the gametophyte produces antheridia in males and archegonia in females. The sperm and eggs are produced mitotically in the antheridia and archegonia, respectively. Sperm are carried to the archegonia in water droplets. After fertilization, the sporophyte generation develops in the archegonium and remains attached to the female gametophyte.

Ferns follow a pattern of development similar to that of mosses, although most (but not all) ferns are homosporous. That is, the sporophyte produces only one type of spore within a structure called the sporangium (Figure 3). A single gametophyte can produce both male and female sex organs. The greatest contrast between the mosses and the ferns is that both the gametophyte and the sporophyte of the fern photosynthesize and are thus autotrophic; the shift to a dominant sporophyte generation is taking place.2


Figure 3 Life cycle of a fern (genus Polypodium). The sporophyte generation is photosynthetic and is independent of the gametophyte. The sporangia are protected by a layer of cells called the indusium. This entire structure is called a sorus. Meiosis within the sporangia yields a haploid spore. Each spore divides mitotically to produce a heart-shaped gametophyte, which differentiates both archegonia and antheridia on one individual. The gametophyte is photosynthetic and independent, although it is smaller than the sporophyte. Fertilization takes place when water is available for sperm to swim to the archegonia and fertilize the eggs. The sporophyte has vascular tissue and roots; the gametophyte does not.

At first glance, angiosperms may appear to have a diplontic life cycle because the gametophyte generation has been reduced to just a few cells (Figure 4). However, mitotic division still follows meiosis in the sporophyte, resulting in a multicellular gametophyte, which produces eggs or sperm. All of this takes place in the organ that characterizes the angiosperms: the flower. Male and female gametophytes have distinct morphologies (i.e., angiosperms are heterosporous), but the gametes they produce no longer rely on water for fertilization. Rather, wind or members of the animal kingdom deliver the male gametophyte “pollen” to the female gametophyte. Another evolutionary innovation found in the gymnosperms and angiosperms is the production of a seed coat, which adds an extra layer of protection around the embryo. A further protective layer, the fruit, is unique to the angiosperms and aids in the dispersal of the enclosed embryos by wind or animals.


Figure 4 Life cycle of an angiosperm, represented here by a pea plant (genus Pisum). The sporophyte is the dominant generation, but multicellular male and female gametophytes are produced within the flowers of the sporophyte. Cells of the microsporangium within the anther undergo meiosis to produce microspores. Subsequent mitotic divisions are limited, but the end result is a multicellular pollen grain. The megasporangium is protected by two layers of integuments and the ovary wall. Within the megasporangium, meiosis yields four megaspores—three small and one large. Only the large megaspore survives to produce the embryo sac. Fertilization occurs when the pollen germinates and the pollen tube grows toward the embryo sac. The sporophyte generation may be maintained in a dormant state, protected by the seed coat.

1Have you ever wondered why there are no moss trees? Aside from the fact that the gametophytes of mosses (and other plants) do not have the necessary structural support and transport systems to attain tree height, it would be very difficult for a sperm to swim up a tree!

2It is possible to have tree ferns, for two reasons. First, the gametophyte develops on the ground, where water can facilitate fertilization. Second, unlike mosses, the fern sporophyte has vascular tissue, which provides the support and transport system necessary to achieve substantial height.


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