meiosis in ﬂowering plants and other green organisms · meiosis in ﬂowering plants and other...

Journal of Experimental Botany, Vol. 61, No. 11, pp. 2863–2875, 2010doi:10.1093/jxb/erq191

FLOWERING NEWSLETTER REVIEW

Meiosis in flowering plants and other green organisms

C. Jill Harrison*, Elizabeth Alvey and Ian R. Henderson*

Department of Plant Sciences, University of Cambridge, Downing Street, Cambridge CB2 3EA, UK

* To whom correspondence should be addressed: E-mail: [email protected] and [email protected]

Received 17 February 2010; Revised 21 May 2010; Accepted 25 May 2010

Abstract

Sexual eukaryotes generate gametes using a specialized cell division called meiosis that serves both to halve the

number of chromosomes and to reshuffle genetic variation present in the parent. The nature and mechanism of the

meiotic cell division in plants and its effect on genetic variation are reviewed here. As flowers are the site of meiosis

and fertilization in angiosperms, meiotic control will be considered within this developmental context. Finally, we

review what is known about the control of meiosis in green algae and non-flowering land plants and discuss

evolutionary transitions relating to meiosis that have occurred in the lineages giving rise to the angiosperms.

Key words: Meiosis, organogenesis, recombination, sex, sporangia.

Introduction

Man has selected from natural genetic variation to breed

crop species useful for agriculture. Considerable genetic

diversity remains to be fully exploited, including variationcapable of increasing yield and broadening crop range.

Sustainable increases in agricultural yield in the range of

50% will be required to feed the 9 billion people estimated to

exist by 2050 (The Royal Society, 2009). To increase the

efficiency of crop breeding, it is important to understand the

mechanism by which variation is generated and transmitted.

Variation is generated in a specialized, reductive cell di-

vision termed meiosis during which recombination betweenhomologous chromosomes, termed crossover (CO), occurs.

CO frequency varies within and between species and can be

limiting such that useful variation is not accessible for

breeding. Although the majority of crops are angiosperms

in which meiosis occurs within floral organs, non-flowering

plants have provided novel insights into the control of plant

meiosis. The nature of the meiotic cell division in plants, the

genetic mechanisms that promote variation, and the de-velopmental context in which meiosis occurs in land plants

and their closest sister groups are reviewed here.

Meiotic cell divisions

Meiosis is a mode of cell division specific to eukaryotic

organisms whereby four haploid daughter cells are produced

from a single diploid parent cell (Villeneuve and Hillers,

2001; Hamant et al., 2006; Mezard et al., 2007). This

reductive division is achieved by a single round of DNA

replication followed by two rounds of chromosome segre-

gation and cell division (meiosis-I and meiosis-II) (Fig. 1).

Meiosis appears to be an ancestral trait within eukaryotes

and is speculated to have arisen close to the group’s origin

(Villeneuve and Hillers, 2001; Cavalier-Smith, 2002). This

idea is strengthened by observations that many unicellular

eukaryotes, once presumed to be asexual, have since been

found to possess conserved meiosis-specific genes (Ramesh

et al., 2005; Malik et al., 2007). The first meiotic division

differs dramatically from mitosis as homologous chromo-

somes pair before segregation. During the second division

sister chromatids segregate to opposite poles in the same

manner as during mitosis. While homologues are paired

during meiosis-I they are tightly associated via the synapto-

nemal complex that forms along their length (Page and

Hawley, 2004). Meiosis-specific expression of an endo-

nuclease, SPO11, causes a large number of double strand

breaks (DSBs) along the paired chromosomes (Villeneuve

and Hillers, 2001). A subset of these break sites are repaired

through recombination pathways that lead to physical

exchange between the paired chromosomes (CO) (Villeneuve

and Hillers, 2001; Hamant et al., 2006; Mezard et al., 2007).

As paired chromosomes segregate during meiosis-I, CO sites

ª The Author [2010]. Published by Oxford University Press [on behalf of the Society for Experimental Biology]. All rights reserved.For Permissions, please e-mail: [email protected]

can be visualized cytologically as chiasmata (Armstrong

et al., 2009). Independent segregation of maternal and

paternal chromosome sets during meiosis in combination

with CO between chromosomes means that gametes are

likely to possess novel combinations of genetic variation

(Fig. 1).

Programmed DNA breakage and repair

An essential first step in achieving CO is the generation of

DSBs throughout the genome by SPO11. SPO11 is related

to the A subunit of archaebacterial topioisomerase IV,which acts to relieve torsion in DNA by generating

transient DSBs (Villeneuve and Hillers, 2001; Cavalier-

Smith, 2002). Hence, components of the meiotic recombina-

tion machinery may have been recruited from prokaryotic

DNA repair mechanisms. Two related orthologues SPO11-

1 and SPO11-2 are non-redundantly required for meiotic

DSBs in Arabidopsis thaliana (Grelon et al., 2001; Stacey

et al., 2006; Hartung et al., 2007). The spo11-1 and spo11-2

mutants lack meiotic DSBs and show an absence of

homologue pairing and synapsis, meaning that univalent

chromosomes segregate at meiosis-I (Grelon et al., 2001;

Stacey et al., 2006; Hartung et al., 2007). As univalent

chromosomes segregate independently from their homolo-

gous partner, spo11 mutants show a high incidence of

chromosomally unbalanced gametes (Grelon et al., 2001;

Stacey et al., 2006; Hartung et al., 2007). The PUTATIVE

RECOMBINATION INITIATION DEFECTS1 (PRD1),

PRD2 and PRD3 genes are also necessary for SPO11-

dependent DSB formation and cause univalent segregation

when mutated (De Muyt et al., 2007, 2009). PRD1 and

PRD3 share sequence similarity with known meiotic

proteins, mammalian Mei1 and rice PAIR1, respectively,

but perform unknown functions during DSB formation

(Libby et al., 2002; Nonomura et al., 2004; De Muyt et al.,2007, 2009).

A second functional class of genes, including MND1,

AHP2, RAD50, RAD51C, XRCC3, MRE11, and COM1/

SAE2, share a mutant phenotype of meiotic chromosome

fragmentation without synapsis, which is suppressed when

combined with spo11 or prd3. (Schommer et al., 2003;

Bleuyard et al., 2004; Bleuyard and White, 2004; Puizina

et al., 2004; Li et al., 2004; Kerzendorfer et al., 2006; Panoliet al., 2006; Uanschou et al., 2007; Vignard et al., 2007).

This indicates that these proteins act to process DSBs

during meiotic recombination, and unrepaired DSBs cause

chromosome fragmentation. Meiotic DSB processing pro-

ceeds via break site resection to form 3’ single-stranded

DNA, which is used to invade the intact duplex of a paired

homologous chromosome (Bhatt et al., 2001; Villeneuve

and Hillers, 2001; Hamant et al., 2006; Mezard et al., 2007).Invasion of DNA duplexes by ssDNA requires the DMC1

and RAD51 recombinases, which are related to prokaryotic

RecA DNA strand exchange proteins (Couteau et al., 1999;

Masson and West, 2001; Villeneuve and Hillers, 2001;

Cavalier-Smith, 2002; Li et al., 2004). Though RAD51

Fig. 1. Chromosome inheritance during meiosis. (A) Maternally

(white) and paternally (black) inherited copies of two chromosome

pairs are shown in a diploid parental cell. (B) During S-phase of

meiosis-I each chromosome is replicated. (C) Maternally and

paternally inherited pairs of homologous chromosomes physically

associate during prophase of meiosis-I. Coincident with pairing,

a large number of DNA double-strand breaks (shown as short

vertical lines) are induced by the SPO11 endonuclease. (D) At the

end of meiosis-I spindle microtubules attach to the centromeres of

paired bivalents and recombination events are completed.

(E) Members of homologous chromosome pairs segregate to

opposite cell poles. (F) Meiosis-I is completed with equal numbers

of replicated, recombined chromosomes in daughter cells.

(G) During meiosis-II cohesion is lost between replicated chroma-

tids, which segregate to opposite cell poles. (H) Four haploid

daughter cells are produced with novel combinations of maternally

and paternally inherited genetic information.

2864 | Harrison et al.

and DMC1 are both thought to act at the step of

ssDNA invasion, rad51 mutants show SPO11-dependent

chromosome fragmentation, whereas dmc1 mutants show

univalent segregation (Couteau et al., 1999; Li et al., 2004).

One explanation for this difference may be that DSBs are

repaired using sister chromatids in dmc1, but not in rad51

(Couteau et al., 1999; Li et al., 2004).

DSBs lead to CO via formation of a double Hollidayjunction (dHJ) between paired homologous DNA duplexes

(Villeneuve and Hillers, 2001). Subsequent to strand in-

vasion, a series of events are required for dHJ formation,

including second end capture and DNA synthesis and

ligation (Villeneuve and Hillers, 2001). These junctions are

ultimately resolved into CO events via an unknown dHJ

resolvase in plants, although the structure-specific endonu-

cleases GEN1/Yen1 and SLX4 serve this function in animals(Svendsen and Harper, 2010). An excess of DSBs are

generated by SPO11 relative to the number of CO events

ultimately observed. The remaining DSBs are repaired via

a gene conversion (also known as a non-crossover, NCO)

pathway, which does not involve the exchange of flanking

genetic markers (Villeneuve and Hillers, 2001). The synthe-

sis-dependent strand annealing pathway appears to be

a major mechanism for NCO formation in Saccharomyces

cerevisiae, which acts independently of dHJ formation

(McMahill et al., 2007). In S.cerevisiae the decision to

repair DSB sites as CO or NCO events is made early in

prophase-I, prior to dHJ formation, so it is unlikely that the

CO/NCO choice represents alternative processing of dHJs

(Allers and Lichten, 2001; Borner et al., 2004).

Control of meiotic crossover frequency

The position of COs can be influenced by other COs on the

same chromosome via a phenomenon known as interfer-

ence. In interference one event inhibits the formation of

adjacent events in a distance-dependent manner meaningthat they are more widely distributed than expected at

random (Sturtevant, 1915; Muller, 1916; Copenhaver et al.,

2002). However, non-interfering COs that have a random

distribution also occur (Copenhaver et al., 2002). In A.

thaliana class I interfering COs are the majority (;75–85%)

and class II non-interfering COs the minority (;15–25%)

(Copenhaver et al., 2002; Higgins et al., 2004; Mercier et al.,

2005; Berchowitz et al., 2007; Higgins et al., 2008a). Class ICOs in A. thaliana require a set of genes including MSH4,

MSH5, MLH3, MER3/ROCK-N-ROLLERS, PARTING

DANCERS, ZIP4/SPO22, RPA1a and SHOC1 (Higgins

et al., 2004, 2008b; Chen et al., 2005; Mercier et al., 2005;

Jackson et al., 2006; Wijeratne et al., 2006; Chelysheva

et al., 2007; Macaisne et al., 2008; Osman et al., 2009).

Knockout of these genes results in a dramatic reduction in

CO frequency and the events that remain are randomlydistributed. The ancestry of meiotic proteins in prokaryotic

DNA repair is again evident as MSH4 and MSH5 proteins

are related to bacterial MutS mismatch repair proteins

(Villeneuve and Hillers, 2001; Cavalier-Smith, 2002). Class

II non-interfering COs require the MUS81 gene, which

encodes a protein similar to structure-specific endonucleases

(Berchowitz et al., 2007; Higgins et al., 2008a).

Paired chromosomes generally show at least one CO

event, the obligate CO, that is required for proper patterns

of chromosome segregation in A. thaliana (Grelon et al.,

2001). Each A. thaliana chromosome shows approximately

1.8 CO per meiosis and very few chromosomes show no CO

(Copenhaver et al., 1998, 2002; Higgins et al., 2004;Drouaud et al., 2006, 2007). CO position is highly variable

and hot- and cold-spots of CO frequency exist along the

chromosomes (Copenhaver et al., 1998, 1999; Drouaud

et al., 2006, 2007). Pronounced increases in CO frequency

and gene density are observed towards the telomeres of

wheat and maize chromosomes (Liu et al., 2009; Saintenac

et al., 2009; Schnable et al., 2009). A recombination hotspot

at the maize Bronze (Bz) locus lies in a gene-rich regionclose to the telomere. This region shows 40–80-fold higher

CO rates than the genome average and is flanked by large

stretches of nested retrotransposon insertions that infre-

quently crossover (Dooner and Martinez-Ferez, 1997; Fu

et al., 2001, 2002). Repetitive regions flanking centromeres

are also suppressed for CO, and centromere-proximal

events associate with chromosome mis-segregation (Koehler

et al., 1996; Lamb et al., 1997; Deng and Lin, 2002;Rockmill et al., 2006).

Differences in crossover frequency may be accounted for

by epigenetic information. For example in mouse and S.

cerevisiae H3 K4 trimethylation marks DSB hotspots, and

disruption of this modification reduces DSBs (Borde et al.,

2009; Buard et al., 2009). Conversely, in A. thaliana the CO-

cold repetitive sequences flanking the centromere are

transcriptionally silenced using epigenetic information in-cluding DNA cytosine methylation and targeted DNA

methylation in Ascobolous immersus is sufficient to repress

CO frequency several hundred fold (Maloisel and Rossi-

gnol, 1998; Zhang et al., 2006; Zilberman et al., 2007).

Repetitive insertions and inversions can also locally sup-

press CO, and play an important functional role in genome

organization (Dooner, 1986; Nacry et al., 1998). This is

illustrated by CO suppression at sex chromosomes, mating-type loci and self-incompatibility loci, where it is important

to maintain linkage between genes required for opposite

mating/incompatibility types (Ferris and Goodenough,

1994; Casselman et al., 2000; Ming and Moore, 2007;

Bergero and Charlesworth, 2009). Hence, CO frequency is

likely to be determined by a combination of local DNA

sequence, trans-factors, and epigenetic information.

Control of meiotic cell cycle progression

Meiosis requires the modification of mitotic cell cycle

control, such that a single S-phase is followed by twosequential rounds of chromosome segregation. Progression

through the cell cycle is controlled by cyclins that interact

with and activate cyclin-dependent kinases (CDKs) which

mediate stepwise transitions through the cycle via phos-

phorylation (Huntley and Murray, 1999). Several genes

Flowering and meiosis | 2865

implicated in the regulation of meiotic progression have

been identified in A. thaliana. A novel, cyclin-like gene

SOLO DANCERS (SDS) is specifically expressed during

meiosis and sds mutants show defects in chromosome

pairing, segregation, and CO (Azumi et al., 2002). Recently,

sds mutants have been observed to form DSBs but repair

them efficiently, most likely via RAD51-mediated inter-

sister repair (De Muyt et al., 2009). The novel geneOMISSION OF SECOND DIVISION1 (OSD1) is required

for meiosis-II and osd1 produces diploid dyad products of

meiosis instead of haploid tetrads (d’Erfurth et al., 2009). A

temperature-sensitive substitution allele of CYCLINA1;2,

termed tardy asynchronous meiosis1 (tam1) causes a delay in

meiotic progression, which also leads to dyad formation

(Magnard et al., 2001; Wang et al., 2004). Regulation of

protein stability is critical for cell cycle control and SKP1-like F-box proteins act to promote ubiquitination and

destruction of target proteins, including cyclins (Bai et al.,

1996). Consistently, mutations in the SKP1-related gene

ASK1 show defects in meiotic chromosome segregation

(Yang et al., 1999a; Zhao et al., 2006). In animals, cell cycle

checkpoints cause later events to depend upon the comple-

tion of earlier events (Murakami and Nurse, 1999). Meiotic

checkpoint mechanisms have not been genetically defined inA. thaliana, although msh4 and asy1 show a significant

delay in meiosis, suggesting feedback on the regulation of

meiotic progression (Higgins et al., 2004; Sanchez-Moran

et al., 2007). Together these results demonstrate that defects

in cell cycle progression can disrupt meiosis.

Key early events in meiosis are the identification of

homologous chromosome partners, pairing, and formation

of the synaptonemal complex (SC) (Page and Hawley,2004). Homologous partner identification occurs by un-

known mechanisms during prophase-I. In polyploid species

pairing is complex as homologues must also avoid pairing

with related homeologues (Martinez-Perez et al., 1999). For

instance, in hexaploid wheat Ph1 affects the stringency of

homologue/homeologue discrimination by influencing chro-

matin remodelling associated with pairing and localization

of the SC component ASY1 (Martinez-Perez et al., 1999;Prieto et al., 2004, 2005; Boden et al., 2009). Ph1 maps to

a repetitive locus containing cell cycle-dependent kinase

genes, which causes down-regulation of unlinked CDK

genes (Griffiths et al., 2006; Al-Kaff et al., 2008). As related

CDK genes in mammals influence meiotic progression,

trans-silencing of CDKs by Ph1 could lead to changes in

homologue pairing (Ortega et al., 2003). Pairing may

depend on DSB formation as in A. thaliana or may occurvia achiasmate mechanisms as in female Drosophila mela-

nogaster (Grelon et al., 2001; Page and Hawley, 2004).

SWITCH1/DYAD encodes a novel protein expressed during

prophase-I, necessary for chromosome pairing, synapsis, and

recombination and in swi1/dyad mutants univalents segregate

at meiosis-I. Studies of a maize homologue, AMEIOTIC1

indicate that these functions are conserved within angio-

sperms (Golubovskaya et al., 1993; Mercier et al., 2001, 2003;Agashe et al., 2002; Ravi et al., 2008; Pawlowski et al., 2009).

POOR HOMOLOGOUS SYNAPSIS1 encodes a second

novel protein required for chromosome pairing and synapsis,

which causes high levels of non-homologous pairing when

mutated (Pawlowski et al., 2004; Ronceret et al., 2009).

Co-incident with pairing, the SC forms between homolo-

gous chromosomes (Page and Hawley, 2004) and SC

components identified in A. thaliana include ASYNAPTIC1

(ASY1), ZYP1A, ZYP1B, SCC3, and REC8/DIF1/SYN1

(Bai et al., 1999; Bhatt et al., 1999; Caryl et al., 2000;Armstrong et al., 2002; Cai et al., 2003; Chelysheva et al.,

2005; Higgins et al., 2005). ASY1 and ZYP1 show distant

identity with the animal HOP1 and ZIP1 SC proteins,

respectively (Caryl et al., 2000; Armstrong et al., 2002;

Higgins et al., 2005). Loss of SC proteins causes failures in

synapsis and CO formation, and can lead to univalent

segregation and chromosome fragmentation (Bai et al.,

1999; Bhatt et al., 1999; Caryl et al., 2000; Armstronget al., 2002; Cai et al., 2003; Chelysheva et al., 2005; Higgins

et al., 2005). Interestingly, increases in CO frequency are

associated with increases in total SC length via an unknown

mechanism (Lynn et al., 2002; Drouaud et al., 2007). These

genetic mechanisms provide novel insights into the in-

terrelated processes of homologue recognition, pairing, and

synapsis in plants.

Correct patterns of chromosome segregation are requiredto generate balanced gametes and depend on regulation of

the SC and chromosome cohesion. During mitosis the

cohesin complex holds sister chromatids together until the

SCC1 subunit is cleaved by SEPERASE1 at anaphase,

allowing chromosome segregation (Uhlmann et al., 1999).

REC8/DIF1/SYN1 is a meiosis-specific orthologue of SCC1

and the rec8/dif1/syn1 mutant disrupts normal SC localiza-

tion of SCC3 (a cohesin subunit shared with mitosis) (Bhattet al., 1999; Cai et al., 2003; Chelysheva et al., 2005). As in

animal systems, A. thaliana REC8 is cleaved by the cysteine

protease SEPERASE1 (ESP1) (Liu and Makaroff, 2006).

Cohesion in the chromosome arms is released by ESP1 at

anaphase-I, but maintained at the centromeres until

anaphase-II (Liu and Makaroff, 2006). In maize, centro-

meric REC8/AFD1 is protected from ESP1 destruction by

the conserved Shugoshin protein (SGO1) during anaphase-I(Hamant et al., 2005; Watanabe, 2005). SGO1 is then

removed and REC8 is destroyed at the centromere during

anaphase-II to allow chromatid segregation. Step-wise

formation and removal of connections between homologues

is thus required for correct patterns of meiotic chromosome

segregation and recombination.

The developmental context for meiosis

Successful completion of meiosis and sexual life cycles

depends on activation of the meiotic cell cycle at the correct

developmental time and place. In many animals separationof a dedicated germ cell lineage from somatic cell types

occurs during embryogenesis, a distinction not observed in

some early diverging animal lineages and plants (Gilbert,

1994; Dickinson and Grant-Downton, 2009). In the algal

sister groups to land plants single-celled zygotes undergo


meiosis immediately following fertilization (Fig. 2A). By

contrast, in all land plants mitotic divisions intercede

fertilization and meiosis, and meiosis occurs after a period

of diploid development in specialized structures termed

sporangia that produce numerous spores (Bower, 1935;

Becker and Marin, 2009). The initiation of sporangium

development pathways often follows a switch in meristem

identity from a vegetative to a reproductive fate (Steeves

and Sussex, 1989). The location and structure of sporangia

vary by plant group and are associated with their secondary

functions, which are spore dispersal and nutrition. Hetero-

morphic sporangia and spores have evolved convergently in

vascular plants and associate with specialized functions

(Fig. 2). Female megasporangia have fewer, larger spores

(megaspores) that may be retained within the parent plant

after fertilization, whereas male microsporangia develop

numerous small spores (microspores) that have a dispersal

function (Bower, 1935). Gender can also influence patterns

of CO frequency (Drouaud et al., 2007). Thus the initiation

and progression of meiosis depend on the developmental

identity of the tissue in which it is activated, discussed by

plant group below.

Meiosis in chlorophyte and charophyte algae

In both chlorophyte and charophyte algal sister groups to

the land plants meiosis occurs immediately following

fertilization (Becker and Marin, 2009). In the single-celled

chlorophyte alga, Chlamydomonas reinhardtii two haploid

mating types, plus and minus, differentiate into gametes

which fuse during fertilization to form a single-celled zygote

Fig. 2. Phylogenetic distribution of characters associated with meiosis in plants. (A) Phylogenetic relationships between major plant groups

showing synapomorphies associated with meiosis. Heterospory has a polyphyletic origin in lycophytes, monilophytes, and spermatophytes,

indicated by asterisks. (B) Reproductive structures of representatives of clades illustrated in (A), and extinct protracheophyte fossil forms.

1. Electron micrograph of Chlamydomonas monoica zygospores (photograph courtesy of Professor Karen P Van Winkle-Swift and by kind

permission of John Wiley and Sons: see VanWinkle-Swift KP, Rickoll WL. 1997. The zygospore wall of Chlamydomonas monoica

(Chlorophyceae): Morphogenesis and evidence for the presence of sporopollenin1. Journal of Phycology 33, 655–665.) 2. Light micrograph

of the haploid plant and oogonium (inset) in the charophyte alga Chara sp. 3. The creeping haploid thallus and a diploid erect and

determinate sporophyte of Pellia epiphylla (photographs of bryophytes courtesy of Li Zhang). 4. The haploid leafy gametophyte of Atrichum

angustatum bearing diploid unbranched determinate sporophytes. 5. The haploid creeping thallus of Folioceros sp. with upright

indeterminate sporophytes. 6. A fossil sporophtye of Cooksonia sp. showing branching with terminal sporangia (photo courtesy of Jenny

Morris and Dianne Edwards). 7. A fossil sporophyte of Zosterophyllum showing lateral sporangia (photo courtesy of Jenny Morris and

Dianne Edwards). 8. A Selaginella kraussiana sporophyte showing vegetative branching habit, an unbranched reproductive strobilus, and

a mega- and microsporangium (inset). 9. Sporangia formed on the stem of Psilotum nudum. 10. A frond of Adiantum mairisi showing

marginal sori that contain sporangia. 11. The terminal cones of Cupressus sp. that contain the mega- and microsporangia. 12. The

stamens and carpels of a Magnolia sp. flower that contain the micro- and megasporangia, respectively.


(Fig. 2B) (Lee et al., 2008). Plus and minus gamete identity

is specified from the MATING-TYPE locus and requires

cytoplasmic accumulation of BEL [GAMETE-SPECIFIC

PLUS1 (GSP1)] and KNOX [GAMETE-SPECIFIC MI-

NUS1 (GSM1)] class homeodomain proteins, respectively

(Ferris and Goodenough, 1994; Lee et al., 2008). Following

fertilization GSP1 and GSM1 heterodimerize, translocate to

the nucleus, and initiate zygotic gene expression patterns(Lee et al., 2008). Constitutive expression of either GSP1 or

GSM1 in the opposite gamete type is sufficient to trigger

zygote development in the absence of fertilization (Zhao

et al., 2001; Lee et al., 2008). Stable C. reinhardtii diploids

that do not initiate meiosis can also be generated, and

constitutive expression of GSP1/GSM1 together in these

cells is sufficient to induce meiosis with normal patterns of

recombination (Lee et al., 2008). This indicates that GSP1/GSM1 homeodomain proteins are potential triggers of

meiosis in a chlorophyte alga. In contrast to chlorophytes,

charophytes have a multicellular haploid body that gener-

ates free-swimming sperm in antheridia and egg cells that

are retained within an oogonium on the parent plant

(Fig. 2B). Egg retention (oogamy) is an innovation shared

with land plants thought to have been a key adaptation in

their evolution (McCourt et al., 2004). As there arecurrently no charophyte genetic models, potential roles for

KNOX/BEL genes are unexplored, and the initiation of

meiosis in charophytes is via an unknown mechanism.

Sporophyte development and meiosis inbryophytes

In contrast to their algal sisters, all land plants have a period

of multicellular diploid growth, the extent of which varies

by plant group (Lewis and McCourt, 2004; McCourt et al.,

2004; Becker and Marin, 2009). The bryophyte sister groups

to the vascular plants exhibit limited post-embryonic de-velopment with no indeterminate apical growth (Mishler

and Churchill, 1985; Shaw and Renzaglia, 2004; Donoghue,

2005). Sporophytes comprise a small single stem with

a terminal sporangium that represents the simplest basal

land plant body plan (Kenrick, 2002; Donoghue, 2005; Qiu

et al., 2006; Boyce, 2008) (Fig. 2). In liverworts and mosses

sporangium development arrests diploid growth, whereas

hornwort sporophytes contribute to their own nutrition andhave sporangia that grow indeterminately from a basal

meristem (Boyce, 2008; Kato and Akiyama, 2005). A sub-

epidermal archesporial cell layer is specified during sporan-

gium development and divides either by meiosis to generate

spores (mosses) or spore mother cells and interspersed elater

cells that perform nutritive or dispersal functions (liverworts

and hornworts). The tissues surrounding the archesporial

cell layer perform dispersal functions specific to eachbryophyte group (Bower, 1935). The genetic and develop-

mental mechanisms that regulate bryophyte sporophyte

development are currently poorly understood, but interest

has recently accelerated due to the establishment of moss

(Physcomitrella patens) and liverwort (Marchantia polymor-

pha) models (Ishizaki et al., 2008; Rensing, 2008). Two gene

classes that affect sporangium development in P. patens are

homologues of A. thaliana LEAFY (LFY) and KNOX genes

(Champagne and Ashton, 2001; Tanahashi et al., 2005;

Sakakibara et al., 2008). A pair of LFY homologues

redundantly control the first zygotic division in P. patens

and double mutants exhibit arrested zygotic development

(Tanahashi et al., 2005). In mutants that do not arrest,sporangium number, initiation, and development are per-

turbed. These defects may arise as a consequence of

abnormal sporophytic development, although spore number

and germination are also highly variable in the mutants,

suggesting meiotic defects (Tanahashi et al., 2005). Similarly

Class I KNOX mutants in P. patens have abnormal

sporangia and reduced spore numbers (Sano et al., 2005;

Sakakibara et al., 2008). Interestingly KNOX expression issporophyte specific in P. patens (Champagne and Ashton,

2001; Sakakibara et al., 2008), and a potential role of

KNOX and BEL genes in diploid development conserved

between algae and bryophytes remains to be explored.

Protracheophytes and seed plant sistergroups

Key features that distinguish vascular plants from bryo-

phytes are the elaboration of an indeterminately growing

and branching diploid body (Mishler and Churchill, 1985;

Donoghue, 2005; Langdale and Harrison, 2008). Fossilplants whose form is not represented in living plants, such

as Cooksonia, have low orders of branching and may have

amplified spore numbers by increasing numbers of terminal

sporangia (Fig. 2B) (Edwards and Feehan, 1980; Graham

et al., 2000; Donoghue, 2005; Gerrienne et al., 2006). These

fossils raise interesting questions about the developmental

nature of the association between axis development, spo-

rangium development, and branching and, intriguingly, rarebryophyte branching mutants strikingly resemble Cooksonia

sporophytes (Fig. 2B). Alternative lateral sporangial place-

ments appear independent of branching and may have

served a similar purpose in other fossil groups (Fig. 2B).

This arrangement is exhibited in modern lycophytes, and

sporangia arise either at the base of leaves or from the stem

via one or two sub-epidermal archesporial cell layers. These

archesporial cells give rise to sporogenous tissue (Lycopo-dium) or sporogenous and tapetal tissues (Selaginella,

Isoetes) (Bower, 1935). Monilophyte sporangia are diverse

in terms of their size, the number of spores produced per

sporangium, and their number and position on the plant

(Fig. 2B). The eusporangiate basal monilophyte grade

comprising marattioid ferns, horsetails, ophioglossoid ferns,

and whisk ferns possess sporangia that develop from several

cells and produce thousands of spores (Bower, 1935;Wagner, 1977; Parkinson, 1987; Pryer et al., 2004). By

contrast, the leptosporangiate ferns develop numerous,

small sporangia from single cells, which typically contain

tens of spores (Bower, 1935; Pryer et al., 2004). Sporangia

may show terminal, adaxial, abaxial, or marginal locations


on leaves (Fig. 2B). With the exception of the leptosporan-

giate and whisk ferns, nutritive tapetal tissues arise from

non-sporogenous tissue (Bower, 1935; Parkinson, 1987). As

in bryophytes, the genetic basis of diploid development is

poorly characterized in lycophytes and monilophytes.

Notably sporophytic KNOX expression is conserved, and

meristematic expression domains suggest likely roles in

indeterminate growth (Bharathan et al., 2002; Harrisonet al., 2005; Sano et al., 2005), although reproductive roles

have not yet been explored. Thus the structure and dispersal

functions of sporangia vary broadly across the land plants

and the developmental context for the initiation of meiosis

is lineage specific. An evolutionary trend towards the

amplification of spore numbers by alterations in body plan,

sporangium size, and the number of sporangia is apparent

(Bower, 1935).

Sporophyte development in seed plants

In seed plants (gymnosperms and angiosperms) a prolongedperiod of vegetative growth is followed by the reproductive

transition. This transition involves a change in meristem

identity and leads to the development of cones or flowers

(Steeves and Sussex, 1989). Seeds develop in the context of

the ovule following fertilization of the female egg cell by

a male sperm cell transferred in pollen, thus dispersal

functions are provided both by haploid pollen and diploid

seed. Ovules are the site of megasporangium (nucellus)development, which precedes meiosis. Whilst in gymno-

sperms one to several nucellar cells enter meiosis, in

angiosperms a single megaspore mother cell undergoes

meiosis to form a tetrad, three members of which de-

generate to form a single functional megaspore, which

divides mitotically to form the embryo sac (Campbell,

1940; Colombo et al., 2008). Pollen sac (microsporangium)

development occurs from a microsporophyll or in theanther in gymnosperms and angiosperms respectively. In

both, sub-epidermal cells are specified as archesporial cells

that divide periclinally to form a layer of parietal cells

surrounding the sporogenous cells (Campbell, 1940; Feng

and Dickinson, 2007). Sporogenous cells may then either

directly enter meiosis or continue to proliferate. Parietal

cells divide further to form a variable number of concentri-

cally arranged cell layers, the innermost of which differ-entiates into the nutritive tapetum (Campbell, 1940;

Feng and Dickinson, 2007). During meiosis sporogenous

cells become encased in an impermeable callose ((1-3)-

b-D-glucan) wall, which later breaks down when members

of the microspore tetrads are released (Gifford and Foster,

1988). Callose appears to play an important role in

sporogenesis, as tapetal expression of callase causes male

sterility in tobacco (Worrall et al., 1992). Following meiosis,the resulting haploid microspores undergo mitosis and

differentiate into pollen grains.

The genetic control of vegetative development, the re-

productive transition, and sporangium formation are well

studied in the angiosperm A. thaliana. Activity of the class I

KNOX gene SHOOTMERISTEMLESS (STM) is neces-

sary for the establishment of an indeterminate meristem

(Long et al., 1996), and STM and BREVIPEDICELLUS

(BP) act redundantly to maintain indeterminacy and repress

determinate leaf development (Byrne et al., 2002). Class I

KNOX proteins promote indeterminacy by dimerizing with

BEL transcription factors and triple bellringer, poundfoolish,

Arabidopsis thaliana homeobox1 BEL mutants phenocopystm mutants (Rutjens et al., 2009). Thus, in A. thaliana

KNOX and BEL genes play key roles in elaboration of the

diploid body, providing the context for later reproductive

development and meiosis. Flower development follows

conversion of indeterminate, vegetative shoot meristems to

reproductive fates. This switch is controlled by a large

network of genes that ensure reproduction is co-ordinated

with environmental and developmental conditions (Baurleand Dean, 2006). These signalling pathways converge on

a key set of transcription factors required for floral

meristem identity, including LEAFY and APETALA1

(Baurle and Dean, 2006). Floral organ identity genes

encode three functional classes of MADS-box transcription

factors (A, B, and C) that are expressed in overlapping

domains to specify the four floral organ types (Coen and

Meyerowitz, 1991). Stamen identity requires overlappingexpression of the B and C class MADS genes PISTILLATA

and APETALA3, and AGAMOUS (AG) (Yanofsky et al.,

1990; Jack et al., 1992; Goto and Meyerowitz, 1994). Carpel

identity requires activity of the C class gene AG, which acts

in conjunction with three additional MADS proteins,

SEEDSTICK, SHATTERPROOF1, and SHATTER-

PROOF2 to specify ovule identity (Yanofsky et al., 1990;

Pinyopich et al., 2003). The KNOX genes STM andKNAT2, and BEL1 genes also play roles in the specification

of carpel and ovule identity (Modrusan et al., 1994; Pautot

et al., 2001; Scofield et al., 2007). Thus genes involved in

meristem identity also play roles in the specification of

reproductive fate.

In both micro- and megasporangia archesporial cell

specification precedes sporangium formation (Gifford and

Foster, 1988). The mechanisms of archesporial cell spec-ification are poorly understood, but one gene, SPORO-

CYTELESS/NOZZLE (SPL/NZZ), functions directly

downstream of the C class organ identity gene AG to

promote sporogenesis (Schiefthaler et al., 1999; Yang et al.,

1999b; Ito et al., 2004). SPL is a nuclear protein with distant

homology to MADS box transcription factors that per-

forms an unknown function (Schiefthaler et al., 1999; Yang

et al., 1999b). spl mutants differentiate microsporangialarchesporial cells that divide once, but fail to form micro-

sporocytes or tapetal cells, causing sterility (Schiefthaler

et al., 1999; Yang et al., 1999b). The spl/nzz mutants are

also female sterile due to nucellar defects, meaning that the

archesporial cells do not differentiate and meiosis fails to

initiate (Schiefthaler et al., 1999; Yang et al., 1999b).

Ectopic activation of SPL in agamous mutants is sufficient

to induce staminoid development and pollen formation (Itoet al., 2004). Together this indicates that SPL/NZZ

performs an upstream meiotic function in both male and


female development. Male sporocyte identity in A. thaliana

is also regulated by the leucine-rich repeat (LRR) receptor

kinase EXTRA SPOROGENOUS CELLS/EXCESS

MICROSPOROCYTES1 (EXS/EMS1) in conjunction with

its small protein ligand TAPETUM DETERMINANT1

(TPD1) (Yang et al., 1999b, 2003; Canales et al., 2002; Zhao

et al., 2002). The first archesporial cell division normally

separates reproductive sporocyte fate from non-reproduc-tive wall and tapetal fates. Microsporangial development is

altered in exs/ems1 and tpd1 mutants such that sporogenous

cells develop at the expense of tapetal cells (Yang et al.,

1999b, 2003; Canales et al., 2002; Zhao et al., 2002). This

implies that EXS/EMS1 kinase signalling is important either

to promote tapetal or to repress sporogenous cell identity.

Although exs/ems1 mutations in A. thaliana show normal

megasporangial development, mutations in the rice homo-logue MULTIPLE SPOROCYTES1 (MSP1) show super-

numerary sporocytes in both the anther and ovule, as does

the multiple archesporial cells1 (mac1) mutant in maize

(Sheridan et al., 1996; Nonomura et al., 2003). Interestingly,

additional LRR receptor kinases have also been implicated

in proper differentiation of the anther cell layers (Albrecht

et al., 2005; Colcombet et al., 2005; Mizuno et al., 2007;

Hord et al., 2008). How these signalling processes areorganized between the cell types within the developing

anther is not yet clear.

Future perspectives

Developmental genetic studies in A. thaliana have signifi-

cantly advanced our understanding of the context in which

meoisis arises in plants. Future goals will be to tie together

our understanding of the context, initiation, and progress of

meiosis in diverse plant groups so that potential variation

can be released to breeding. During plant diversification,

genes that may have originally been involved in reproduc-tive development have been co-opted to vegetative de-

velopment pathways. Whilst KNOX/BEL proteins may

trigger meiosis in a chlorophyte alga, their role is unknown

in charophytes and most bryophytes and thus the point of

functional diversification remains to be identified. The

mechanisms for archesporial cell development are not yet

fully characterized in flowering plants and are unknown in

non-flowering plants. The derivation of nutritive tapetaltissues in different plant groups may or may not be

independent of archesporial lineages. It will be interesting

to test homology between archesporial and tapetal cell types

by testing the function of SPL, EXS, and TPD1 homo-

logues in different lineages. A detailed picture is emerging of

the mechanisms that control plant meiotic chromosome

pairing, synapsis, recombination, and segregation in A.

thaliana. Understanding how these mechanisms integrateduring progression of the meiotic cell cycle will be a major

challenge. Equally, the pattern of CO hot- and cold-spots is

complex and the mechanisms that determine plant CO

frequency remain to be determined. Knowledge of these

mechanisms may allow CO to be targeted during crop

breeding and facilitate the generation of novel high-yielding

agricultural strains.

Acknowledgements

We thank the Royal Society and the Gatsby Charitable

Foundation for funding, Karen Van Winkle-Swift, Vasily

Kantsler, Li Zhang, Jenny Morris, and Dianne Edwards for

photographs, and two anonymous reviewers for helpful

comments on the manuscript.

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