Analyse fonctionnelle du rôle de la chromatine dans le contrôle de l ...

UNIVERSITÉ DE PERPIGNAN VIA DOMITIA
ECOLE DOCTORALE SIBAGHE
Laboratoire Génome et Développement des Plantes CNRS/IRD/UP 5096
THÈSE
Pour obtenir le grade de
DOCTEUR DE LʼUNIVERSITÉ DE PERPIGNAN VIA DOMITIA
Discipline Biologie Intégrative des Plantes
présentée et soutenue publiquement par
Marcelina GARCIA-AGUILAR
le 28 Janvier 2010
ANALYSE FONCTIONNELLE DU RÔLE DE LA CHROMATINE
DANS LE CONTRÔLE DE LʼAPOMIXIE ET DE LA SEXUALITÉ
Thèse dirigée par Daniel GRIMANELLI & Alain GHESQUIERE
Jury:
Thomas DRESSELHAUS!! ! ! ! ! ! Rapporteur
Raphäel MERCIER! ! ! ! ! ! ! Rapporteur
Jean-Marc DERAGON! ! ! ! ! ! ! Examinateur
Célia BAROUX! ! ! ! ! ! ! ! Examinateur
Alain GHESQUIERE! ! ! ! ! ! ! Directeur de Thèse
Daniel GRIMANELLI! ! ! ! ! ! ! Directeur de Thèse

“Todas las personas al comienzo de su juventud, saben cual es su
leyenda personal. En ese momento de la vida todo es claro, todo es
posible, y ellas no tienen miedo de soñar y desear todo aquello que
les gustaría hacer en sus vidas. El alma del mundo es alimentada por
la felicidad de las personas. O por la infelicidad, la envidia, los celos.
Cumplir su leyenda personal es la única obligación de los hombres.
Y cuando una persona desea realmente algo, el universo entero
conspira para que pueda realizar su sueño”
Paolo Coelho
2

A la memoria de mi Padre,
Por enseñarme que la motivación es el mejor
camino para lograr nuestros sueños, porque
siempre viviste compartiendo tu alegría, por
heredarme tu amor al maíz.
Donde estés, Te amo Papá.
3

Résumé
De nombreuses données expérimentales suggèrent que les phases de transition au cours de
la reproduction sexuée sont régulées épigénétiquement par des modifications de la structure
de la chromatine. Chez les Angiospermes, l’apomixie est un mode de reproduction asexuée
par graines conduisant à la production de descendances génétiquement identiques à la plante
mère. Contrairement à ceux obtenus par reproduction sexuée, qui résultent de l’union de deux
produits de la méiose, les embryons apomictiques sont produits sans méiose ni fécondation.
L’objectif de cette étude était de déterminer le rôle de la structure de la chromatine au cours
de la reproduction des plantes et de tester l’hypothèse d’un lien fonctionnel entre
l’établissement de ces structures chromatidiennes et la différentiation entre sexualité chez le
maïs et apomixie chez son apparenté sauvage, Tripsacum. Dans un crible portant sur 319
enzymes modificatrices la chromatine (CME), nous avons pu identifier six loci dont
l’expression dans les ovules diffère entre plantes sexuées et plantes apomictiques. Quatre
d'entre eux codent pour des protéines homologues de composants de la voie de méthylation de
l'ADN dépendante de l'ARN (RdDM) chez Arabidopsis thaliana. L’analyse fonctionnelle de
DMT102, l’homologue de CHROMOMETHYLASE 3, et de DMT103, l’homologue de
DOMAIN REARRANGED METHYLTRANSFERASE 2, montre qu’une perte de fonction
induit la production de gamètes non-réduits et la formation de sacs embryonnaires multiples
dans l'ovule, deux aspects clés du développement apomictique. Des expériences
d'immunolocalisation dans des ovules de maïs indiquent que DMT102 est impliquée dans la
définition d’un état de quiescence transcriptionelle affectant un nombre limité de cellules, y
compris le gamète femelle, et que la perte de méthylation de l'ADN observée dans le mutant
DMT102 conduit à une altération de cet état répressif. Finalement, la formation au sein de
l’ovule d’un domaine cellulaire restreint, renfermant les cellules reproductrices et
transcriptionellement quiescent, est abolie au cours du développement apomictique chez
Tripsacum. Ces observations suggèrent que le mutant dmt102 reproduit l’état chromatidien
prévalent dans les ovules apomictiques. Nos résultats montrent qu’une voie de type RdDM
spécifique des gamétophytes est impliquée chez le maïs dans la définition des cellules
reproductrices et que son altération participe à la différenciation entre reproduction sexuée et
reproduction apomictique.
Mots clefs: reproduction sexuée, apomixis, structure de la chromatine, regulation
epigénétique, maïs, Tripsacum.
4

FUNCTIONAL AND COMPARATIVE ANALYSIS OF CHROMATIN CHANGES
ASSOCIATED WITH THE CONTROL OF SEXUAL AND APOMICTIC REPRODUCTION
5

Summary
Several lines of evidences suggest that transitions during sexual plant reproduction are
epigenetically controlled through dynamic changes in chromatin structure. In asexual
reproduction by seed, or apomixis, female gametes avoid meiosis and double fertilization,
developing by parthenogenesis offspring that are genetically identical to the mother plant. It
has been hypothesized that apomixis could be a temporal or spatial deregulation of the sexual
reproductive pathway. Our objective was to determine the role of chromatin dynamics during
plant reproduction, and in the molecular switch between sexual reproduction in maize, and
apomictic development in maize-Tripsacum hybrids. In a transcriptional screening of 319
chromatin-modifying enzymes (CME), we identified six loci that are specifically downregulated
in ovules of apomictic plants. Four of them share strong homology with members of
the RNA directed DNA methylation (RdDM) pathway, which in Arabidopsis is involved in
genes silencing via DNA methylation. The functional analysis of plants defective for
DMT102 (homologous to Arabidopsis CHROMOMETHYLASE 3) and DMT103 (homologous
to DOMAIN REARRANGED METHYLTRANSFERASE 2) reveals production of unreduced
gametes, and formation of multiple embryo sacs in the mutant ovules, both key features of
apomictic development. Immunolocalization experiments indicate that DMT102 is involved
in regulating chromatin state and transcription in a few cells in the maize ovule, including the
female gametes. Loss of DNA methylation in a DMT102 mutant line results in the localized
release of a repressive chromatin state, which also mimics features of apomictic ovules. Our
results suggest that reproductive development in maize requires an gametophyte-specific
RdDM-like pathway, whose regulatory role is critical to the differentiation between apomictic
and sexual reproduction.
Key words: sexual reproduction, apomixis, chromatin structure, epigenetic regulation,
maize, Tripsacum.
6

TABLE OF CONTENTS
ABBREVIATIONS 9
CHAPTER 1. General Introduction 11
1.1. Background information 12
1.1.1. Developmental Events During Sexual Plant Reproduction 12
1.1.2. Acquisition of Cell Fate in the Ovule and the Female Gametophyte 14
1.1.3. Apomictic Developments 16
1.1.4. The Genetic Control of Apomixis 19
1.1.5. Role of Hybridization and Polyploidy in Apomicts 21
1.1.6. Epigenetic, Cell Fate and Apomixis 21
1.1.7. Mechanisms of Chromatin Dynamics 23
1.1.8. Modifying Enzymes and Chromatin Dynamics 25
1.1.8.1. DNA methylation 25
1.1.8.2. Histone modifications 27
1.1.8.2.1. Histone acetylation 28
1.1.8.2.2. Histone methylation 29
1.1.8.2.3. Phosphorylation 29
1.1.8.2.4. Ubiquitylation and SUMOylation 30
1.1.8.3. ATP-dependent chromatin remodeling 30
1.2. Thesis Proposal: Chromatin dynamics, an unexplored frontier toward
understanding asexuality in plants 31
CHAPTER 2. Expression of Chromatin Modifying Enzymes during Apomictic
and sexual Development 33
2.1. Introduction 34
2.2. Results 34
2. 2.1. Selection of Chromatin Modifying Enzymes (CME) from ChromDB 34
2.2.2. Microarray analysis of Chromatin Modifying Enzymes (CME) 38
2.2.3. Preliminary analysis of selected genes 40
2.3. Discussion 42
CHAPTER 3. Inactivation of a Reproductive DNA Methylation Pathway in
Maize Results in Apomixis-Like Phenotypes 44
3.1. Introduction 47
3.2 Results 49
3.2.1. Identification of CMEs differentially expressed in sexual and apomictic plants 49
7

3.2.2. Apomixis correlates with the downregulation of CMT3 and DRM1/2 homologues in
both Tripsacum and Boechera 53
3.2.3. dmt102 and dmt103 are expressed in the reproductive cells in the maize ovule 53
3.2.4. Downregulation of dmt102 and dmt103 in sexual maize promotes the formation of
unreduced gametes 54
3.2.5. Downregulation of dmt102 and dmt103 in sexual maize induces the formation of
multiple embryo sacs in the ovule 57
3.2.6. Patterns of chromatin modification in dmt102 mutant lines mimic the effect of
apomixis during sporogenesis and gametogenesis 60
3.2.7. Transcription patterns in the mature ES and the early seed suggest quiescence in the
egg cell and the sexual zygote, but strong activity in the apomictic pro-embryos 62
3.2.8. Parthenogenetic pro-embryos have lost gametic identity, but have not established
embryonic patterning 65
3.3 Discussion 66
3.4 Methods 71
3.4.1. Plant Materials 71
3.4.2. Genotyping 71
3.4.3. Selection of CMEs for RT-PCR analysis 72
3.4.5. RT-PCR 72
3.4.6. Imaging of histone modifications and transcriptional activity in ovules and seed
tissues 73
3.4.7. Whole mount clearing of ovule samples 74
3.4.8. Whole mount in-situ mRNA hybridizations 74
3.5. Tables 75
3.6. Supplemental tables and figures 78
CHAPTER 4. Discussion and Perspectives 90
ACKNOWLEDGMENTS 95
REFERENCES 97
8

ABBREVIATIONS
AdoMet: S-adenosyl-L-methionine
AGO1: ARGONAUTE1
AGO4: ARGONAUTE 4
Arg/R: Arginine
ATO: ATROPOS
ATP: Adenosine Triphosphate
ChIP: Chromatin ImmuniPrecipitation
ChromDB: Chromatin Database
CLO: CLOTHO
CME: Chromatin Modifying Enzymes
CMT: Chromomethyltransferases
CMT3: CHROMOMETHYLASE 3
DAP: Days After Pollination
DCL3: DICER-LIKE 3
DDM1: DECREASE IN DNA METHYLATION1
DMT: DNA methyltransferases
DMT102: DNA Methyltransferase 102
DMT103: DNA Methyltransferase 103
DMS3: DEFECTIVE IN MERISTEM SILENCING3
DRD1: DEFECTIVE IN RNA-DIRECTED DNA METHYLATION1
DRM: Domains Rearranged Methyl Transferases
DRM2: DOMAIN REARRANGED METHYLTRANSFERASE 2
dsRNA: double strand RNA
ES:
Embryo Sac
FIE1: FERTILIZATION INDEPENDENT ENDOSPERM1
FIS:
FERTILIZATION INDEPENDENT SEED
FIS2: FERTILIZATION INDEPENDENT SEED2
H1: Histone 1
H2A: Histone 2A
H2B: Histone 2B
H3: Histone 3
H4: Histone 4
9

HAT: Histones Acetyltransferase
HDAC: Histone Deacetylase
HDA6: HISTONE DEACETYLASE 6
H3K9: H3 methylation lysine 9
HME: Histone Modifiers Enzymes
HMT: Histone Methyltransferases
K: Lysine
KYP: KRYPTONITE
LIS:
LACHESIS
Lys:
Lysine
MEA: MEDEA
MET1: METHYLTRANSFERASE1
m5C: Cytosine 5 methylated
MMC: Megaspore Mother Cell
MOM: MORPHEUS MOLECULE 1
PCR2: Polycomb Complex 2
PKL: PICKLE
RdDM: RNA-dependent DNA Methylation
RDR2: RNA-DEPENDENT RNA POLYMERASE 2
RDR6: RNA-DEPENDENT RNA POLYMERASE 6
RPD1: RNA POLYMERASE D1
RT-PCR: Reverse Transcription Polymerase Chain Reaction
SDC: SUPRESSOR of drm1drm2cmt3
SAM: S-adenosyl-L-methionine
SDE3: SILENCING DEFECTIVE 3
Ser:
Serine
SET:
Su(var)3-9, E(Z) and Trithorax
SGS3: SUPPRESSOR OF GENE SILENCING 3
siRNA: small interfering RNA
SUVH: Su(var)3-9 Homologs
SYD: SPLAYED
10

CHAPTER 1. General Introduction

The unique plants life strategy, with alterning sporophytic and gametophytic generations,
requires the correct activation or repression of transcriptional programs specific to each
developmental stage. In plants and other eukaryots, developmental programs imply preestablished
transcription patterns mediated through an epigenetic cellular memory, which is
controlled by dynamic changes or alterations at the chromatin structural level (Hsieh and
Fisher, 2005). These processes have been extensively studied at the mechanistic level. For
example, in flowering plants, Polycomb group proteins are involved in the epigenetic control
of genes throughout a repressive Polycomb complex 2 (PCR2) regulating vegetative and
reproductive programs (Leroy et al., 2007; Review in Pien and Grossniklaus, 2007). Although
it is also known that significant transcriptional changes occur during the switch between the
diploid sporophytic and haploid gamethophytic generation (Poethig, 2009), a key remaining
question is to define the mechanisms controlling the orderly transitions during the
reproductive process. In both Arabidopsis and maize, in particular, chromatin remodeling is
involved in controlling the orderly progression of vegetative development; however their
potential role during female meiosis, gametogenesis and fertilization remains poorly
understood. Here, we studied the epigenetic mechanisms controlling developmental
transitions during reproduction in flowering plants. The results also shed light onto the
molecular and cellular events leading to asexual reproduction in apomixis, a promising highvalue
trait for crops biotechnology.
1.1. BACKGROUND INFORMATION
1.1.1. Developmental Events During Sexual Plant Reproduction
The reproductive cycle of flowering plants occurs in specialized reproductive organs, the
anthers and the ovules. There, sexuality depends on the development of an haploid
(gametophytic) generation in which haploid gametes are formed by means of two consecutive
steps: sporogenesis (spore formation) and gametogenesis (gamete formation). The
gametophytic generation consists of specialized multicellular structures called gametophytes,
whose formation arises from cell fate changes turning somatic sporophytic cells into
primordial germ cells. The male gametophytes (or pollen grains) develop into the anthers. In
the archesporial tissue of the anther, a large number of microspore mother cells differentiate
(Figure 1A). Each of those cells undergoes a meiotic division to produce four haploid
12

microspores. All four meiotic products then go through two consecutive mitoses to produced
two spermatic cells enclosed into the cytoplasm of a vegetative cell, that collectively form the
mature pollen grain (McCormick, 1993). In the ovule, megasporogenesis or female
sporogenesis is initiated with the differentiation of the megaspore mother cell (MMC) within
the nucellus (Figure 1B).
Figure 1. Sporogenesis and gametogenesis in male (A) and female (B) reproductive organs of
flowering plants.
In the male reproductive organs, sporogenesis initiates with the differentiation of numerous pollen
mother cells (PMCs) within the tapetum of the anther; female sporogenesis in the ovule takes place in
a single cell, the megaspore mother cell (MMC). The outcome of meiosis in both PMCs and MMCs is
a tetrad of micro- and megaspores, respectively. A) Development of the male gametophyte (taken
from McCormick 2004). In the male organs, each free microspore undergoes mitosis I, producing the
bicellular pollen grain. The second mitosis gives rise to a tri-cellular mature pollen grain containing
two cell types, a vegetative cell and two gametes or spermatic cells. (B) Female megagametogenesis
occurs only in the functional megaspore, and three consecutive mitosis give rise to the mature embryo
sac or female gametophyte.
Only a single MMC differentiates in each ovule, and undergoes meiosis. As a result of
meiosis, four haploid megaspores are formed, three of them degenerate and only one, at the
chalazal pole, differentiates as a functional megaspore to give rise to the embryo sac (ES) or
female gametophyte. In the Polygonum type of ES development, the most common in
13

angiosperms (including maize and Arabidopsis) the nucleus of the functional megaspore
undergo three rounds of free nuclear divisions to produce eight haploid nuclei. Shortly after,
these nuclei initiate cellularization, which is concomitant to the acquisition of cell fate. At the
end of gametogenesis, the resulting mature ES contains seven cells: two synergids cells, three
antipods and two kinds of female gametes, the haploid egg cell, and the homo-diploid (or dihaploid)
central cell, resulting from the fusion of the two haploid polar nuclei (Figure 2).
At fertilization, the diploid sporophytic generation is reestablished after the pollen tube
reached the embryo sac and the double fertilization takes place. The two spermatic cells are
released into the ES; one fuses with the egg cell and the other fuses with the central cell,
producing the diploid zygote and the triploid endosperm respectively. Then, the fertilized
ovule gives rise to the seed by the synchronized growth of the embryo, the endosperm, and
the surrounding maternal tissues.
Ovary
Synergid
Mature Embryo Sac
Antipods
Central Cell
Synergid
Egg Cell
Figure 2. Schematic representation of
a Polygonum type mature embryo
sac.
In the ovary, a mature ovule containing
the cellularized embryo sac. Two
synergids together with the female
gamete form the egg apparatus. The egg
cell contains a prominent vacuole
localized toward the micropylar pole.
Three antipods are differentiated at the
chalazal end while in the center the binucleated
central cell constitutes the
biggest cell of the mature embryo sac.
1.1.2. Acquisition of Cell Fate in the Ovule and the Female Gametophyte
Contrary to animals, which set aside their germ cells early during embryonic development
by establishing clearly defined germ lines (Seydoux and Braun, 2006; Nokamura and
Seydoux, 2008), plants define their reproductive cells late during development, and do not
have germ-lines as such (Walbot and Evans, 2003). Rather, somatic cells switch
14

developmental programs to produce reproductive cells. The resulting Polygonum type of
female gametophyte has four cell types showing particular attributes, and fulfilling
specialized functions during the reproductive process (Figure 2).
One egg and two synergids form collectively the egg apparatus, which is localized at the
micropylar end of the ovule. Cytological descriptions of the egg reveal a highly polarized
cytoplasm content. While ultrastructural descriptions of the egg cytoplasm vary among
species, a relative physiological quiescence is consistently detected in the unfertilized eggs
(Huang et al., 1992; Russell, 1993). The egg is flanked by two synergids cells that function in
the attraction of pollen tube and constitute the target site for spermatic cells delivery
(Higashiyama, 2002; Marton et al., 2005). Synergids differ in size, cytoplasm, and nucleus
polarity with respect to the egg cell. While in some species they are indistinguishable from the
egg, in others species, like maize, mature synergids are characterized by a filiform apparatus,
a cell ingrowths with unclear function. The central cell is the largest cell in the embryo sac,
and a big vacuole fills almost 80% of its volume. The central cell contains two haploid nuclei
that may or may not be fused before fertilization, but are invariably positioned close to the
egg cell. Finally, three antipodal cells are localized to the chalazal end of the ovule (opposite
side from the micropyle). They display particular characteristics like a dense cytoplasmic
content and small vacuoles and nuclei. Cytochemical data also reveals high metabolic activity
but no clear function has been assigned to these cells. In some species they degenerate before
fertilization but in others, for example maize, they proliferate before fertilization. Antipods
and synergids cells are ephemerals, they collapse and degenerate during early stages of seed
development, presumably by following a (not well understood) cell death program (Wu and
Cheung, 2000).
The mechanisms determining the transition from somatic to germinal fate, and the number
of germ cells in the male and female archesporial tissues is not understood yet, even though
some mutants developing more than one MMC have been described in Poaceae (Nonomura et
al., 2003; Sheridan et al., 1996). Cell fate determinism in the female gametophyte is better
defined. It has been shown that it depends on nuclei positioning along its chalazal-micropilar
axis, whose establishment is linked to auxin distribution within the gametophyte, the auxin
gradient acting as a morphogen (Pagnussat et al., 2007; Pagnussat et al., 2009). While these
results suggest a model for gametophyte patterning in Arabidopsis, the mechanisms regulating
this intra-cellular polarization of auxin, and the acquisition of individual cell identity in the
embryo sac remain unclear. Arabidopsis mutants affecting the LACHESIS (LIS) gene switch
15

central cell to egg cell identity. LIS, and two other mutants in the same class, CLOTHO (CLO)
and ATROPOS (ATO) are also necessary for the restricted expression of egg- and central-cell
fate. All three genes are core spliceosomal components, suggesting that pre-mRNA splicing
mechanisms regulate temporal and spatial cell fate in the female gametophyte (Grob-Hardt et
al., 2007; Kägi and Grob-Hardt, 2007; Moll et al., 2008). The relationship between
spliceosome activity and cell identity is unclear, though. Collectively, our understanding of
cell fate determinism during reproductive development is thus extremely limited.
1.1.3. Apomictic Developments
Bi-parental sexual reproduction is overly dominant in flowering plants. Alternatively, some
plant species have modified their reproductive process and acquired means of reproducing
asexually. One such strategy is apomixis, a natural process of cloning by seeds (Nogler,1984).
In apomictic plants, female germ cells avoid the two hallmark components of sexual
reproduction, meiosis and fertilization (Figure 3). This results in the production of embryos
that are genetically identical to their mother plants.
There are two main apomictic mechanisms (reviewed in Nogler, 1984; Grimanelli et al.,
2001; Koltunow and Grossniklaus, 2003), involving either somatic (sporophytic apomixis)
and/or reproductive cells (gametophytic apomixis). In sporophytic apomixis, somatic cells in
the mature ovule acquire the ability to produce adventitious embryos, without the formation
of a gametophytic generation. The embryo arises spontaneously from the nucellar cells during
early stages of the sexual embryo development. Like zygotic embryos, survival of the
adventitious embryos depend of the normal growth of an endosperm and therefore, most, if
not all of these embryos co-exist with sexual development within the seed, and emerge in
positions adjacent to the sexually derived ES.
Two different manners of gametophytic apomixis, apospory and diplospory, have been
described in diverse apomictic species. Both apospory and diplospory develop an unreduced
embryo sac, originated from either a megaspore mother cell (diplospory) or a somatic cell
from the ovule (apospory). In the diplosporous type of apomixis, which occurs for example in
Tripsacum dactyloides (Leblanc et al., 1995), a wild relative of maize, or Boechera holboellii
(Naumova et al., 2001), that is related to Arabidopsis, the MMC differentiates but fails to
initiate or complete meiosis. Instead, meiosis is substituted by apomeiosis, a mitotic-like
division characteristic of apomicts. These apomeiotic MMCs, or derivatives, acquire the
identity of a functional megaspore, and similarly to sexual plants, they follow a wild type
16

megagametogenesis to produce a Polygonum type of embryo sac. Cell fate and organization
in the apomeiotic embryo sac is broadly similar in sexually and apomictically derived
gametophyte. However, specific abnormalities, such as lack of polarization during
megagametogenesis, number and/or size of free nuclei, inverted embryo sacs or abnormal
location of the egg and central cells are frequently found in apomicts (Bicknell and Koltunow,
2004). Apomeiosis is often considered as a female-specific reproductive phenotype, but
unreduced pollen grains formation has also been reported in apomictic Boechera (Schranz et
al., 2006) and Tripsacum (Grimanelli et al., 2003), suggesting that male and female gametes
are affected in similar ways.
At the end of the formation of the diplosporous female gametophyte, autonomous divisions
of the unfertilized egg cell produce a parthenogenetic pro-embryo, while the unreduced
central cell either requires (pseudogamy) or does not require (autonomous) fertilization to
initiate endosperm development. In individual ovules, diplosporic development recruits the
MMC to apomixis, and thus excludes sexual embryo sac formation. In some species,
however, displosporous development coexists with aposporic embryo sac formation (Bonilla
and Quarin, 1997).
Aposporic embryo sacs, by contrast, originate from somatic cells, and therefore often
coexist with a sexual embryo sacs derived from the MMC. Here, megasporogenesis is omitted
and one or more functional megaspores develop directly from somatic cells in the nucellus
(aposporous initials), arising adjacent to the meiotically reduced megaspore. As in diplospory,
the aposporous initials proceed normally through megagametogenesis, and several unreduced
embryo sacs are seen in individual ovules. The structural abnormalities found in diplospory
are usually found with a higher frequency in apospory. Similarly, the embryo develops
parthenogenetically and the endosperm is either pseudogamous, or autonomous.
In most, if not all documented cases, apomixis and sexuality are not mutually exclusive
since they usually coexist in the same ecotypes with varying levels of relative expression
(Nogler, 1984; Bicknell et al., 2003; Barcaccia et al., 2006). Similarly, in the case of
gametophytic apomixis, the formation of multiple embryo sacs is a hallmark of aposporous
apomixis, but some diplosporous plants also develop extra embryo sac-like structures. This
collectively suggests that the frontier between both types of apomixis, and between apomixis
and sexually, is relatively blurry, fueling the perception that apomixis might be a deregulated
form of sexual reproduction rather than a new function (Grimanelli et al., 2001; Grimanelli et
17

al., 2003; Tucker et al., 2003; Koltunow and Grossniklaus, 2003; Ozias-Akins and Van Dijk,
2007).
Figure 3. Alterations of the sexual reproductive pathway in the different apomictic routes
(from Bicknell and Koltunow, 2004). The typical progression of the sporophytic and gametophytic
generations are schematically represented during the life cycle of sexual plants (yellow line). The
gametophytic routes of apomixis, diplospory (purple line) and apospory (red line) bypass both meiosis
(megasporogenesis) and double fertilization but they follow the successive developmental steps of the
life cycle. In the sporophytic type (or adventitious embryony; green line), somatic embryos are formed
from somatic cells of the ovule and develop alongside the normal, reduced, female gametophyte..
While bi-parental sexual reproduction is overly dominant, in both plants and animals, it
also entails short-term disadvantages over asexual reproduction (Partridge and Hurst, 1998).
Chiefly, obligate meiosis in sexually reproducing organisms prevents the perpetuation of
phenotypes with exceptional fitness. In asexual reproduction, by contrast, organisms
perpetuate themselves without need of a partner, leading to offspring that are genetically
identical. This provides a strong selective advantage to exceptionally fit asexual ecotypes.
The evolutionary theory, however, also predicts that the long-term cost of asexuality in
wild population will erase its immediate benefits. In controlled environments or experimental
populations, however, cloning has clear values, be it in medicine (therapeutic cloning), and
agriculture (plant and animal products). The potential applications of apomixis in agriculture
18

are quite obvious in terms of plant breeding and seed production methods, and have been
discussed in details in several reviews (for example Spillane et al., 2001; Spillane et al.,
2004). However, although apomixis occurs in more than 400 angiosperm species, it is not
found in major agriculture crops, and attempts to introduce or engineer apomixis in otherwise
sexual plants have proved difficult.
The work described in this document uses maize, and Tripsacum, as its key experimental
systems. Only few agriculturally important species have close apomictic relatives (Bicknell
and Koltunow, 2004). This includes maize (Zea mays), a crop which has a wider range of uses
than any other cereal, and constitutes the staple food in many American and African countries.
In Mexico, maize is in fact much more than food or feed. Rather, it has been at the root of its
cultural heritage since the Maya and Aztec civilizations (Mangelsdorf, 1974), and is an
essential component of the pre-colonial cosmogony. Tripsacum is an apomictic wild relative
of maize, endemic of Mesoamerica. Both species are genetically quite close, and apomictic
maize-Tripsacum hybrid plant can be obtained relatively easily (Leblanc et al., 1996).
Apomictic hybrids obtained with maize and Tripsacum reproduce by diplosporous,
pseudogamous gametophytic apomixis. Giving the wealth of genetic, genomic and cell
biology tools available in maize, such combination of a good experimental system together
with a close apomictic relative provides an attractive model to study apomixis. Our group has
been using this model for a long time, and contributed some interesting data regarding the
genetic control of apomixis (Leblanc et al., 1995; Leblanc et al., 1996; Leblanc et al., 2009;
Grimanelli et al., 1998a; Grimanelli et al., 1998b; Grimanelli et al., 2001; Grimanelli et al.,
2003; Grimanelli et al., 2005).
1.1.4. The Genetic Control of Apomixis
It is widely accepted that apomixis is under genetic control. The inheritance of apomixis
has been studied using segregating population in diverse species. The results indicate
commonalities, but also divergences (Reviewed in Ozias-Akins and Van Dijk, 2007).
Inheritance and mapping studies in aposporic species such as Pennisetum (Ozias Akins et al.,
1998), Paspalum (Martinez et al., 2001) and Hieracium (Bicknell et al., 2000) indicate that
apomixis segregates as a single dominant genomic region, suggesting monogenic inheritance,
or at least a limited number of dominant alleles. However, in other species such as Taraxacum
officinale, or Poa pratensis, segregation of the different components of apomixis imply
several independent genes contributing to the entire apomictic process (van Dijk et al., 2003;
19

Matzk et al., 2000). Furthermore, studies in Tripsacum (Grimanelli et al., 1998a), later
confirmed in other species, including Pennisetum (Ozias-Akins et al., 1998) and Paspalum
(Martinez et al., 2003), indicate that the chromosomal segment co-segregating with apomixis
is likely a large block of non-recombining DNA, presumably consisting mostly of gene-poor
heterochromatin (Conner et al., 2008). Overall, our understanding of apomixis from a genetic
standpoint remains thus extremely limited.
Interestingly, some studies showed that apomixis and sexuality share key regulatory
mechanisms (Tuckey et al., 2003). Thus, it has been proposed that apomixis could result from
a temporal or spatial deregulation of the transcriptional programs controlling sexual
reproduction (Koltunow and Grossniklaus 2003; Grimanelli et al., 2003; Bicknell and
Koltunow, 2004; Bradley et al., 2007). The model postulates that sexual reproduction involves
several transitions (from somatic cells to reproductive cells, from sporogenesis to
gametogenesis, from gametogenesis to embryogenesis) whose orderly progression is altered
in apomicts, resulting in heterochronic expression of the core developmental programs.
Several comparative transcriptome profiling experiments have been carried out between
sexual and apomictic plants with the aim of recovering «switches» differentiating sexual and
apomictic reproduction (Albertini et al., 2004; Grimanelli et al., 2005; Sharbel et al., 2009).
The results showed that only an surprisingly reduced proportion of the global transcript
populations are differentially expressed between sexual and apomictic plants. Indeed, only
few candidate genes have been identified, and their involvement in the control of apomixis
remains largely speculative.
Thus, neither genetic nor profiling approaches have helped much unlocking the molecular
determinants of apomixis. By contrast, however, the analysis of sexual Arabidopsis mutants
exhibiting phenotypes that resemble apomixis in some ways, and particularly diplospory, have
generated interesting candidates, possibly involved in regulating key elements of apomixis
(Ravi et al., 2008; d’Erfuth et al., 2009). Still at early stages, these alternative approaches,
toward the engineering of de novo versions of apomixis by combining mutant phenotypes in
sexual model species, promise an interesting alternative for manipulating apomixis.
20

1.1.5. Role of Hybridization and Polyploidy in Apomicts
The natural occurrence of apomixis has been linked to so-called «genomic shocks», that is,
genome wide changes in genome structure and dynamic, following major disruptive events,
such as interspecific hybridization, or changes of polyploidy. The argument was initially
suggested by the fact that most apomictic species are polyploid and highly heterozygous,
often of interspecific origin (Carman, 1997). The role of ploidy response or hybridization as
causes or consequences of apomixis however, have never been fully, functionally tested.
The close similarities between sexual and apomictic reproduction were emphasized in an
evolutionary theory of apomixis proposed by J. Carman (1997), “the duplicate-gene
asynchrony hypothesis”. This hypothesis postulates that apomixis result from the
hybridization of related species showing differences in reproductive behavior. Carman
suggested that specific combinations of hybrid genomes may result in unstable expression of
reproductive traits, and consequently, alterations in the temporal and/or spatial patterns of
reproductive genes expression. Indirect support for this hypothesis can be found in hybrids of
sexual progenitors displaying distinct reproductive anomalies as a result of changes in genes
expression patterns following for example allopolyploid formation (Chen and Ni, 2006; Chen,
2007; Chen et al., 2008). Similarly, abnormal meiotic behavior can be found in autotetraploids
(Carvalho et al., 2009), and heterochronic expression of reproductive traits is a common
observation in sexual diploid Tripsacum accessions (Bradley et al., 2007). More recently it
has been proposed that mechanisms affecting the fate of orthologous and homoeologous
genes in allopolyploids could provide a flexible mean to respond to polyploid and genomic
shock in hybrids (Chen, 2007; Gaeta et al., 2007; Gaeta et al., 2009), allowing polyploid
hybrids to adapt to new reproductive strategies, possibly including apomixis. This places
hybridization and ploidy as consequences rather than cause of apomixis.
1.1.6. Epigenetic, Cell Fate and Apomixis
Underlying all the previous assumptions, clearly, is the concept that genomic shocks acts to
modify the epigenetic background in which reproductive development operates. Starting with
the landmark characterization of the FIS (FERTILIZATION INDEPENDENT SEED) class of
genes in Arabidopsis (reviewed in Baroux et al., 2007, Autran et al., 2005), the idea that
epigenetic-level regulation might play a critical role in differentiating apomixis and sexual
reproduction has gained considerable traction. The FIS class of genes MEA (MEDEA;
Grossniklaus et al., 1998), an enhancer of Zeste-like SET-domain protein, FIE1
21

(FERTILIZATION INDEPENDENT ENDOSPERM, Ohad et al., 1999), WD-40 repeat protein,
and FIS2, a Zinc-finger transcription factor (Luo et al., 1999), were initially identified in
genetic screens for apomixis-like behavior. They are involved in the regulation of cell division
in the female gametophyte by the Polycomb Group complex, to whom both MEDEA and FIS2
belong, and their mutant alleles produce partial seed development in absence of fertilization.
Both MEDEA and FIS2 are imprinted, that is, show parent of origin specific expression.
Collectively, they offered the first clues that epigenetic regulators were important to seed
development.
Epigenetic control has been known for a long time to play a crucial role in defining cell
fate in both plants and animals (Bernstein et al., 2006; Buszczack and Spradling, 2006; Costa
and Shaw, 2006; Caro et al., 2007; Hajkova et al., 2008; Gereige and Mikkola, 2009). Such
epigenetic regulation involves changes in gene expression without changes in DNA sequence,
and depends on stable, mitotically (and eventually meiotically) heritable marks on DNA or
chromatin (see below). In the ovule as in any organs, the definition of each cell lineage is the
consequence of a finely tuned control of “on” and “off” gene expression states, a process
which is intrinsically epigenetic. Conceptually, at least three transitions take place during
reproductive development involving changes in cell identity, and each could differ between
apomictic and sexual plants: (1) the transition from the soma to the germ cells, at the
beginning of megasporogenesis, (2) the transition between the meiotic products and the
functional megaspore, at the beginning of megagametogenesis, and (3) the transition from the
gamete to the zygote, at the initiation of embryo development.
A phenomenon akin to transdifferentiation (the conversion to a cell or one tissue lineage
into a cell of a completely different lineage; Costa and Shaw, 2006) has been proposed for a
long time to explain apomixis (Peacock, 1992; Carman, 1997). It offers an explanation for the
cellular fate of apomeiotic MMCs, or the direct determination of an unreduced functional
megaspore from the nucellar cells, and eventually, the triggering of parthenogenesis in the
unreduced germ cell.
So could the orchestration of the transition between these different states by epigenetic
means play a role in defining apomixis? Recent studies have clearly demonstrated that indeed,
epigenetic mechanisms have a crucial role in developmental transitions during vegetative
growth (Poethig, 2003; Peragine et al., 2004; Bezhani et al., 2007; Poethig, 2009). Similarly,
several lines of evidences suggest that transitions during reproduction and early seed
development are epigenetically controlled through dynamic changes in chromatin state
22

(Huanca-Mamani et al., 2005; Xiao et al., 2006; Baroux et al., 2007; Curtis and Grossniklaus,
2008). The hypothesis is clearly attractive, but whether alterations in the epigenetic regulation
orchestrating these transitions converge in a cellular transdifferentiation, and whether they are
involved in the differentiation between apomictic and sexual reproduction is currently
unknown.
1.1.7. Mechanisms of Chromatin Dynamics
Chromatin represents the compacted form of genomic DNA in the nucleus and its basic
unit is the nucleosome (Figure 4A). The nucleosome is composed by a core particle consisting
of an octamer of histones, that are surrounding by 147 pb of DNA wrapped in 1.65 turns. The
octamer contains two molecules each of the highly conserved histone proteins, H2A, H2B, H3
and H4. Toward their C termini region, the histones have fold domains of about 70 amino
acids, which mediate the interactions between the core of histones and between histones and
DNA. The nucleosomes are interconnected by a linker of 10-80 pb DNA. The H1 histone
connects the outside region of the core particle and interact with about 20 pb of linker DNA.
Its function is associated with the modulation of chromatin compaction (Clausell et al., 2009).
The N terminal region of the core histones has lysine-rich tails which are highly conserved
between species and are the target sites of a high variety of covalent modifications. These tails
play an important role in the highest-order of chromatin structure, as their covalent
modifications altering significantly the stability of the nucleosome. The combination of these
covalent modifications represent a key feature of epigenetic marking, and is often referred to
as the «histone code» (Strahl and Allis, 2000; Jenuwein and Allis, 2001), which plays a
functional role in nucleus organization, the control of transcription, and the definition of cell
fate (Lee et al., 2006; Costa and Shaw, 2007; Earley et al., 2007; Corpet and Almouzni, 2009;
Hublitz et al., 2009; Mohn and Schübeler, 2009).
Two distinct states of chromatin are traditionally defined (although these states are in fact
highly dynamic structures), euchromatin and heterochromatin (Figure 4B). Heterochromatin
is a highly condensed state of chromatin, associated to late replication, mainly localized at
centromeres and telomeres. It usually presents a low gene content, a high content of repetitive
sequences, reduced meiotic recombination and low transcriptional activity. In Arabidopsis,
the factors implicated in heterochromatin formation are repetitious DNA, DNA methylation,
H3 methylation on lysine 9 (H3K9), and small RNAs (Grewal and Moazed, 2003; Grewal and
Elgin, 2007; Grewal and Jia, 2007; Hale et al., 2009; Hublitz et al., 2009). Genes found in
23

constitutive heterochromatin are usually uniform in their nucleosome array and can remain in
an inactive state as heterochromatin permanently. By contrast, facultative heterochromatin
can switch to a euchromatic state and allow gene expression, by removing the histones
modifications and the proteins associated with the inactive state. The euchromatin is a generich
chromatin, less condensed, localized at the chromosome arms, is enriched in unique
sequences, replicated throughout S phase, broadly constitutes the transcriptionally active
component of the genome, and recombines during meiosis (Grewal and Elgin, 2007).
Chromatin structure represents a highly dynamic system in which arrays of nucleosomes
are disrupted, modified or removed by the chromatin remodeling factors (Jerzmanowski,
2007), and histone proteins modified by different chemical groups. Temporal and/or spatial
chromatin modifications provide a variety of mechanisms to control switch during cell
differentiation and development (Corpet and Almouzni, 2009). Such dynamic state, which is
determined by chromatin modifying enzymes (CME), tunes the accessibility of the DNA
molecule to a high variety of regulatory enzymes, including transcription factor (Greval and
Moazed, 2003). Genome-wide analysis in both plants (Zilberman et al., 2007) and animals
(Schübeler et al., 2004, and numerous references since then) have confirmed the
interdependent relationship between dynamic chromatin structures and transcription. Here, we
will briefly summarize the state of the art regarding two key modifications that are important
to this work, DNA methylation and Histone tail modifications. The field is an extremely
dynamic one, and this presentation is necessarily partial.
24

A
B
Heterochromatin
Me
Me
Me
Transcription
repression
Euchromatin
Ac
Ac
Ac
Transcription
activation
Figure 4. Chromatin Structure. A) Genomic DNA is packaged into the nucleosomes; an individual
nucleosome is composed by the core of histones which is formed by two molecules each of H2A,
H2B, H3 and H4. The histone octamer together with 147 bp of DNA and other proteins constitute the
chromatin structure; image taken from Morgan, 2008. B) The genome is organized into different
compartments, corresponding to different levels of chromatin organization.
1.1.8. Modifying Enzymes and Chromatin Dynamics
1.1.8.1. DNA methylation
In eukaryots, cytosine residues are modified by methylation, one of the most studied
epigenetic mechanisms (for a textbook: Allis, Jenuwein and Reinberg, 2007). The process
involves the covalent addition of a methyl group to the cytosine five (C5m) by DNA
methyltransferases, which transfer a methyl group from S-adenosyl-L-methionine (AdoMet)
to cytosine residues. DNA methylation occurs at genes sequences, repeated and transposon
sequences. Methylation at 5’ control regions is generally associated with transcriptional
repression and/or silencing of genes. Biological consequences of loss of DNA methylation
include arrest of embryo development, switch-on of apoptotic effects, and usually lethality in
mammals. Numerous effects on morphology and development have been also reported in
plants (Gehring and Henikoff, 2007).
25

While many plant and animal species use DNA methylation as an epigenetic mark, plant
DNA methylation has unique features. DNA methylation in plants takes place on all three
possible cytosine contexts: CG, CHG and CHH, where H represent A,T, or C. Repeats in
general are highly associated with non-CG methylation (but also CG), while genes are
characterized by CG (and limited non-CG) methylation. The role of CG methylation in the
coding region of the genes is not absolutely clear.
In Arabidopsis, where the process is best understood, at least three classes of DNA
cytosine methyltransferases (DMTs) intervene in methylation pathways: DRM, CMT, and
MET. DRM1 and DRM2 (DOMAINS REARRANGED METHYL TRANSFERASE1 and 2) take
part in the so called RdDM (RNA-directed DNA methylation) pathway and are the main de
novo DMTs involved in all sequence contexts (Cao and Jacobsen, 2002). DRMs also play a
role, together with CMT3 (CHROMOMETHYLTRANSFERASE3), in the maintenance of
methylation at non-CG sites (Cao et al., 2003) and are required for maintaining genes
silencing (Lindroth et al., 2001). Curiously, it is only in a triple drm1drm2cmt3 mutant that a
loss-of-function of DRM genes have any phenotypic effects. The effects results from the loss
of non-CG methylation at the endogenous gene SUPRESSOR of drm1drm2cmt3 (SDC;
Henderson and Jacobsen, 2008). Phenotypic effects of CMT3 are mild, and only detected
during early embryogenesis (Xiao et al., 2006). Finally, MET1 (METHYLTRANSFERASE1) is
involved in maintaining DNA methylation at CG dinucleotides, is required to maintain
methylation patterns during male and female gametogenesis (Saze et al., 2003; Saze, 2008)
and is essential for transgenerational stability of epigenetic information (Mathieu, et al.,
2007). Additionally, DDM1 (DECREASE IN DNA METHYLATION1), a SWI2/SNF2-like
chromatin-remodeling factor, also participates in the maintenance of methylation at both CG
and non-GC sites (Jeddeloh et al., 1999). Mutations in DDM1 cause a loss of both DNA and
H3K9 methylation in somatic tissues (Johnson et al., 2002; Gendrel et al., 2002; Schoft et al.,
2009), and play a crucial role in maintaining genome integrity in the sperm cells during
genome reprogramming (Slotkin et al., 2009; Schoft et al., 2009). Remarkably, gametophytic
effects in DRM1, DRM2, CMT3 or DDM1 have not been reported (Saze, 2008), which would
suggest that DNA methylation is fully dispensable during gametogenesis, or, alternatively,
that an additional, reproduction-specific pathway complements the «somatic» (DRM2, CMT3
dependent) pathways.
Collectively with DMTs, small interfering RNAs (siRNAs), histone modifying enzymes
and RNAi proteins, the RdDM pathway is involved in mediating both DNA and histone
26

methylation (Huettel et al., 2007). This mechanism establishes de novo DNA methylation in
all sequences contexts (Matzke at al., 2007; Saze, 2008), protecting the genome from
transgenerational epigenetic defects (Texeira et al., 2009). In Arabidopsis RdDM initiate
when siRNA are generated by the action of the putative DNA-directed RNA polymerase Pol
IV, RDR2 (RNA-dependent RNA polymerase 2), and DCL3 (DICER-LIKE3) (Matzke et al.,
2009). Then, the siRNAs are loaded into ARGONAUTE proteins (AGO4 mainly) to direct
DNA methylation by DRM2 (Figure 5). In addition, others components as an SNF2-like
chromatin remodeling protein DRD1, and an SMC-like protein DMS3 (Kanno et al., 2008),
play roles in dsRNA-induced de novo methylation at non-CG contexts (Saze, 2008). The
silencing effect of the RdDM pathway is reinforced by the establishment of repressive histone
marks, in particular methylation of lysine 9 on histone H3 (H3K9me), by the activity of KYP
(KRYPTONITE), the main H3K9 methyltransferase in Arabidopsis (Jackson et al., 2002), and
HDA6 (HISTONE DEACETYLASE 6) (Probst et al., 2004).
1.1.8.2. Histone modifications
The amino-terminal tails of histones is the second major covalent regulatory system of
chromatin controlling genes expression (see Allis et al., 2007, for detailed informations).
Histone tails may be modified by acetylation, methylation, phosphorylation, ubiquitination, or
SUMOylation (Figure 6). Patterns of these post-translational modifications in the
nucleosomes constitute a histone code, which defines chromatin states as either conducive
(permissive) or inhibitory (repressive) to transcription (Jenuwein and Allis, 2001). Histone
modifications and DNA methylation are not independent processes, as the histone code is
characterized by a close interaction between both chromatin marks (Jackson et al., 2002). The
«complete» code also includes the effect of numerous other proteins. Their complete
description is beyond the scope of this introduction, and only a short summary is provided
below.
27

Arabidopsis thaliana
DNA methylation occurs at CG and non-
CG (CHG, CHH) sequences contexts
(yellow boxes). In Arabidopsis,
establishment or de novo methylation is an
exclusive activity of the DNA
methyltransferases D O M A I N S
REARRANGED METHYLTRANSFERASE
1 and 2 (DRM1/DRM2). Maintenance of
methylation requires not only DNA
methyltransferases but also other CMEs,
including at CG sites a histone deacetylase
( H D A 6 ) , a S W I / S N F c h r o m a t i n
remodeling DECREASE IN DNA
METHYLATION 1 (DDM1) and the DNA
m e t h y l t r a n s f e r a s e
METHYLTRANSFERASE 1 (MET1).
Maintenance of methylation at non-CG
RNAgenerating
pathway
RDR2
DCL3
RPD1
AGO4
RDR6
SGS3
SDE3
AGO1
Other?
RNA proteins
DNA methyltransferases
SWI/SNF chromatin remodeling ATP-depending
Histone methyltransferases
Histone deacetylases
Establishment
si RNA
DRM1
DRM2
CHH
CHG
DDM1
CG
Maintenance
DRM1/DRM2
CMT3
MET1
HDA6
si RNA
KYP
SUVH4
SUVH?
Figure 5. DNA methylation pathways in
sites involves CHROMOMETHYLASE 3
(CMT3) and DRM1/DRM2. CMT3 methylation is directed by KRYPTONITE (KYP) and other
unknown histone methyltransferases. All methylation pathways, but maintenance of methylation at
CG, involve siRNA generated from different branches of the RNAi pathway (dark blue boxes). AGO1,
ARGONAUTE1; AGO4, ARGONAUTE 4; DCL3, DICER-LIKE 3; RDR2, RNA-DEPENDENT RNA
POLYMERASE 2; RDR6, RNA-DEPENDENT RNA POLYMERASE 6; RPD1, RNA POLYMERASE
D1; SDE3, SILENCING DEFECTIVE 3; SGS3, SUPPRESSOR OF GENE SILENCING 3. siRNA,
small interfering RNA; SUVH, Su(var)3-9 homologs. Figure modified from Chan et al., 2005.
1.1.8.2.1. Histone acetylation
Acetyl groups are added by histones acetyltransferases (HATs), which catalyze the transfer
of the acetyl moiety acetyl-CoA to the E-amino group of lysine (Lys) residues (Lahm et al.,
2007). Histones can be acetylated at multiple sites. H4 for example is acetylated at Lys5,
Lys8, Lys12, and Lys16, and H3 is acetylated at Lys9, 14, 18, 23 and 27, with the list growing
rapidly. In the cell, the acetylation state can be dynamic and reflect the balance of HATs and
histone deacetylases (HDACs) activity. Together, HAT and HDAC play a key role in
transcriptional regulation, since strong correlation has been established using chromatin
Immuniprecipitation (ChIP) between transcriptional activity and the distribution patterns of
histone acetylation (Bhat et al., 2003; Ng et al., 2006; Earley et al., 2007).
28

1.1.8.2.2. Histone methylation
The histone methyltransferases (HMTs) are responsible for the transfer of methyl group
donor from S-adenosyl-L-methionine (SAM) to either lysine (Lys,K) or arginine (Arg/R)
residues. Lysines can be mono-, di- or three-methylated while arginines can be mono- and
dimethylated. These histone modifications have been correlated with both transcriptionally
silenced or active chromatin states, but the rules are complex. For instance, H3K9 methylation
is associated with silenced promoters on the inactive X chromosome in mammals, whereas
H3K4 methylation is found at active genes (Boggs et al., 2002). Changes in histone
methylation are mediated by the activity of histone demethylases acting in the coordinated
cross-talk of the chromatin states. KYP is the main H3K9 methyltransferase in Arabidopsis.
1.1.8.2.3. Phosphorylation
Histone phosporylation is mediated by kinases, and has been associated with chromosomal
condensation and segregation, and also with genes regulation. Phosphorylation is particularly
found in threonines, and serine (Ser) 10 and 28 of H3 where has been associated to
transcriptional activity. This modification is frequently codependent on histone acetylation.
Figure 6. Histone modifications
The figure illustrates the main known covalent modifications at the amino-terminal domain of the
histones. Some modification occurs also at the globular domain (blue sphere at H2A-H2B, and green
Me hexagone at H3). Active marks include acetylation (turquoise flag), arginine methylation (yellow
hexagon) and some lysine methylation (green hexagon). Repressive marks are represented as red
hexagons. Numbers below the protein sequence indicate the position of the modified aminoacid. Figure
taken from Allis et al., 2007.
29

In H4 it is only found in Ser1 (Berger and Gaudin, 2003). Dynamic phosphorylation is
provided by phosphatases which remove phosphate groups (Berger, 2007).
1.1.8.2.4. Ubiquitylation and SUMOylation
In contrast to Acetylation, methylation and phosphorylation which modify chromatin with
small chemical groups, ubiquitylation and SUMOylation add large moieties of around twothirds
the size of the histone proteins. Ubiquitylation forms an isopeptide bond with ubiquitin
at the histone lysine residues. Ubiquitylation is associated with transcriptionally active
chromatin state in H2B Lys 123, but it correlates with repressed chromatin in H2A Lys 119
(Berger, 2007). SUMOylation correlates with repressive chromatin and it seems to be
mutually exclusive of acetylated chromatin. This modification occurs in H2A and H2B where
it is correlated with transcription repression.
1.1.8.3. ATP-dependent chromatin remodeling
This chromatin modification is performed by a multiprotein complexes which modify
histone-DNA interactions using ATP hydrolysis. They destabilize nucleosome structure by
introducing superhelical torsion into DNA, leading temporal accessibility to other modifier
factors or specific enzymatic reactions. Although the Arabidopsis genome encodes more than
40 SNF2-like proteins, most of them have been no analyzed yet. However, some studies
carried out in PICKLE (PKL), DDM1, MOM and SPLAYED (SYD), suggest important roles
during plant development (Ogas et al., 1999; Vongs et al., 1993; Amedeo et al., 2000; Wagner
and Meyerowitz, 2002).
30

1.2. THESIS PROPOSAL: CHROMATIN DYNAMICS, AN UNEXPLORED FRONTIER TOWARD
UNDERSTANDING ASEXUALITY IN PLANTS
Results that we obtained and published prior to formally initiating this PhD program were
essential for the design of this project. These results suggested two important hypotheses. The
first point was that apomixis is the result of a spatial and/or temporal deregulation of the
orderly progression of developmental transitions that occurs during sexual reproduction.
(Grimanelli, D., Garcia-Aguilar, M., et al. 2003. Heterochronic expression of sexual
reproductive programs during apomictic development in Tripsacum. Genetics 165: 1521–
1531). The paper presents data suggesting the deregulation of sexual reproductive programs
during apomictic reproduction in Tripsacum, and the idea that apomixis might be a classical
case of heterochronic mutation.
The second point was that, similarly to vegetative development, the key developmental
transitions during reproduction might be mediated by epigenetic, chromatin-based regulatory
processes. (Huanca-Mamani, W., Garcia-Aguilar, M., et al. 2005. CHR11, a chromatinremodeling
factor essential for nuclear proliferation during female gametogenesis in
Arabidopsis thaliana. Proc. Natl. Acad. Sci. USA 102: 17231-17236). This paper presents
data suggesting a crucial role for chromatin-based regulation at the sporogenesisgametogenesis
transition.
The following document is therefore mostly the description of our efforts in trying to
connect the dots between both papers. The purpose in this work was to analyze the role of
chromatin dynamics in the control of developmental transitions in reproductive cells of sexual
and apomictic plants, and to identify candidate chromatin-level regulators of apomixis. To
achieve this, we addressed the three following questions:
(1) are the various developmental transitions characterized by specific chromatin states?
(2) are there differences in chromatin states between sexual and apomictic plants?
(3) could these variations be causal factors in determining whether a plant reproduces
sexually or apomictically?
31

In the second chapter, we describe a profiling experiment used to identify CMEs showing
differential expression in sexual and apomictic ovules. Out of 319 CMEs, we selected 56
candidates for further profiling during key developmental steps of sexual and apomictic
reproduction. This work led to the identification of a limited number of candidates out of
which two, DMT102 and DMT103, were further characterized using both genetical and
immunochemical approaches. The results of these functional analyses are presented in
Chapter 3, as a manuscript submitted for publication. In Chapter 4, we discuss the hypothesis
we put forward for a siRNA silencing pathway acting specifically in plant ovules and
controlling reproductive cells differentiation.
32

CHAPTER 2. Expression of Chromatin
Modifying Enzymes during Apomictic and sexual Development
33

2.1. INTRODUCTION
To summarize, we know that apomixis can be achieved through diverse, complex
pathways, all of which affect the orderly progression of the key developmental transitions that
take place during sexual reproduction: megaspore mother cell differentiation,
megagametogenesis and embryogenesis. Although several studies have documented the role
of chromatin-mediated control in genes expression during early seed development (reviewed
in Baroux et al., 2007), the role of chromatin in the earlier stages of reproductive development
(from the ovule primordium until the formation of the mature gametophyte) is poorly
understood. We used two sources of information to screen maize chromatin modifying
enzymes (CMEs) for candidates involved in the regulation of transitional changes during
sexual reproduction. The first one was the Chromatin Database (ChromDB, University of
Arizona; www.chromdb.org). The second one was a set of microarray data comparing mRNA
populations extracted from sexual maize and apomictic maize-Tripsacum ovules. Collectively,
we identified 56 candidate CMEs for further transcriptional analyses during plant
reproduction, with a particular emphasis on the key steps that differentiate sexuality in maize
from the apomictic pathway in Tripsacum.
2.2. RESULTS
2. 2.1. Selection of Chromatin Modifying Enzymes (CME) from ChromDB
ChromDB contains 358 expressed CME sequences belonging to 74 proteins families
(updated data as of April 2007; Table 2.2.1). Of these, 185 corresponded to fully sequenced
and annotated genes while only partial information was provided for the remaining 173 genes.
Expression data was available for 79 entries in maize, including RNA expression patterns for
several tissues (ear, tassel, kernel, seedling leaf, callus and mature leaf), and 23 genes for
which RNAi lines are available from the Maize Genetic Stock Center (McGinnis et al., 2007).
We used RNA expression patterns as the main criterion for selection, discarding genes
showing no expression in reproductive tissues. However, due to the growing evidence for a
regulatory role for chromatin in cell fate differentiation, and the hypothesis of a causal effect
of apomixis on somatic cell identity, we also retained genes specifically expressed in
undifferentiated callus tissues, assuming that the capability for totipotency might contribute to
apomixis expression (ex. CHR106, CHR120, HAG102). The figure 2.2.1 reproduces the
expression pattern of several selected genes as shown on the ChromDB site. Furthermore, as a
34

control for transcriptional behavior of constitutive genes into an apomictic background, we
also included genes highly expressed in all tissues except mature leaves, such as HDT103 and
NFA101 (Table 2.2.2). A total of 45 genes encoding CMEs, covering 15 of the 74 CME
families and showing interesting expression in reproductive tissues were selected (Table 2.2.1,
Table 2.2.2). Note that RNAi lines are available for 23 out of the 45 selected loci (Table 2.2.2
and www.chromdb.org).
Probe
1 2 3 4 5 6
BRAT 101
CHR106
NFE 101
SDG 122
Figure 2.2.1. Expression of several chromatin modifying enzymes in different maize tissues by
Northern analyses. Original pictures and experimental protocols can be found at www.chromdb.org.
1: ear, 2: tassel, 3: kernel, 4: seedling, 5: callus, 6: mature leaf.
35

Table 2.2.1. List of Chromatin Modifying Enzymes as provided by the Chromatin
Database. 1 Annotations of protein class according to ChromDB. 2 Genes number correspond
to total identified genes at ChromDB.
Protein Class 1 Genes 2 No. Genes with RNA data No. Selected genes
AGO 16 ND 0
ARP 6 ND 0
BRAT 1 1 1
BRD 4 2 1
CHB 4 2 1
CHC 2 1 1
CHE 1 ND 0
CHR 44 11 8
CPC 2 ND 0
CPD 1 ND 0
CPG 1 ND 0
CPH 1 ND 0
CRD 1 ND 0
DCL 4 ND 0
DMT 7 5 7
DNG 4 ND 0
DRB 6 ND 0
EBP 5 ND 0
EPL 1 1 0
GTA 3 ND 0
GTB 1 1 0
GTC 1 1 0
GTE 11 2 0
GTF 1 ND 0
GTG 1 ND 0
HAC 5 1 1
HAF 1 ND 0
HAG 6 3 1
HAM 2 ND 0
HCP 2 ND 0
HDA 10 4 2
HDMA 3 ND 0
HDT 4 4 3
HEN 1 ND 0
HF0 15 ND 0
HIRA 2 ND 0
HMGA 3 ND 0
HMGB 11 6 0
HON 6 4 2
HTA 16 ND 0
HTB 13 ND 0
HTR 14 ND 0
HUPA 1 ND 0
HUPB 2 ND 0
HXA 2 1 1
JMJ 4 ND 0
MBD 14 5 2
MFP 1 ND 0
NFA 5 2 2
NFB 1 ND 0
NFC 5 3 2
NFE 1 1 1
NFF 3 ND 0
PAFA 1 ND 0
PAFB 1 ND 0
PAFC 1 ND 0
PAFD 1 ND 0
PAFE 1 ND 0
PGE 2 1 0
PRMT 2 ND 0
RBL 2 ND 0
RDR 4 ND 0
RPDA 2 ND 0
RHEL 2 ND 0
SDG 35 15 7
SGA 3 ND 0
SGF 1 1 0
SMH 6 ND 0
SNT 2 ND 0
SRT 1 ND 1
SWDB 3 ND 0
SWDC 2 ND 0
TAFV 1 ND 0
VEF 3 1 1
36

Table 2.2.2. Genes selected from the Chromatin Database.
Gene Protein Tissue
RNAi
Family Ear Tassel Kernel Seedling Callus Leaf line
BRD101 Diverse bromodomain NSD NSD NSD NSD NSD NSD *
BRAT101 Bromodomain ++ ++ ++ *
CHB102 SWI/SNF (Swi3/Rsc8) ++ ++ + +
CHC101 SWI/SNF (Swp73/Rsc6) NSD NSD NSD NSD NSD NSD *
CHR101 SNF2 Super family ++ ++ ++
CHR106 SNF2 Superfamily ++
CHR110 SNF2 Superfamily ++ + +
CHR112 SNF2 Superfamily +
CHR113 SNF2 Superfamily ++ ++
CHR120 SNF2 Superfamily + ++
CHR125 SNF2 Superfamily + +
CHR126 SNF2 Superfamily + + + + *
DMT101 DNA Methyltransferase ++ + ++ ++ ++
DMT102 DNA Methyltransferase + + + + *
DMT103 DNA Methyltransferase ++ ++ ++ + ++ *
DMT104 DNA Methyltransferase + + + +
DMT105 DNA Methyltransferase ND ND ND ND ND ND
DMT106 DNA Methyltransferase ++ + + + ++
DMT107 DNA Methyltransferase ND ND ND ND ND ND
HXA102 Hist. acetyltransferase ++ ++ ++ + + *
HAG102 Hist. acetyltransferase + *
HAC 101 Hist. acetyltransferase ++ + ++ + +
HDA102 Histone deacetylase ++ ++ ++ ++ *
HDA110 Histone deacetylase + + + *
HDT101 Histone deacetylase + + ++ + + *
HDT103 Histone deacetylase +++ +++ +++ +++ +++ + *
HDT104 Histone deacetylase +++ +++ +++ +++
SRT101 Histone deacetylase ND ND ND ND ND ND *
HON101 Histone H1 +++ + ++ ++ + +
HON104 Histone H1 +
MBD101 Methyl binding domain +++ + + + ++ *
MBD109 Methyl binding domain +++ +++ + ++ ++ + *
NFA101 Nucleosome assembly +++ +++ +++ +++ +++ *
NFA104 Nucleosome assembly ++ + + + *
NFC102 NURF complex +++ +++ +++ +++ *
NFC104 NURF complex +
NFE101 Nucleosome positioning +++ ++ ++
SDG101 SET domain +++ + +
SDG102 SET domain ND ND ND ND ND ND *
SDG107 SET domain + + + + + *
SDG110 SET domain +++ +++ + + + *
SDG118 SET domain + *
SDG122 SET domain +
SDG129 SET domain +
VEF101 VEF family ++ +++ + + ++ + *
Levels of RNA expression is indicated by (+) weak, (++) medium and (+++) strong expression.
NSD no signal detected. ND no data
37

2.2.2. Microarray analysis of Chromatin Modifying Enzymes (CME)
A key feature of apomictic plants in Tripsacum is the precocious development of the
unreduced egg cell, which divides several days prior to fertilization to form a "proembryo"
consisting of 8 to 64 cells, and then remains quiescent until the endosperm develops following
fertilization (Grimanelli et al., 2003). We used dissected ovules of wild type maize and
apomicts maize-Tripsacum hybrids to compare on a genome scale gene expression at 0 DAP
(i.e. mature unfertilized ovules) between apomictic (containing a proembryo) and sexual
plants (contain an egg cell). The arrays were kindly provided and hybridized at the Syngenta
Biotechnology Institute (RTP, North Carolina) by Tong Zhu. We used custom-made 45K
maize Affymetrix chips, which contain a large representation of the maize transcriptome, and
include most expressed maize CMEs. The experiment consisted of 2 biological repetitions
mixed together for each sample and of 2 technical repetitions and allowed the comparison of
c.a. 45,000 probes between apomictic and sexual plants. Normalization was performed
according to standard Affymetrix protocols, and differentially expressed genes were identified
using ANOVA as described in Grimanelli et al. (2005). Using BLAST analyses, we then
extracted from the raw data set the probes corresponding to CME candidates.
Table 2.2.3. CME families and numbers of candidates identified based on microarrays
analyses.
PROTEIN FAMILY # of CMEs DIFF.EXPRESSION
BROMODOMAIN 19 0
SWI/NSF2 AND RSC COMPLEX 1 0
SNF2 SUPERFAMILY 4 1
DNA METHYLTRANSFERASE 3 0
HISTONE ACETYLTRANSFERASE 82 5
HISTONE DEACETYLASE (HD2 FAMILY) 27 2
HISTONEH1 15 0
METHYLBINDING PROTEIN 4 0
POLYCOMB GROUP 8 0
NURF COMPLEX 8 1
SET DOMAIN 69 5
Total 240 14
38

Histone Acetyltransferase
SNF2
ZM018126_AT
ZM051672_AT
ZM076171_AT
ZM043878_AT
ZM002094_AT
ZM064220_AT
ZM011705_AT
ZM011153_AT
ZM015023_AT
0 100 200 300 400 500 600 700 800 900
ZM079691_AT
NURF complex
ZMU16254_AT
ZM072252_AT
ZM023133_AT
ZM073333_AT
0 100 200 300 400 500 600 700 800
ZM064192_AT
SET domain
AY093418_AT
ZM037415_AT
AF545814_AT
ZM019715_AT
ZM052636_AT
ZM022324_AT
ZM022595_AT
ZM021320_AT
ZM016159_AT
Histone Deacetylase
ZM074706_X_AT
AF384033_AT
ZM016392_S_AT
ZM047050_AT
ZM069978_X_AT
ZM069978_AT
ZM076663_AT
AF384034_AT
ZM013552_AT
0 50 100 150 200 250 300 350
0 100 200 300 400 500 600 700 800 900
0 100 200 300 400 500
SEXUAL
APOMICTIC
Figure 2.2.2. Expression profiles within CMEs families that contain members showing differential
expression in microarrays analyses. x axis: expression values (fluorescence units). y axis: gene ID
(note that names are not shown for all probes analyzed). See also Table 2.2.4.
Collectively, on the 45k slides, differentially expressed genes represented only c.a. 1% of
the transcriptome, a very low proportion, but consistent with previously published work
(Grimanelli et al., 2005; Sharbel et al., 2009). Differential expression could be observed for a
limited number of CME probes (n=14) for which BLAST analyses revealed that they
39

elonged to 5 CME families only (Table 2.2.3 and 2.2.4): histone acetylases, histone
deacetylases (HDAs), SET domain proteins, NURF complex, and SNF2 superfamily.
Expression data are presented Figure 2.2.2. Interestingly, most differences corresponded to a
complete down regulation in the ovules of apomicts (e.g. #ZM064192) while the opposite
seldom occurred (e.g. HDA #ZM076663).
Table 2.2.4. Description of CMEs selected from microarrays experiments.
Maize ID
BLAST/ all organism DB
BLAST/ Plant DB
NSF2
ZM011153_AT
HAT
ZM001919_AT
Similar to A.thaliana; SWI2/SNF2-Like protein
Similar to O.sativa genomic DNA, chromosome 1
Similar to putative DNA dependent ATPase (Oryza sativa)
Similar to O.sativa putative taxadien-5-alpha-ol O-AT
ZM020825_S_AT Similar to ANASP Q8yva8 a arginine biosynthesis probable glutamate/ornithine acetyltransferase
ZM013490_S_AT Similar to Yeast Q03503 Saccharomyces cerevisiaeSimilar to At2g38130(GCN5-related N-acetyltransferase)
ZM071056_AT
ZM076568_AT
HDA
ZM076663_AT
Similar to O.sativa genomic DNA chromosome 7
Similar to O. sativa genomic DNA,chromosome 7
Similar to Zea mays HD2 type histone deacetylaseHDA106
Similar to N-acetyltransferase (Oryza sativa)
No description
No description
ZM033694_AT
NURF COMPLEX
ZM064192_AT
SET DOMAIN
ZM019715_S_AT
AF545813_AT
ZM047301_AT
No description
Similar to Zea Mays NFC102; WD40 proteins
Similar to A. thaliana unknow protein
Zea mays Set domain protein (sdg 113)
No Description
Similar to HDAC_Maize probable histone deacetylase
Putative DNA polymerase (O. sativa)
Similar to SET domain-containing protein-like (O. sativa)
Similar to SET domain 113
Similar to SET domain-containing protein-like (O. sativa)
ZM023509_AT Similar to A.thaliana BAC F16A14 from Chrom 1 Similar to SET domain-containing protein-like (O. sativa)
ZM080896_AT
Similar to mez2
Similar to MAIZE protein EZ2
2.2.3. Preliminary analysis of selected genes
Preliminary validation by RT-PCR was performed using ovule tissues collected from
sexual maize and apomictic maize-Tripsacum, precisely timed at three critical developmental
stages: megasporogenesis, mature embryo sac and 3 days after pollination (DAP). A detailed
description of the most interesting candidates is provided in Chapter 3. The results presented
here focus on the comparison of the global transcription behavior of the 56 selected CMEs
between apomictic and sexual developments, and between developmental stages within each
reproductive pathway.
In sexual plants, overall transcriptional activity (steady-state RNA levels) for all 56 genes
appeared surprisingly stable across all three developmental stages, with only a few genes
40

changing expression profiles as development proceeded (in this case, silenced or showing
reduced expression; Figure 2.3.1).
The situation looked very different in apomictic plants, particularly during early
development. We detected 9 CMEs that were totally down-regulated during apomeiosis,
included in a total of 20 CMEs with some reduction in expression during apomeiosis. It is
remarkable that only one locus showed up-regulated expression at this stage. Surprisingly, this
trend towards globally reduced CME activity in apomicts vanished during later stages. In
mature embryo sacs, 9 of the 20 down-regulated candidates recovered identical expression in
apomictic and sexual plants (Table 2.2.5). At 3 DAP both reproductive modes displayed
similar transcriptional patterns within ovules (Figure 2.3.1), with only few exceptions.
Interestingly, only 7 CMEs remained down-regulated during all three developmental
stages, and only one, HXA102, remained up-regulated during all three stages of apomictic
development. The analysis of over-expressed chromatin regulators could lead to interesting
insights into apomictic reproduction, since apomixis has been largely conceived as a gain of
function phenotype.
100%
90%
80%
70%
60%
50%
40%
30%
20%
10%
0%
3
3
3
5
42
9
7
2
11
27
2
2
6
44
Sexual 1 Apo 2 Sexual 3 Apo 4 Sexual 5 Apo6
7
1
9
38
4
1
6
9
36
7
5
1
7
36
Série5
Série4
Série3
Série2
Série1
MEIO
MES
3DAP
Figure 2.3.1. Global transcription patterns of selected CMEs in sexual and apomictic (Apo) ovules
collected at three developmental stages: megasporogenesis (MEIO), mature embryo sac (MES), and 3
DAP. Transcription levels were determined by RT-PCR and classified according to the scale shown on
right. Pictures illustrate reproductive stages and were obtained from cleared ovules.
41

Table 2.2.5. Chromatin Modifying Enzymes Deregulated during Apomictic Development.
EXPRESSION
STAGES
PROFILE APOMEIOSIS MES 3DAP
CHB102 CHB102 CHR106
CHR106 CHR106 CHR120
CHR20 CHR120 DMT102
CHR125 DMT102 DMT103
DMT102 DMT103 DMT105
DMT103 DMT105 BT066941
DMT105 HON101 HDT104
DMT107 MBD109 HON101
Down
HAG102 NFC104 NFA104*
HAG101 SDG118* NFC104
BT066941 AW288923 DR959742*
AY110756
EE168341*
HDT104
EC897242
HON101
MBD109
NFC104
SDG129
AW288923
EC897242
VEF101
HXA102 HXA102 BRD101
AC157319 CHR126
AY104727 HXA102
HDT104 HAG102
Over
SDG107 SDG102
SDG107
SDG110
AW288923
VEF101
(*) CME not registered in apomeiosis
2.3. DISCUSSION
A commonly accepted hypothesis for the molecular bases of apomixis advocates that it
results from the deregulation of genes controlling sexual reproduction (reviewed Koltunow
and Grosniklaus, 2003). Surprisingly then, global transcription profiling of sexual and
apomictic ecotypes in different systems have not in past experiments revealed many
significant differences (Grimanelli et al., 2005; Sharbel et al., 2009; Yamada-Akiyama et al.,
2009).
Results presented here point out a similar behavior; on the complete 45k arrays, the
differences between apomictic and sexual ovules represented c.a. 1% of the transcriptome,
below our false discovery rate. Only 6% of the CMEs analyzed using microarrays (before
validation) showed differential expression. This is much higher than the overall RNA
population, but remains a low proportion. These results might suggest that only a few key
42

genes, acting on regulatory developmental checkpoints, are altered in apomicts. Alternatively,
this might suggest that the key differences might occur at a post-transcriptional level. Another
explanation is that the differentiation between apomixis and sexuality occurs at a much earlier
stage of development, and does not require significant transcriptional changes at later stages.
Global differences became more evident, in fact, when analyzing more closely for selected
CMEs. The most striking differences between apomictic and sexual plants were found at very
early stages (meiosis vs. apomeiosis). This either suggests a deregulation in apomeiosis or
alternatively indicates that many of these CMEs are specific or highly expressed at meiosis
and thus not found in apomictic plants. Remarkably, more that half of the CMEs differentially
expressed during sporogenesis recovered identical expression during gametogenesis, and
virtually all of them did so during early embryogenesis.
By contrast, only marginal differences were detected between developmental stages during
sexual reproduction. This might suggest, but requires additional analyses, either that the bulk
of CME transcripts is deposited early in the reproductive cells, and carried over throughout
the gametophytic phase, or, alternatively, that most changes in the recruitment of specific
genes occurs at the post-transcriptional level. In any case, these data suggest certainly that the
key differences between apomixis and sexuality are established early during reproductive
development, likely during or before sporogenesis, with consequences throughout
reproduction.
Most CMEs with differential patterns showed a down-regulation phenotype in apomictic
plants when compared to sexual ones. Only a reduced group (6), sharing similar functions
(see chapter 4), remained downregulated during all developmental stages. Only one
(HXA102) remained up-regulated during the whole apomictic development. The follow step,
thus, was to assess the functional link between changes in the activity of these CMEs and
apomixis. For this, we are using two different approaches. For the genes that are up-regulated
in apomictic plants, we are using transgenic RNAi maize line to induce dominant loss-offunction
alleles into an apomictic background, using a crossing strategy previously described
(Leblanc et al., 1996). Alternatively, as will be presented in the next chapter, we are using
loss-of-function maize mutants to analyze the effect of down-regulating candidates in a sexual
context.
43

CHAPTER 3.
Inactivation of a Reproductive DNA Methylation
Pathway in Maize Results in Apomixis-Like Phenotypes
(manuscript currently under revision for publication)
44

Inactivation of a Reproductive DNA Methylation Pathway in
Maize Results in Apomixis-Like Phenotypes.
Marcelina Garcia-Aguilar, Olivier Leblanc and Daniel Grimanelli*
Institut de Recherche pour le Développement, UMR 5096 (IRD/CNRS/Université de
Perpignan), Montpellier, France
Correspondance to Daniel Grimanelli, Institut de Recherche pour le Développement,
UMR5096, 911 Avenue Agropolis, 34394 Montpellier France. T: +33 (0)4 67 41 63 76.
daniel.grimanelli@ird.fr
Running tittle: Regulation of DNA Methylation and Apomixis
Estimated lenght: 12 pages
The author responsible for distribution of materials integral to the findings presented in this
article in accordance with the policy described in the Instructions for Authors is: Daniel
Grimanelli, daniel.grimanelli@ird.fr
45

SUMMARY
Apomictic plants reproduce asexually through seeds by avoiding both meiosis and
fertilization. Although apomixis is genetically controlled, its core genetic component(s) has
not been determined yet. Using profiling experiments comparing sexual development in
maize (Zea mays) to apomixis in maize-Tripsacum hybrids, we identified six loci that are
specifically downregulated in ovules of apomictic plants. Four of them share strong homology
with members of the RNA directed DNA methylation (RdDM) pathway, which in Arabidopsis
is involved in silencing via DNA methylation. Analyzing loss-of-function alleles for two
DNA methyltransferase genes, dmt102 and dmt103, respectively homologous to Arabidopsis
CHROMOMETHYLASE3 (CMT3) and DOMAINS REARRANGED
METHYLTRANSFERASE2 (DRM2), revealed phenotypes highly reminiscent of apomictic
developments, including the production of unreduced gametes and formation of multiple
embryo sacs in the ovule. Loss of DMT102 activity in ovules resulted in the establishment of
a transcriptionally competent chromatin state in the archesporial tissue and in the egg cell that
mimics the chromatin state found in apomicts. Interestingly, dmt102 and dmt103 expression in
the ovule is found in a restricted domain in and around the germ cells, indicating that a DNA
methylation pathway active during reproduction is essential for gametophyte development in
maize, and likely plays a critical role in the differentiation between apomictic and sexual
reproduction.
46

3.1. INTRODUCTION
Sexual reproduction in angiosperms takes place within a multicellular ovule in which the
formation of the female gametes entails two consecutive steps: megasporogenesis (spore
formation) and megagametogenesis (gamete formation). Megasporogenesis initiates with the
differentiation of the megaspore mother cell (MMC), which undergoes meiosis, producing
four haploid spores. Three of them usually degenerate, leaving a single functional megaspore.
In the Polygonum type of megagametogenesis, the most common in angiosperms, the
megaspore undergoes three rounds of mitotic divisions to form the embryo sac (ES)
containing the female gametes (the egg cell and the central cell), two synergids at the
micropylar pole and three antipodal cells at the chalazal pole (Reiser and Fischer, 1993). In
male reproductive organs (the anthers), all four meiotic products form male gametophytes
(pollen grains), which consist of two reproductive sperm cells embedded in the vegetative
cell. The fertilization of the egg cell and the central cell gives rise to the embryo and the
endosperm respectively.
Apomixis refers to a diverse group of reproductive behaviors, resulting in asexual
reproduction through seeds (Nogler 1984). Apomictic plants bypass both meiotic reduction (a
process called apomeiosis) and egg-cell fertilization (via parthenogenesis), thus producing
offspring that are exact genetic replicas of the mother plant. In the diplosporous type of
apomixis, which occurs for example in Tripsacum (Leblanc et al., 1995), a wild relative of
maize, or Boechera species (Naumova et al., 2001), that are related to Arabidopsis, the MMC
differentiates but fails to complete meiosis and finally produces an unreduced functional
megaspore. Megasporogenesis then proceeds similarly to sexual plants, but the embryo
develops from the unreduced egg cell without fertilization, i.e. by parthenogenesis. In the
aposporous type of apomixis, megasporogenesis is omitted and one or more ESs can develop
directly from somatic nucellar cells (aposporous initials), adjacent to the surviving
megaspore. The embryo then similarly develops parthenogenetically. Development of the
endosperm of apomictic plants is either pseudogamous, i.e. it arises from the fertilization of
the central cell by one male sperm, or autonomous when it occurs without fertilization of the
central cell.
47

In many apomictic species, meiotic abnormalities are not restricted to the female
reproductive cells. Unreduced pollen grain formation has been reported for example in
Boechera (Schranz et al., 2006) and Tripsacum (Grimanelli et al., 2003), suggesting that male
and female reproductive paths are affected in similar ways. Also, in most if not all
documented cases, apomixis and sexuality are not mutually exclusive since they usually
coexist in the same ecotypes with varying levels of relative expressivity (Nogler, 1984). Most
diplosporous apomicts produce apomeiotic (unreduced) and meiotic (reduced) spores and
female gametes. Similarly, in the case of aposporous apomixis, the ovule usually contains
both meiotically derived and apomeiotically derived ESs. Interestingly, while the formation of
multiple ESs is a hallmark of aposporous apomixis, some diplosporous plants also develop
extra embryo sac-like structures (e.g., Paspalum minus; Bonilla and Quarin, 1997). Therefore,
the frontier between both types of apomixis and between apomixis and sexuality, appears
relatively blurry, fueling the perception that apomixis might be a deregulated form of sexual
reproduction rather than a new function. Indeed, previous studies showed that apomixis and
sexuality share key regulatory mechanisms (Tucker et al., 2003). Thus, it has been proposed
that apomixis could result from a temporal or spatial deregulation of the transcriptional
programs controlling sexual reproduction (Koltunow and Grossniklaus 2003; Grimanelli et
al., 2003; Bicknell and Koltunow, 2004; Bradley et al., 2007). The model postulates that
sexual reproduction involves several transitions (from somatic cells to reproductive cells,
from the sporogenesis to the gametogenesis, from gametogenesis to embryogenesis) whose
orderly progression is altered in apomicts, resulting in ectopic or heterochronic expression of
the core developmental program.
Recent studies have further demonstrated the crucial role of chromatin-based regulation in
vegetative developmental transitions (Poethig, 2003; Bezhani et al., 2007). Similarly, several
lines of evidence suggest that transitions during reproduction and early seed development are
epigenetically controlled through dynamic changes in chromatin state (Huanca-Mamani et al.,
2005; Xiao et al., 2006; Baroux et al., 2007; Curtis and Grossniklaus, 2008). Whether
alterations in the epigenetic regulation orchestrating these transitions are involved in the
differentiation between apomictic and sexual reproduction is currently unknown.
In this study, we compared the expression patterns of diverse chromatin modifying
enzymes (CMEs) during reproduction in sexual and apomictic plants to identify possible
chromatin-level regulators of apomixis. Two key mechanisms affecting the transcriptional
competence of chromatin are covalent modifications of histone tail residues and DNA
48

(Cytosine) methylation (Vaillant and Paskowski, 2007). DNA methylation in plants takes
place on CG, CNG and CNN. In Arabidopsis, where the process is best understood, at least
three classes of DNA methyltransferases (DMTs) intervene in methylation pathways. DRM2
(DOMAINS REARRANGED METHYL TRANSFERASE2) is the main de novo DMT, involved
in all sequence contexts (Cao and Jacobsen, 2002). DRM2 also plays a role, together with
CMT3 (CHROMOMETHYLTRANSFERASE3), in the maintenance of methylation at non-CG
sites (Cao et al., 2003) Another methyltransferase, MET1 (METHYLTRANSFERASE1), is
involved in maintaining DNA methylation at CG dinucleotides. Finally, DDM1 (DECREASE
IN DNA METHYLATION1), a SWI2/SNF2-like chromatin-remodeling factor, also participates
in maintenance of methylation at both CG and non-GC sites (Jeddeloh et al., 1999). Along
with DMTs, small interfering RNAs (siRNAs), histone modifying enzymes and RNAi
proteins are involved in mediating both DNA and histone methylation via the so-called RNAdirected
DNA methylation (RdDM) pathway (Huettel et al., 2007). The silencing effect of the
RdDM pathway is reinforced by the establishment of repressive histone marks, in particular
methylation of lysine 9 on histone H3 (H3K9me), by the activity of KYP (KRYPTONITE), the
main H3K9 methyltransferase in Arabidopsis (Jackson et al., 2002).
Here, we show that apomictic and sexual development differ for the expression of a few
CMEs that are specifically expressed in reproductive cells of maize and downregulated in
ovules of apomictic ecotypes. We further demonstrate that dmt102 and dmt103, respectively
homologous to CMT3 and DRM2 in Arabidopsis, play key roles in ovule development in
maize. Loss-of-function mutants result in phenotypes that are strikingly reminiscent of
apomictic development, suggesting that, in addition to a crucial role in gametophyte
development, DNA methylation in the maize ovule might regulate transcriptional expression
of genes involved in the differentiation between apomixis and sexuality.
3.2 RESULTS
plants
3.2.1. Identification of CMEs differentially expressed in sexual and apomictic
To address the possible role of chromatin structure in the regulation of apomixis, we first
performed a comparative analysis of transcription activity of CMEs during sexual and
apomictic reproduction. We used the B73 maize inbreed as a sexual control. The apomictic
plant used in this analysis, referred to as 38C, is an apomictic hybrid obtained from an
original F1 plant between maize and its apomictic relative, Tripsacum dactyloides 65-1234,
49

and expresses apomixis with high penetrance (>99%). 38C plants contain one diploid set of
chromosomes from maize (2n=2x=20) and one haploid set of chromosomes from Tripsacum
(1n=1x=18). They reproduce via diplospous apomixis; an unreduced ES is derived from a
MMC and premature divisions of the unreduced egg cell result in a precocious,
parthenogenetic pro-embryo, formed before fertilization. This apomictic model plant has been
described in previous publications (Leblanc et al., 1996; Grimanelli et al., 2003; Grimanelli et
al., 2005; Leblanc et al., 2009).
We re-analyzed a set of microarray data comparing non-pollinated mature ovules of maize
and 38C to identify maize CMEs expressed during sexual and apomictic development
(Grimanelli et al., 2005). In addition, we included in the analysis a set of CMEs reported in
The Chromatin Database (www.chromdb.org) as expressed in maize ears. Both data sets
contained collectively 384 loci annotated in the B73 genome as CMEs (Supplemental Table
1). Based on data indicative of expression within reproductive tissues (see Methods), we
selected 56 loci, belonging to 15 different protein families (Supplemental Table 2). We then
used RT-PCR to monitor, at precise stages of female reproductive development, the
expression of the corresponding maize alleles in sexual maize and after their introduction in
the 38C apomictic background. These stages included: dissected ovules during sporogenesis
(including thus meiosis and apomeiosis), mature ESs before fertilization (containing either an
egg cell or a parthenogenetic pro-embryo in sexual and apomictic plants, respectively) and
seeds collected 3 days after pollination (DAP) (Supplemental Figure 1). Differences in
expression profiles therefore indicated that the introduction of a given maize allele in the
apomictic plants resulted in altered expression at the same developmental stages, and served
as markers to screen for loci subjected to differential regulations potentially linked, directly or
indirectly, to divergent reproductive behaviors.
As previously described (Grimanelli et al., 2005), global expression profiles between
apomictic and sexual plants differed only marginally and, accordingly, we identified only six
loci with clear qualitative differential expression between the two reproductive modes
(Supplemental Figure 2). For all six loci, the PCR primer pairs amplified both the maize and
Tripsacum alleles on genomic DNA, suggesting that the differences in expression affected
both the maize and Tripsacum alleles. Four of the six loci showed complete downregulation at
all stages while the two remaining ones exhibited heterochronic expression (Figure 1A).
BLAST analyzes against the Arabidopsis and maize databases indicated probable functions
for all six proteins (Table 1). dmt102 and dmt105 encode DMT enzymes, both homologous to
50

Arabidopsis CMT3 (Supplemental Figure 3). DMT102 is required for cytosine methylation at
CNG sites and it is likely involved in a maintenance function (Papa et al 2001; Makarevich et
al., 2007). dmt103 is a close homologue of DRM1 and 2 (Supplemental Figure 3); it encodes a
putative DMT that shows rearranged catalytic domains conserved in others eukaryotic
proteins, but its function remains uncharacterized in maize. chr106 is homologous to
Arabidopsis DDM1. All four genes were broadly expressed during sexual development in
maize, but totally absent during apomictic reproduction. The set of deregulated genes also
included hdt104 a member of the plant-specific histone deacetylase family (homologous to
AtHD2A), and hon101, a histone H1 linker protein gene. Furthermore, DMT102, DMT103,
DMT105 and CHR106 are clearly homologous to well-characterized enzymes involved in
DNA methylation in Arabidopsis. Interestingly, dmt101, the closest homologue of the
Arabidopsis MET1 gene, showed similar expression patterns during sexual and apomictic
developments (Supplemental Figure 2).
Previous studies have linked changes in DNA methylation with changes in polyploidy
levels (Wang et al., 2004). It is also known that interspecific hybridizations can induce
epigenetic variations (Madlung et al., 2002; Chen et al., 2008). Interestingly, both
hybridization and polyploidy have been proposed as direct causes for the induction of
apomixis, through unknown routes (Carman, 1997; Koltunow and Grossniklaus, 2003; Paun
et al., 2006). Apomictic 38C was derived by interspecific hybridization between maize and
Tripsacum and it carries a polyploid set of chromosomes (Leblanc et al., 1996). To evaluate
the possibility that interspecific hybridization or polyploidy, or both, might have induced the
expression patterns detected in 38C, we compared the expression of dmt102, dmt103 and
chr106 in mature embryo sacs of Tripsacum dactyloides 65-1234, the natural apomictic plant
used to produce the 38C hybrid, with that observed in diploid and tetraploid sexual maize
(B73 inbreed and N108A stocks, respectively) and in the maize-Tripsacum 38C hybrid
(Figure 1B). We found that expression of chr106 was identical in diploid and tetraploid sexual
maize lines. chr106 was also similarly downregulated in 65-1234 and 38C apomicts for both
the maize and Tripsacum alleles. Thus, the deregulation of chr106 in apomicts is independent
of either polyploidy or interspecific hybridization. By contrast, while dmt102 and dmt103
transcripts corresponding to maize and Tripsacum alleles were neither detected in Tripsacum
nor in 38C apomictic contexts, both genes were strongly expressed in diploid maize but
downregulated in the autotetraploid. Their expression is therefore ploidy-dependent, but is
independent of interspecific hybridization effects.
51

A
chr106
B73 sexual
SPO MES 3DAP
38C apomict
SPO MES 3DAP
dmt102
dmt103
dmt105
htd104
hon101
actin1
B
gDNA
cDNA
38C Td
H 2
O B73
38C Td
4x H 2
O B73
dmt102
dmt103
chr106
actin1
C
A.thaliana B.holboellii gDNA
SPO MES SPO MES At Bh
DRM1
DRM2
CMT3
DDM1
ACTIN11
Figure 1. RT-PCR of CMEs deregulated during apomictic reproduction.
(A) Expression of CMEs in ovules during sporogenesis (SPO), in ovules containing mature embryo
sacs (MES) and in 3 DAP seeds in maize (B73) and apomictic 38C plants.
(B) Effect of ploidy and inter-specific hybridization on the expression of dmt102, dmt103 and chr106
in apomictic (38C and Td) and sexual (B73 and 4x) ovules containing mature embryo sacs. gDNA:
genomic DNA. Td: Tripsacum dactyloides ecotype 65-1234. 4x: autotetraploid of B73.
(C) Expression of DRM1, DRM2, CMT3 and DDM1 in Arabidopsis thaliana and its apomictic relative
Boechera holboellii using ovules containing megaspore mother cells (SPO) and mature embryos sacs
(MES). gDNA: genomic DNA. At: Arabidopsis thaliana. Bh: Boechera holboellii.
52

3.2.2. Apomixis correlates with the downregulation of CMT3 and DRM1/2
homologues in both Tripsacum and Boechera
Apomixis in plants can occur by different developmental routes and it is an open question
whether the different flavors of apomictic development rely on similar or unrelated regulatory
pathways. To address this, we analyzed the expression of DRM1 and DRM2, CMT3, and
DDM1 in growing ovules of Arabidopsis thaliana and of Boechera holboellii, a diplosporous
apomictic relative. We performed RT-PCR on ovules dissected at the MMC and mature ES
stages using primer pairs that amplified genomic fragments in both species (Figure 1C). We
found that both DRM genes were deregulated, though in a different manner, in apomictic
Boechera when compared to sexual Arabidopsis. DRM2 was downregulated during
sporogenesis in apomeiotic ovules of Boechera but not in mature embryo sacs. DRM1 on the
other hand showed clear expression at both stages in sexual plants but no expression in
apomictic ovules. Finally, CMT3 and DDM1 had similar expression patterns in both species,
suggesting differential regulation in maize and Arabidopsis. However, we cannot discard a
possible downregulation specifically within Boechera gametophytes that would not affect
expression in the surrounding somatic tissues, and more detailed studies are required to assess
their role in apomixis in Boechera. These results nonetheless show that at least DRM1 and
DMR2, or their homologues, are deregulated in two unrelated apomictic species.
3.2.3. dmt102 and dmt103 are expressed in the reproductive cells in the maize ovule
RT-PCR (Figure 1; Supplemental Figure 2) showed that in sexual maize dmt102 and
dmt103 are not expressed in somatic tissue such as leaves, and are mainly expressed during
early reproductive stages, including in male gametophytes (data not shown). To further
characterize their expression patterns in reproductive tissues, we performed in situ RNA
hybridization in ovules of wild-type maize plants (Figure 2). Both were expressed in a
restricted portion of the nucellus during megasporogenesis. dmt102 expression domain
encompassed the reproductive cell and a few layers of nucellar cells around it, while dmt103
was detected only in the epidermal layer and integuments of the growing ovule. dmt102
conserved the same spatial expression during early gametogenesis, while dmt103 expression
expanded to include the ES. At the end of gametogenesis, the expression of both genes
became confined to the ES area with stronger signals in the chalazal region. In Arabidopsis,
the detailed expression pattern of CMT3 or DRM1/2 during reproductive development have
not been reported. However, consistent with their maintenance function, these genes are
53

oadly expressed in somatic tissues (e.g., leaves, seedlings) according to the Arabidopsis
expression databases (http://bbc.botany.utoronto.ca/efp/cgi-bin/efpWeb.cgi), by contrast to
dmt102 and dmt103, as shown here. This suggests a specialization in the maize genome of at
least a subset of DNA methylation enzymes to reproductive functions.
3.2.4. Downregulation of dmt102 and dmt103 in sexual maize promotes the
formation of unreduced gametes
To further define the biological role of this DNA methylation pathway in reproductive
development, we wanted to analyze the phenotypes of loss-of-function lines for dmt102 and
dmt103 in sexual maize. For dmt102, we used a previously characterized mutant line. It
carries a Mutator element inserted within the DMT domain and produces an aberrant protein
(Papa et al., 2001). The mutant line has no apparent morphological or seed phenotype and the
dmt102::Mu alelle was normally transmitted via both male and female gametes in reciprocal
crosses to wild type (B73 line) plants. For dmt103, we analyzed two dmt103-specific RNAi
lines, hereafter referred to herein as D103-2 and D103-4, produced by the Maize Chromatin
Consortium (McGinnis et al., 2005) and one EMS allele (dmt103-6) recovered from the maize
TILLING population that contains two non-synonymous substitutions in the coding region of
54

the gene (R49I and R272Q). None of these dmt103 loss-of-function lines showed
morphological phenotypes during vegetative development. T3 progenies of both RNAi lines
exhibited severe effects on seed morphology. When used as females, all seeds produced by
heterozygous D103-4 T3 lines were at least partially defective, with miniature and aborted
kernels, the latter representing 53% of the kernels (Supplemental Figure 4). Crossing
heterozygous D103-2 with wild type pollen resulted in normal (21%), miniature (37%) and
aborted (42%, n=169) kernels (Supplemental Figure 4). This suggests at least a maternal
sporophytic effect of Dmt103 inactivation when the RNAi-inducing sequence is driven by the
35S promoter. However, additional gametophytic effects could not be excluded.
The three dmt102 and the dmt103 mutant lines produced abundant pollen grains following
normal anther dehiscence. However, cytological analyses of male gametophytes in
dmt102::Mu mutants and in D103 lines revealed a high proportion of abnormally large pollen
grains, i.e. ~40% and ~25% respectively (Figure 3A-B-C). To verify whether large pollen size
correlated with unreduced gametes formation, we quantified DNA contents in bulks of mature
pollen grains by flow cytometry. Quantification of relative fluorescence intensity confirmed
the production of reduced, but also unreduced and aneuploid pollen grains at a frequency that
matched the proportion of large gametophytes (Figure 3C-D). This indicates that
downregulation of both dmt102 and dmt103 resulted in the production of unreduced male
gametes.
Whether unreduced female gametes were also produced was technically more difficult to
assess. All the viable seeds produced in both mutants contained diploid embryos when
pollinated by wild type pollen (data not shown), showing that the viable embryos originated
from reduced female gametes. dmt102::Mu homozygous lines produced nearly full seed sets,
indicating that most, if not all female gametes were meiotically derived. In contrast, the high
proportion of abortive seeds in D103 lines might reflect endosperm defects resulting from
altered genome parental dosage due to the occurrence of unreduced or aneuploid female
gametes, or poor viability of unreduced or aneuploid gametes (Birchler, 1993). Interestingly,
cleared ovules from both D103 lines at the mature gametophyte stage exhibited surprisingly
large female gametes (Figure 3E). Whether they reflect non-reduction is unclear and requires
further examination.
55

A
B73
1C
B
dmt102::Mu
B73
1C
200
D103-4
2C
100
dmt102::Mu
0
dmt102::Mu
Dmt103-4
WT
C
B73
D103-4
EC
PN
EC
PN
Figure 3. dmt102 and dmt103 mutants produce unreduced male gametes.
(A) From left to right: morphology, and 1C DNA contents in wild type pollen grains.
(B) From left to right: morphology, DNA contents in mutant pollen grains (the 2C peak represents
unreduced and aneuploid pollen grains), and size distribution (gray bars: small grains; white bars:
large grains).
(C) Whole mount ovule clearing of a D103-4 mature ovule and wild type maize; an abnormally
large female gamete is visible at the micropylar pole; EC: egg cell; PN: polar nuclei.
DNA content in pollen grain were determined by flow cytometry. The peak corresponding to
haploid DNA content (1C) is determined in the B73 sample. The extra peak detected in the mutant
is centered on 2C. x axis: relative fluoresecence intensity. y axis: number of nuclei. Bars: 100!m.
56

3.2.5. Downregulation of dmt102 and dmt103 in sexual maize induces the formation
of multiple embryo sacs in the ovule
To better characterize the effect of dmt102 and dmt103 loss of function on ES
development, we cleared ovules of the mutant lines at different developmental stages.
Interestingly, we observed related phenotypes in ESs of both mutants. We found that ESs of
dmt102::Mu plants exhibited normal development until cellularization. The mature ES
conserved the Polygonum type organization occurring in maize. In particular, the three
antipodal cells proliferated before pollination (Figure 4A), thus producing an ephemeral
structure of approximately 50-100 cells, which shares protoplasm content because of
incomplete cell wall formation (Diboll and Larson, 1966). ESs in dmt102::Mu/dmt102::Mu
ovules followed the wild type behavior with antipodal cells proliferation. However, at late
stages (mature ES), we observed the development at the chalazal pole of large structures (in
c.a. 25% of ES; Figure 4B), presumably originating from abnormal growth of antipodal cells
and strongly resembling small uni-nucleated ESs, as typically seen before the first division of
the functional megaspore. No further development was observed since they remained arrested
as large chalazal cells until the multicellular antipodal structure degenerated. This suggests
that DMT102 might be involved in the maintenance of antipodal cell fate during late
gametogenesis.
In D103 lines, a similar but much more severe phenotype was found and, despite normal
development of ovule primordia at early stages, we observed remarkable sporophytic and
gametophytic defects by the time megagametogenesis took place (Figure 4; Table 2 and 3).
First, we noticed an incomplete ovule rotation that caused drastic changes in ovule polarity
and in ES position, which frequently extruded from the nucellus or collapsed on the ovary
wall (Supplemental Figure 4). Furthermore, the total number of gametophytic nuclei was
unpredictable. These were clustered in central position within ESs, eventually acquiring the
typical shape and large nucleoli associated with polar nuclei (Figure 4C). In particular,
individual ES with up to 8 unfused polar nuclei were observed. Proliferating nuclei of unclear
identity were also observed in atypical positions within central cells (Figure 4E), similar to
what has been described in the ig1 (indeterminate gametophyte1) mutant of maize (Evans,
2007). Extra egg cells, however, were not seen in individual ESs. In addition, loss of polarity
of the ES was observed in morphologically normal ovules, resulting in an inverted orientation
of the cells (Figure 4D). Strikingly, D103-2 individuals also showed frequent production of
57

extra ESs, usually positioned at the chalazal pole (Figure 4E-K, Table 2). The number,
position and size of these extra ESs were variable, but the presence of differentiated egg and
polar nuclei was a consistent feature of these structures. Contrary to the ES positioned at the
micropyle, the apical-basal polarity of extra ESs usually was inverted (Figure 4G). The
multiple ES phenotype was clear in D103-2 but rare for D103-4 plants (Table 2). However,
observation of cleared sections of dmt103-6 ovules also revealed abnormal gametophytic
development, including multiple ESs formation (Table 2). Collectively, this suggests that
DMT103 is essential for several aspects of gametophytic development in maize, with a
specific role in limiting the number of ES in the ovule.
An important question regarding the extra ESs is whether they originated from haploid
gametophytic cells or from diploid somatic (nucellar) cells in the ovule. Our observations
suggest that both developments are possible. At maturity, it was hard to differentiate ESs
originating from either cell types, because the supernumerary gametophytes occupied a large
nucellar volume adjacent to the micropylar ES (Figure 4F-G) and collapsed antipodal and
nucellar cells were difficult to characterize. However, a large proportion of the mature ESs
seen in the most severely affected dmt103 RNAi line (D103-2) was deprived of antipodal
cells. Instead, and very similar to the dmt102::Mu mutant, large undifferentiated cells were
localized at the chalazal pole, in the position normally occupied by the antipodal cells (Figure
4I-J). This suggests that the extra ESs were derived from the cellularized gametophyte, either
before or after the definition of the antipodal cells. However, we also observed the
development of large undifferentiated cells in the nucellus, similar to uninucleate ESs in wild
type plants. They were separated from the ES proper by one or a small number of cell layers
and thus clearly distinct from the antipodal cells (Figure 4I-J). At even earlier stages, we also
noticed the formation of multiple uninucleate ESs, arising from distinct cells (Figure 4K).
This indicates that extra ESs can be produced from both the gametophytic cells and the
nucellar cells at the chalazal pole of the ovule.
To further investigate the function of DMT102 and DMT103, we generated Dmt102/
dmt102::Mu individuals carrying the RNAi transgene from D103-4. While Dmt103
inactivation in the D103-4 line showed lower frequency of multiple ESs compared to that of
D103-2, these plants exhibited an increased severity of the dmt103 RNAi phenotype as the
frequency of multiple ESs increased significantly (from 15% to 40%; Tables 2 and 3). Thus,
combining loss of function for both genes significantly enhanced the phenotypic effect
observed in D103-4. We next examined segregating F2 families derived from two F1
59

individuals (F2#3 and F2#12). We found that 78% of the plants produced unreduced male
gametophytes, but without additive effects since the proportion of unreduced gametophytes
remained similar in the double or simple mutants. Multiple ES development was found in
28% of the F2s and co-segregated with the RNAi transgene only (Figure 5). These data
indicate that DMT102 and DMT103 likely act on a common process in the ovule, consistent
with their mRNA colocalisation in this tissue, but that the formation of fully developed
supernumerary ESs seems to be more specific of dmt103 loss of function.
3.2.6. Patterns of chromatin modification in dmt102 mutant lines mimic the effect
of apomixis during sporogenesis and gametogenesis
Since key DMT proteins were deregulated in apomictic 38C, we were interested in
comparing DNA methylation patterns in the reproductive cells of sexual, mutant and
apomictic plants. The restricted domains of expression of both Dmt102 and Dmt103 within
60

A
H3K9ac
DAPI
B
H3K9ac
DAPI
An
B73
B73
An
dmt102::Mu
dmt102::Mu
D
CC
H3K9me2
Syn
38C
EC
C
CC
H3K9me2
EC
DAPI
Relative Intensity, EC/CC
2-
1-
0-
DAPI
B73 B73
dmt102::Mu
H3K9me2
dmt102::Mu
Figure 6. Patterns of H3K9 acetylation and dimethylation in dmt102::Mu mutant lines mimic the
effect of apomixis during sporogenesis and gametogenesis.
(A) H3K9 acetylation in the ovule during sporogenesis. The dash lines indicate the nucellar domain
enclosed within the inner integuments.
(B) H3K9 acetylation in mature ovules. An: antipodal cells nuclei.
(C, D) H3K9 dimethylation in ESs of wild type (C) and mutant (D) plants; EC: egg cell nucleus;
CC: central cell nuclei; Syn: synergid nucleus; Quantification of signals intensity measures the
ration of egg cell to central signal per unit of DNA, to take into account the dihaploid nature of the
central cell. The same measure performed for DAPI signals produced the expected 1:1 ratio. The
quantification shows a significant reduction of relative H3K9me2 in the egg cell in the mutant
plant.
the ovule, however, rendered chromatin-IP or related experiments technically challenging.
Alternatively, we used immunolocalization experiments to determined chromatin states during
sporo- and gametogenesis by checking the global distribution of two antagonist marks,
61

dimethylation of H3K9 (a repressive mark) and acetylation of H3K9 (a permissive mark).
Previous studies in Arabidopsis have demonstrated the close interplay of DNA and histone K9
covalent modifications (Jackson et al. 2002; Soppe et al., 2002; Tariq et al., 2003; Lindroth et
al., 2004; Mathieu et al., 2005; Johnson et al., 2007). For these experiments, we used the
stable dmt102::Mu mutant line rather than the D103 RNAi lines, reasoning that the relative
instability of RNAi induction might result in excessive variability when comparing individual
ovules.
H3K9 acetylation (H3K9ac) patterns differed markedly between wild type and
dmt102::Mu maize ovules. During sporogenesis in wild type ovules, the reproductive cells, as
well as surrounding somatic cells, were conspicuously deprived of H3K9ac signals (Figure
6A, n>25). This pattern followed closely the spatial localization of Dmt102 mRNA, as
observed by in situ hybridization analysis at the same developmental stage (Figure 2). In
contrast, H3K9ac signals in dmt102::Mu line, were clearly visible in the reproductive cells
and the surrounding somatic cells (Figure 6A, n>25). Thus DMT102 is necessary to maintain
a deacetylated H3K9 state in a spatially limited domain of the ovule, which contains the
reproductive cells and expresses Dmt102 in sexual maize. Strikingly, the pattern seen in the
dmt102::Mu mutant line was also observed during apomeiosis and gametogenesis in
apomictic 38C plants (Figure 6A, n>25), with the spatial domain encompassing the
archespore/MMC similarly hyperacetylated.
Patterns of H3K9me2 in wild type B73 and dmt102::Mu ovules were similar during
gametophyte development (Supplemental Figure 5). However, H3K9me2 signals in
dmt102::Mu mature ESs differed from that observed in wild type (Figure 6, n=25). In wild
type mature ES, H3K9me2 signal was enhanced in the egg cell as compared to the central
cell. By contrast, in dmt102::Mu, H3K9me2 in the egg cell was significantly reduced relative
to central cell, suggesting that DMT102 activity is important for maintaining a repressive state
of egg cell chromatin.
3.2.7. Transcription patterns in the mature ES and the early seed suggest
quiescence in the egg cell and the sexual zygote, but strong activity in the apomictic proembryos
The differences detected at the level of histone marks suggested that apomictic and sexual
plants likely differ in transcriptional activity in and around the germ cells in the ovule. To test
this prediction, we analyzed global transcriptional patterns in ovules and early seeds of sexual
62

plants (Methods and Supplemental Figure 5). Staining of ovules containing a mature ES with
4H8, an antibody against the heptamer repeats in the C-terminal domain of the main subunit
of RNA POLYMERASE II (POLII), showed that most cells within the ovule contained some
degree of POLII (n>25), including the egg cell and the central cell (Figure 7A). However, a
clear difference could be observed between both cell types when immunostained with H5,
which recognizes the same heptamers as 4H8 when phosphorylated on serine 2.
Phosphorylation of ser2 occurs concomitantly with transcript elongation and, therefore, H5 is
a mark of POLII molecules engaged in transcription (Palancade and Bensaume, 2003). While
central cells showed strong staining for both 4H8 and H5, egg cells never produced a signal
detectable above background level with H5 (Figure 7A, n>25). Similar results were obtained
with an antibody against and H3K9ac and H3K4me3, two marks also broadly associated with
a transcriptionally competent chromatin state. This is consistent with the pattern of the
repressive mark H3K9me2 indicating a much higher level of repressed chromatin in the egg
cell than in the central cell (Figure 6 C, D). Thus, complementary information suggests that
the two gametes in the ES have very distinct transcriptional activity, with an active central cell
and a more quiescent egg cell coexisting in the ES.
To determine whether these two patterns were maintained following fertilization, we
probed 1 and 2 DAP seeds. Patterns of H5 (Figure 7B) were similar and consistent: the
dividing endosperm nuclei produced a strong signal, which was not detected in the zygote.
This indicates that the egg cell-to-zygote transition occurs without massive transcriptional
activation of the embryo genome.
To determine more precisely the timing of zygotic genome activation, we probed growing
seeds at various stages from 3 to 6 DAP with the same antibodies. The results showed that
most embryos at 3 DAP (>90%, n>50) had staining patterns similar to the zygote (Figure 7C),
suggesting that zygotic transcriptional quiescence lasted at least until 3 DAP. A striking
contrast, however, was clearly visible with most (>90%, n>50) embryos at 5 DAP where most
if not all cells in the embryo showed strong H5 (Figure 7C) and H3K4me3 staining.
Consistent with these observations, H3K9me2 staining showed a marked reduction at 5 DAP
(Figure 7C). These data collectively suggest that between 3 and 5 DAP the embryo genome in
sexual maize undergoes massive chromatin changes involving the release of repressive marks
and global transcriptional activation.
63

To further compare apomictic and sexual plants, we looked at transcriptional activity in
64

parthenogenetic embryos of the apomictic 38C ecotype. Pro-embryos in 38C develop
precociously as part of the maturation of the ES. Thus, the mature ES contains a pro-embryo,
arrested at the 16 to 64-cell stage, and an unfertilized central cell (Figure 7D). These
precocious arrested proembryos resume development following fertilization of the central
cell. Immunostaining of H5 and H3K4me3 (Figure 7D) showed a strong signal in all
proembryos (n>50) regardless of their size (from 16 to c.a. 60 cells). These results
demonstrate that the precocious parthenogenetic embryos are transcriptionally active.
3.2.8. Parthenogenetic pro-embryos have lost gametic identity, but have not
established embryonic patterning
During embryo development, specific patterns of genes expression are tightly regulated in
a spatial and temporal manner to either maintain or initiate changes in cell fate and
development. We were thus interested in checking whether the differences that we observed
between apomictic and sexual development correlated with changes in the expression patterns
of gametic or embryonic cell identity markers. In particular, it remained unclear based on the
above results whether the precocious embryo is an embryo or an over-proliferating gamete.
We analyzed the expression of a reporter gene consisting of the promoter region of the Zea
mays embryo sac1 (Zm ES1) gene fused to the green fluorescent protein gene (Cordts et al.,
2001; Figure 8A). In sexual maize, Zm ES1 is specific to the egg apparatus and its expression
decreases strongly immediately after fertilization (Cordts et al., 2001). In the apomictic
ecotype, expression was visible in the ES at the stage when the apomictic «gamete» consisted
of a single cell, but was absent in apomictic pro-embryos (n=20) (Figure 8A). This indicates
that pro-embryos lost egg cell identity as they transitioned from unicellular to multicellular
structures. To verify whether they acquired proper embryonic identity, we further used in situ
hybridization and localized transcripts of Zea mays WUSCHEL-related homeobox2 (Zm
Wox2) in parthenogenetic and sexual pro-embryo (Figure 8B). Zm Wox2 has been previously
characterized in maize (Nardmann et al., 2007) and shows a pattern similar to its Arabidopsis
homologue: it is expressed specifically in the apical cells of the pro-embryo and marks the
acquisition of clear apical-basal polarity in the embryo proper. Here, we found that Zm Wox2
expression was not specific to the apical cells in the apomictic pro-embryo but rather
expressed in the entire pro-embryo (Figure 8B). This suggests that the parthenogenetic proembryos
have not acquired the organization typically found in the sexual embryo.
65

3.3 DISCUSSION
By comparing transcription profiles of a diverse set of maize CMEs in sexual and
apomictic ovules, we identified a limited set of enzymes showing qualitative expression
differences between both reproductive strategies. Four of these CMEs (CHR106, DMT102,
DMT105 and DMT103) are predicted, or have been shown to control DNA methylation and
share close homology with the Arabidopsis DDM1, CMT3 and DRM2 proteins, respectively.
DDM1 is involved in the maintenance of methylation at both CG and non-CG sites, while
66

CMT3 and DRM2 genes act redundantly to maintain non-CG methylation. Finally, de novo
methylation is also a function of DRM2 in all sequences contexts (Cao and Jacobsen, 2002;
Cao et al., 2003). Among the many different families of CMEs tested, the differentially
expressed genes were not selected ex ante based on predicted functions and, therefore, it is
remarkable that most of them function in DNA methylation.
It has been advocated that apomixis might be a consequence of «epigenomic shocks», such
as interspecific hybridization and polyploidization, resulting in a broad deregulation of
reproductive development. Because our experimental model consists in an apomictic hybrid
of polyploid nature (as virtually all known apomictic plants) and interspecific origin, some or
all the variation detected in gene expression patterns might be only indirectly related to
apomixis. However, three arguments suggest otherwise. First, similar deregulation pattern of
DRM genes occurred in the two models we tested, Tripsacum dactyloides and Boechera
holbellii, indicating that it might denote a conserved feature among apomictic plants. Second,
deregulation affected genes with clear reproductive expression as dmt102 and dmt103
transcription domains in maize ovules were confined to the germ cells and a few surrounding
somatic nucellar cells. Finally, we also found that individual or combined downregulation of
both genes resulted in phenotypes highly reminiscent of apomictic developments. DMT102
loss-of-function resulted in the production of unreduced male gametes and in abnormal
patterns of antipodal cell differentiation. It further resulted, as in apomictic 38C, in a
hyperacetylated state of H3K9 in the archesporial domain of the ovule. Similar but more
severe phenotypes were observed for dmt103 mutant lines, which additionally resulted in
strong defects in ovule development: ESs with supernumerary gametes, formation of extra
ESs, possibly of both somatic and gametophytic origins, unreduced male gametes, and
possibly unreduced female gametes. The presence of multiple ESs suggests that DMT103,
and possibly DMT102, function in nucellar cells to ensure that a single gametophyte develops
within each ovule. In addition, the occurrence of either differentiated or undifferentiated extra
nuclei indicates that they might also regulate both cell identity and cell proliferation patterns
during megagametogenesis. In addition to this novel role in gametophytic development,
observation of unreduced gametes and supernumerary ESs is strikingly reminiscent of
apomictic reproduction. Therefore, we propose that the inactivation of this specific set of
genes might represent a crucial difference between apomixis and sexuality. Further work will
require to demonstrate that restoring their activity in an apomictic background can re-install
sexual reproduction.
67

Interestingly, the extra ES phenotype observed in dmt103 mutants is suggestive of
aposporous development, which is not found in diplosporous Tripsacum. It has been an open
question whether the different modes of apomictic development are genetically related. The
strongest argument for a close relationship between apospory and diplospory is the
coexistence of both forms in some species, for example in Paspalum minus (Bonilla and
Quarin, 1997). While it is possible that mutations allowing both developments might have
accumulated independently in the same ecotypes, our data support the more parsimonious
hypothesis that deregulating DNA methylation in reproductive cells participates in both
phenotypes.
The restricted expression domains of dmt102 and dmt103 also suggest the specialization, at
least in maize, of a DNA methylation pathway acting specifically in reproductive cells.
DMT102 in maize and the Arabidopsis homolog of DMT103 are involved in non-CG
methylation (Papa et al., 2001; Vaillant and Paskowski, 2007). Interestingly, DMT101, the
maize homolog of MET1 (the key enzyme for the maintenance of methylation at CG sites)
showed no difference in expression pattern in sexual and apomictic ovules. Although a direct
effect on non-CG methylation of DMT103 still requires confirmation, this points to a specific
role for non-CG methylation, and our results suggest that, during apomictic development in
Tripsacum, a small ovule domain containing the reproductive cells possibly endures
alterations in DNA methylation patterns at non-CG sites. Additionally, DRM2 is the main de
novo methyltransferase in all sequences contexts, and we cannot exclude the possibility that
de novo methylation activity might also be affected in apomictic progenies. Indeed, previous
data showed that apomictic reproduction faithfully reproduces the genome through
generations, but often fails to properly replicate DNA methylation patterns (Leblanc et al.,
2009).
Comparative cell-specific analysis of chromatin states within ovules revealed differences
between sexual maize on the one hand and dmt mutants and apomictic 38C hybrids on the
other. First, we observed that, in contrast to sexual maize, apomictic 38C plants and
dmt102::Mu lines suffered ectopic H3K9 hyper-acetylation in a nucellar domain overlapping
that of dmt102 transcription. While changes in H3K9ac are likely an indirect consequence of
DMT102 loss of function or apomixis, it represents a well-established mark of
transcriptionally competent chromatin. Thus, this indicates that transcriptional activity in and
around the germ cells in the ovule likely differs between apomictic and sexual plants.
Second, transcriptional activity in the early sexual embryo and the parthenogenetic embryo at
68

a similar stage of development strongly differed. Consistently with the aforementioned H3K9
hyper-acetylation, methylation states at H3K9 and H3K4 indicated that the mature female
gamete and the early seed in sexual maize are relatively quiescent until at least 3DAP, when a
major burst of transcriptional activity takes place. By contrast, parthenogenetic embryos
showed active transcription at early stages. We previously demonstrated using microarray
analysis (Grimanelli et al., 2005) that the mRNA populations present in ovules containing a
parthenogenetic embryo and in sexual maize ovules are essentially similar. This showed that
transcriptional activity in precocious pro-embryos was not the result of an early zygotic
genome activation. Furthermore, this heterochronic activation of transcription does not
correspond to the prolongation of a gametic program, as shown by the loss of egg cell-specific
marker expression, and likely interferes with the correct expression of embryo patterning
genes such as Zm WOX2. Whether the same difference in transcriptional activity occurs at
earlier stages of development was indirectly suggested by the differences observed for
H3K9ac during sporogenesis. Although technically difficult, determining directly the extent
of transcriptional repression during reproductive development remains critical in the
understanding of both sexual and apomictic reproduction.
What could be the role of a relative quiescence in the reproductive cells? In animals,
POLII-dependent transcription is repressed in the germ cells, a phenomenon which involves
both inhibition of phosphorylation in the CTD of POLII and large scale chromatin
remodeling. This repression is thought to be important for the establishment and maintenance
of germ-line fate, by preventing somatic differentiation (Seydoux and Braun, 2006). When
transcription is not repressed, germ cells do not form, due to the transformation of their
precursors into somatic cells. Transcriptional repression may also be key to establish
transcriptional profiles compatible with totipotency (Seydoux and Braun, 2006). Plants do not
have germ-lines as such. Rather, somatic cells switch programs late during development to
produce reproductive cells. Furthermore, the products of meiosis, which directly develop into
gametes in animals, undergo additional cell division and differentiation steps to form
multicellular gametophytes containing the gametes. Despite these differences, recent data
indicate that animal and plant germ cells development share mechanistic similarities, as
illustrated by the essential silencing function of DDM1 in plant male gametes, reminiscent of
piRNA pathway function in animals (Slotkin et al., 2009) and the importance of Argonaute
proteins members in both systems (Nonomura et al., 2007; Yin and Lin, 2007). Thus, is a
relative transcriptional quiescence required for (sexual) plant female gametes definition?
69

Here, we observed that parthenogenetic embryos, which develop without fertilization, are
transcriptionally active, by contrast to sexual ones. Whether continuous transcription is
sufficient to induce parthenogenesis is unclear; such a gain-of-function phenotype does not
offer clear functional evidences. Yet, the data on both dmt- mutants suggest strongly that both
gametophyte and egg cell development requires transcriptional quiescence.
The genetic mechanism(s) underlying apomictic developments in plants remains
undetermined. Specific genes or alleles controlling apomixis expression in any plant species
have yet to be discovered. Although important advances have been made recently with the
engineering of apomeiotic genotypes in Arabidopsis (Ravi et al., 2008; d'Erfurth et al., 2009),
it is still unclear whether these phenotypes are related to apomixis in wild species.
Furthermore, the genotypes generated so far in Arabidopsis alter sporogenesis without
affecting gametogenesis or embryogenesis. Here, we took the reverse approach and analyzed
apomixis regulation in an artificial interspecific hybrid as well as in natural ecotypes. Our
data suggest that the downregulation of a reproductive RdDM-like pathway can result in both
sporogenesis (unreduced gametes) and gametogenesis (extra embryo sacs) alterations. This is
in line with the fact that apomeiosis and parthenogenesis in Tripsacum are genetically linked
(Leblanc et al., 2009). On the other hand, neither DMT102 nor DMT103 inactivation resulted
in the production of fully apomictic maize progenies. To validate our hypothesis that the
downregulation of an ovule-specific chromatin-based silencing pathway in maize would
generate apomictic reproduction, it will be necessary to identify the other members involved
in this pathway and to explore the effects of their inactivation either individually or all at
once.
70

3.4 METHODS
3.4.1. Plant Materials
The 38C apomictic maize-Tripsacum hybrid has been previously described in details
(Leblanc et al., 1996; Grimanelli et al., 2003, Leblanc et al., 2009). Transgenic version of 38C
expressing pES1-GFP were produced as described by Leblanc et al. (1996) using a pZm ES1-
GFP maize line provided by Thomas Dresselhaus (University of Regensburg, Germany).
Plants carrying a dmt102::Mu defective allele (zmet2-m1::Mu; Papa et al., 2001) were
provided by SM Kaeppler. In all experiments, we used seeds derived from a single
dmt102::Mu/dmt102::Mu plant. The dmt103 RNAi materials (D103-2 and D103-4) were
generated by the Maize Chromatin Consortium (for vectors and procedures, see McGinnis et
al., 2005). These lines, the dmt103-6 tilling allele, stocks for the B73 maize inbreed and
N108A stocks (autotetraploid line derived from B73) were provided by the Maize Genetic
Cooperation Stock Center. The Tripsacum dactyloides 65-1234, which was used to derive 38C
is maintained at CIMMYT, Mexico. Experiments in Arabidopsis thaliana were done with the
Col(0) ecotype. The apomictic Boechera holbollii accession was obtained from Enrico Perotti
(Australian National University, Canberra).
3.4.2. Genotyping
The genotype of dmt102 locus was determined by PCR using a Mu specific primer and a
dmt102 internal primer as described in Papa et al. (2001). To genotype dmt103 RNAi lines,
we took advantage of the herbicide resistance gene present in the transgenic construct
(McGinnis et al., 2005). We first screened for herbicide resistance a pool of T2 plants and
verified by PCR the presence of the transgene. We then maintained hemizygous plants by
crossing transgenic T2 plants as male or female progenitors with B73 plants. All subsequent
analyzes were performed in the resulted T3 segregating progeny, following the same
procedure (herbicide resistance and transgene genotyping). Genotyping of the TILLed
dmt103-6 allele is described elsewhere (www.genome.purdue.edu/maizetilling). Primers
sequences for genotype analysis are listed in Supplemental Table 3.
71

3.4.3. Selection of CMEs for RT-PCR analysis
To identify CMEs expressed during sexual and/or apomictic ovule development, we reanalyzed
a set of microarrays from which global transcription profiles have been previously
reported (Grimanelli et al., 2005). The original experiment compared differential expression
between ovules at the mature embryo sac stage in apomictic and sexual plants. We used the
results of BLAST searches to identify probes in the arrays corresponding to CMEs expressed
either in sexual plants, apomictic plants, or both. Out of these, we selected all loci (11)
showing putative differential expression between the two reproductive modes. Next, we
identified using The Chromatin Database (www.chromdb.org) an additional set of 45 genes,
with available RNA expression patterns suggesting higher expression in reproductive tissues
than in vegetative parts. ID, loci accession numbers and the corresponding protein
classification are shown in Supplemental Table 1 and 2.
3.4.5. RT-PCR
Immature ovaries were collected and carefully dissected to eliminate non-reproductive
tissues. Bulks of isolated ovules were directly frozen in liquid nitrogen for further nucleic acid
extraction. In order to precisely determine the developmental stage within bulks (see
Supplemental Figure 1 for maize), small samples were set apart prior to freezing and cleared
using benzyl:benzoate solutions (maize and Tripsacum; Leblanc et al., 1995) or Herrs’s
solution (Arabidopsis and Boechera) to precisely determine the developmental stage of each
sample. RNA extractions were performed with TRIZOL reagent (Invitrogen) followed by
DNAse treatment (Deoxyribonuclease I, Invitrogen). RNA was reverse transcribed using the
SUPERSCRIPT III RT-PCR kit (Invitrogene), following the provider’s instructions. For PCR,
we used 10µl reactions containing 1µl of cDNA, 2µl of ReadyMix TM Taq PCR Reaction Mix
(Sigma-Aldrich), 1µl of 10µM each forward and reverse primers and 5µl Millipore water.
PCR cycling program included 35 cycles of 94 o C for 30s, 60 o C for 30s and 72 o C for 30s,
followed by an additional cycle of 2min at 72 o C. For all experiments, we used maize actin1 or
Arabidopsis ACTIN11 as a cDNA amplification control. Primers were designed to span an
intron, when possible, in order to identify genomic DNA contaminations. Primer sequences
are provided in Supplemental Table 3.
72

3.4.6. Imaging of histone modifications and transcriptional activity in ovules and
seed tissues
We used an approach first described by Laurie et al. (1999), referred to as "semi-isolated"
ESs. Fresh ovules or 1 DAP seeds were sliced with a vibratome (LeicaVT1000E) to produce
sections containing intact ESs or developing embryos and endosperms. As described in the
original article, these sections contain living materials that can be either fixed for further
processing or grown onto a basic growth medium. In our hands, most in vitro cultured
sections harvested post fertilization (>60%, n>100) mimicked normal seed development
during at least 6 DAP, therefore covering the time frame of our experiments. This allowed
precise sampling and 3D imaging of intact, whole-mount ES, developing embryos and
syncitial endosperm tissue (Supplemental Figure 6). We thus combined this procedure and
immunostaining to monitor transcriptional activity and histone modifications during
reproductive cells development. To achieve this, we used commercial antibodies (Abcam)
available for transcriptional activity (4H8 and H5) and for several histone marks, i.e.
H3K9me2, H3K9ac and H3K4me3. All these antibodies are routinely used in other organisms
and produced (see below) results consistent with the literature (Supplemental Figure 6).
Embryo sacs and early seeds were prepared following Laurie et al. (1999). Vibratome sections
(c.a. 300 µm) were fixed for 3 hours in 4% paraformaldehyde: 1xPBS: 2% Triton fixative,
washed twice in 1xPBS. Samples were digested in a enzymatic solution (1% driselase, 0,5%
cellulase, 1% pectolyase, 1%BSA, all from Sigma) for 15 to 45 min at RT, depending on the
developmental stage, subsequently rinsed 3 times in 1xPBS and permeabilized for 1 to 2 h in
1xPBS, 2% Triton, 1% BSA. They were then incubated overnight at 4°C with the primary
antibodies (all from Abcam), used at the following concentrations: 1:400 for H3K9ac, 1:200
for 4H8, H5, H3K9me2 and H3K4me3, The sections were washed daylong in 1xPBS; 0,2%
Triton and incubated with secondary antibodies (either FITC conjugate or Cy3 conjugate,
both from Sigma, or Alexa Fluor 488 conjugate, Molecular Probes) used at 1:400 dilution.
After washing in 1xPBS; 0,2% Triton for a minimum of 6 h, the sections were incubated with
DAPI (1ug/ml in 1xPBS) for 1h, washed for 1h in 1xPBS and mounted in PROLONG
medium (Molecular Probes). A minimum of 50 ovules or seeds with interpretable staining
patterns was scored for each developmental stage reported in this paper. Images were captured
on a LEICA epifluorescence microscope equipped with a color CCD Leica camera and
appropriate DAPI, FITC or Cy3 filters. 3D ovule images were captured on a laser scanning
confocal microscope (Leica SP2) equipped for 405nm (DAPI), 488 nm (FITC/Alexa488) and
73

525 nm (Cy3/Alexa568) excitation and either 20X, 40X or 63X objectives. Maximumintensity
projections of selected optical sections were generated for this report and then edited
using Graphic Converter (lemkeSOFT).
3.4.7. Whole mount clearing of ovule samples
Ovaries were fixed in FAA (50% absolute ethanol, 5% glacial acetic acid, 10%
formaldehyde and 35% H2O) for 24h at RT and stored in ethanol 70%. Fixed ovaries were
sectioned by vibratome as described above. Then, sections were dehydrated throughout
successively ethanol dilutions of 70%, 85% and 100% and cleared according to Leblanc et al.
(1995). Cleared sections were mounted in the clearing solution between two blocks of
coverslips with similar thickness to that of samples and covered by a third coverslip. Samples
were observed with differential interference-contrast optics (DIC) in a ZEISS Axio Imager.A1
microscopy.
3.4.8. Whole mount in-situ mRNA hybridizations
Vibratome sections of fresh ovaries were recovered and fixed as for immunolocalization
assays. We then followed the standard in situ protocol as described Garcia Aguilar et al.,
(2005) using (LNA) DNA probes which substitutions were as follows: dmt102:
TAGGAALTGTCZCGTCCLAC, dmt103: TCGATCTZGTGATZGGTGGEAGTC, Zm
Wox2: CGCCGLGCCCPGCTCTCCEGC, with E=A-LNA, L=C-LNA, Z=T-LNA, P=G-
LNA.
74

3.5. TABLES
Table 1. BLAST analysis of selected maize CMEs with the Arabidopsis protein database.
The maize locus nomenclature is according to CHROMDB. % of identity was calculated
using the complete length of the predicted maize protein.
Protein Maize At homologue At gene Function % E values
Family locus
Identity
SNF2 chr106 AT5G66750 DDM1 Maintenance of Cyt methylation 63 e-126
DMT dmt102 AT1G69770 CMT3 Maintenance of CNG methylation 50 0
DMT dmt103 AT5G14620 DRM2 de novo methylation 51 e-151
DMT dmt103 AT5G 15380 DRM1 de novo methylation 51 e-149
DMT dmt105 AT4G19020 CMT3 Maintenance of CNG methylation 51 0
HDA hdt104 AT3G44750 HD2A Histone Deacetylase 42 2e-51
Histone hon101 AT2G18050 HIS1-3 Histone linker 52 e-24
75

Table 2. Ovule phenotypes observed in individuals from dmt103 single and double
mutants. ES: embryo sacs
PHENOTYPES
Lines # WT Abnormal Aborted Extra ES n
D103-2 4 28 6 0 4 38
D103-2 7 10 0 4 0 14
D103-2 11 22 12 2 7 43
D103-2 16 25 2 0 2 29
D103-2 22 22 3 0 1 26
Total 107 23 6 14 150
% 71 15 4 9
D103-4 2 21 5 0 0 26
D103-4 3 2 0 33 0 35
D103-4 4 17 13 0 0 30
D103-4 6 30 0 0 0 30
D103-4 8 1 12 0 0 13
D103-4 9 9 2 0 0 11
D103-4 10 6 0 0 1 7
Total 86 32 33 1 152
% 57 21 22 1
dmt103-6 1 3 15 0 3 21
dmt103-6 4 7 14 0 1 22
dmt103-6 6 5 12 0 4 21
Total 15 41 0 8 64
% 23 64 0 13
Dmt102/dmt102::Mu D103.4 12 9 0 0 8 17
% 53 0 0 47
F2#3 1 16 1 0 0 17
F2#3 6 18 1 0 0 19
F2#3 11 19 0 1 0 20
F2#3 12 28 0 2 0 30
Total 81 2 3 0 86
% 94 2 3 0
F2#12 1 10 6 0 0 16
F2#12 2 9 4 1 0 14
F2#12 3 4 6 0 1 11
F2#12 4 6 1 1 2 10
F2#12 5 12 1 0 11 24
F2#12 6 8 2 0 0 10
F2#12 7 28 0 0 0 28
Total 77 20 2 14 113
% 68 18 2 12
76

Table 3. Embryo sac phenotypes observed in individuals from dmt103 single and double
mutants. nuc: nucellus; ES: embryo sacs; An: antipods
PHENOTYPES
Lines # Normal Extra nuc. Inverted ES Abnormal An Multiple ES n
D103-2 1 28 2 0 0 2 32
D103-2 6 25 0 0 0 1 26
D103-2 11 22 2 1 0 8 33
D103-2 16 25 0 0 2 0 27
D103-2 17 18 0 0 2 0 20
D103-2 18 27 0 0 1 0 28
D103-2 22 22 1 0 0 2 25
D103-2 23 13 1 0 2 0 16
D103-2 24 21 0 0 1 0 22
Total 201 6 1 8 13 229
% 88 3 0 3 6
DMT103-4 4 6 0 0 0 1 7
DMT103-4 8 1 0 0 0 0 1
DMT103-4 9 9 0 2 0 0 11
Total 16 0 2 0 1 19
% 84 0 11 0 5
Dmt102/dmt102::Mu
D103.4 12 10 0 0 0 7 17
% 59 0 0 0 41
77

3.6. SUPPLEMENTAL TABLES AND FIGURES
Supplemental Figure 1. Definition of developmental stages in the ovule.
Supplemental Figure 2. Expression patterns of selected CMEs during sexual and apomictic
reproduction as determined by RT-PCR.
Supplemental Figure 3. Cladogram showing the relationship among selected DMTs in maize
and homologs in Arabidopsis.
Supplemental Figure 4. Phenotypes of D103 mutant lines.
Supplemental Figure 5. H3K9me2 detection during megagametogenesis in sexual maize
(B73) and dmt102 mutant lines and in 38C apomicts.
Supplemental Figure 6. Analysis of chromatin states in whole mount seed tissues.
Supplemental Table 1. List of CMEs evaluated for for reproductive tissue expression.
Supplemental Table 2. CMEs selected for transcriptional analysis.
Supplemental Table 3. List of primers.
78

MMC
INT
Early sporogenesis
Mature embryo sac
PN
Embryo
Early embryogenesis
Supplemental Figure 1. Definition of developmental stages in the ovule.
79

A
GENE SEXUAL APOMICTIC
SPO MES 3DAP LEAF SPO MES 3DAP LEAF
brd101
brat101
chb102
chc101
chr101
chr106
chr110
chr112
chr113
chr120
chr125
chr126
dmt101
dmt102
dmt103
dmt104
dmt105
dmt106
dmt107
hxa102
hag102
hac 101
AC157319
BT066941
AY110756
AY104727
EY951031
hda102
hda110
hdt101
hdt103
hdt104
DR958383
srt101
hon101
hon104
mbd101
mbd109
nfa101
nfa104
nfc102
nfc104
nfe101
sdg101
sdg102
sdg107
sdg110
sdg118
sdg122
sdg129
DT942012
AW288923
DR959742
EE168341
EC897242
vef101
4
3
2
1
0
B
100%
90%
80%
70%
60%
50%
40%
30%
20%
10%
0%
100%
90%
80%
70%
60%
50%
40%
30%
20%
10%
0%
100%
90%
80%
70%
60%
50%
40%
30%
20%
10%
0%
SPOROGENESIS
3
3
3
5
42
sexual
9
7
2
11
27
apo
MATURE EMBRYO SAC
2 7
2
2
1
6
9
44
sexual
3 DAP
38
apo
4
1
7
6
5
1
9 7
36 36
sexual
apo
Supplemental Figure 2. Expression patterns of selected CMEs during sexual and apomictic
reproduction as determined by RT-PCR.
(A) Profiling of individual CMEs in ovules at three developmental stages and in leaf tissue as
sporophytic control. SPO: meiosis/apomeiosis, MES: mature embryo sac, DAP: days after pollination.
Color code for the classification of transcription expression is shown below. Red (0) indicates null
amplification while dark green (4) represents the highest expression levels..
(B)Global transcriptional behavior during sporogenesis, mature ES and 3 DAP in sexual and apomictic
plants. The number of genes corresponding to each color code is indicated in bars.
80

Supplemental Figure 3. Cladogram showing the relationship among selected DMTs in maize and
homologs in Arabidopsis. The tree was build using MacVector version 7.2 (ClustalW procedure,
Neighbor Joining method). Numbers indicate distance.
81

A
B73
B
B73
ES
B73 X D103-2
Ov
D103-2
ES
Ov
B73 X D103-2
D103-4 X B73
D103-2
Ov
ES
D103-4 X B73
Supplemental Figure 4. Phenotypes of D103 mutant lines.
(A) Ears from reciprocal crosses between wild type maize (B73) and D103 lines.
(B) Abnormal ovule development resulting in mispositioned of mature ESs in D103-2 lines. In both
cases, rotation
Supplemental
in the ovule
Figure
fails
4.
to
Phenotypes
complete,
of
and
D103
development
mutant lines.
of the gametophyte is arrested at early
stage. ES: (A) embryo Ears from sac; reciprocal Ov: ovary. crosses between wild type maize (B73) and D103 lines.
(B) Abnormal ovule development resulting in mispositioned of mature ESs in D103-2 lines. In both
cases, rotation in the ovule fails to complete, and development of the gametophyte is arrested at
early stage. ES: embryo sac; Ov: ovary.
82

B73 dmt102::Mu 38C
Supplemental Figure 5. H3K9me2 detection during megagametogenesis in sexual maize (B73)
and dmt102 mutant lines and in 38C apomicts. Green: H3K9me2; Blue: DAPI.
83

A
B
End
ES
Emb
C
Nuc
Ant
4 DAP endosperm nuclei
4H8
Mi
DAPI
Supplemental Figure 6. Analysis of chromatin states in whole mount seed tissues.
Whole mount tissues consisted in vibratome sections of living ovules or seeds. As shown in (A),
ovules sections contained intact ESs within nucellus. Cultured sections produced from 1DAP seeds
and visualized after 4 days of growth recapitulated in vivo development of both the endosperm (End)
and the embryo (Emb) as shown in (B). (C) shows whole mount endosperm and embryo tissues after
confocal imaging of a DAPI-stained seed section collected at 3 DAP (Ant: antipodal cells; Nuc:
nucellar cells; Mi: micropyle). Efficiency testing of immunostaining on vibratome sections of maize
developing seeds was performed by examining the staining patterns of well characterized antibodies in
dividing cells within the nucellus (the maternal tissue surrounding the female gametophyte) or in
dividing endosperm nuclei. Since mitosis involves a generalized repression of gene expression
(reviewed in Gottesfeld and Forbes, 1997), we first used two antidodies raised against the carboxyterminal
domain (CTD) repeats of the main sub-unit of RNA polymerase II (POLII) : H5 that
recognizes the phosphorylated form on serine 2 of the CDT, a widely accepted landmark of POLII
molecules engaged in transcription (Palancade and Bensaume, 2003), and 4H8 that recognizes POLII
CTD repeats regardless of ser2 phosphorylation and thus detects both isoforms (active and inactive) of
the protein. The level of sequence conservation at the CTD of POLII among eukaryotes, as well as
experimental evidences, indicate that both antibodies recognize similarly plant and animal POLII
isoforms (Koiwa et al., 2004; Gullerova et al., 2006). As shown in (D), 4H8 strongly stained
84

interphase nuclei (n>500) but showed decreasing staining patterns as nuclei entered into mitosis and
was finally undetectable on metaphase chromosomes (arrow). Similarly, H5 was never detected on
mitotic chromosomes. The pattern on interphase nuclei was more complex, with a graduation from
nuclei showing a bright staining to undetectable signal. These results were then mirrored by the
staining patterns of antibodies raised against posttranslational histone H3 modifications functionally
associated with chromatin transcriptional competence: H3K4me and H3K9ac for active states and
H3K9me2 for repression.
85

Supplemental Table 1. List of CMEs evaluated for reproductive tissue expression.
Protein Class (1) Genes (2) Selected CME Protein Class (1) Genes (2) Selected CME
AGO 16 0 HMGB 11 0
ARP 6 0 HON 6 2
BRAT 1 1 HTA 16 0
BRD 4 1 HTB 13 0
CHB 4 1 HTR 14 0
CHC 2 1 HUPA 1 0
CHE 1 0 HUPB 2 0
CHR 44 8 HXA 2 1
CPC 2 0 JMJ 4 0
CPD 1 0 MBD 14 2
CPG 1 0 MFP 1 0
CPH 1 0 NFA 5 2
CRD 1 0 NFB 1 0
DCL 4 0 NFC 5 2
DMT 7 7 NFE 1 1
DNG 4 0 NFF 3 0
DRB 6 0 PAFA 1 0
EBP 5 0 PAFB 1 0
EPL 1 0 PAFC 1 0
GTA 3 0 PAFD 1 0
GTB 1 0 PAFE 1 0
GTC 1 0 PGE 2 0
GTE 11 0 PRMT 2 0
GTF 1 0 RBL 2 0
GTG 1 0 RDR 4 0
HAC 5 1 RPDA 2 0
HAF 1 0 RHEL 2 0
HAG 6 1 SDG 35 7
HAM 2 0 SGA 3 0
HCP 2 0 SGF 1 0
HDA 10 2 SMH 6 0
HDMA 3 0 SNT 2 0
HDT 4 3 SRT 1 1
HEN 1 0 SWDB 3 0
HF0 15 0 SWDC 2 0
HIRA 2 0 TAFV 1 0
HMGA 3 0 VEF 3 1
Protein class information (1) and counts (2) are according to ChromDB.
86

Supplemental Table 2. CMEs selected for transcriptional analysis.
LOCUS PROTEIN GROUP SOURCE
brd101 Diverse bromodomain ChromDB
brat101 Bromodomain with AAAATPase ChromDB
chb102 SWI/SNF and RSC chromatin remodeling complex (Swi3/Rsc8) ChromDB
chc101 SWI/SNF and RSC chromatin remodeling complex (Swp73/Rsc6) ChromDB
chr101 SNF2 Super family ChromDB
chr106 SNF2 Superfamily ChromDB
chr110 SNF2 Superfamily ChromDB
chr112 SNF2 Superfamily ChromDB
chr113 SNF2 Superfamily ChromDB
chr120 SNF2 Superfamily ChromDB
chr125 SNF2 Superfamily ChromDB
chr126 SNF2 Superfamily ChromDB
dmt101 DNA Methyltransferase ChromDB
dmt102 DNA Methyltransferase ChromDB
dmt103 DNA Methyltransferase ChromDB
dmt104 DNA Methyltransferase ChromDB
dmt105 DNA Methyltransferase ChromDB
dmt106 DNA Methyltransferase ChromDB
dmt107 DNA Methyltransferase ChromDB
hxa102 Histone acetyltransferase ChromDB
hag102 Histone acetyltransferase ChromDB
hac 101 Histone acetyltransferase ChromDB
AC157319 Histone acetyltransferase Microarrays
BT066941 Histone acetyltransferase Microarrays
AY110756 Histone acetyltransferase Microarrays
AY104727 Histone acetyltransferase Microarrays
EY951031 Histone acetyltransferase Microarrays
hda102 Histone deacetylase ChromDB
hda110 Histone deacetylase ChromDB
hdt101 Histone deacetylase ChromDB
hdt103 Histone deacetylase ChromDB
hdt104 Histone deacetylase Microarrays
DR958383 Histone deacetylase Microarrays
srt101 Histone deacetylase ChromDB
hon101 Histone H1 ChromDB
hon104 Histone H1 ChromDB
mbd101 Methyl binding domain ChromDB
mbd109 Methyl binding domain ChromDB
nfa101 Nucleosome/chromatin assembly complex ChromDB
nfa104 Nucleosome/chromatin assembly complex ChromDB
nfc102 NURF complex ChromDB
nfc104 NURF complex ChromDB
nfe101 Nucleosome positioning into regular spaced arrays ChromDB
sdg101 SET domain ChromDB
sdg102 SET domain ChromDB
87

sdg107 SET domain ChromDB
sdg110 SET domain ChromDB
sdg118 SET domain ChromDB
sdg122 SET domain ChromDB
sdg129 SET domain ChromDB
DT942012 SET domain Microarrays
AW288923 SET domain Microarrays
DR959742 SET domain Microarrays
EE168341 SET domain Microarrays
EC897242 SET domain Microarrays
vef101 VEF family (VRN2,EMF2,FIS2) ChromDB
88

Supplemental Table 3. List of primers.
PRIMERS SEQUENCES PRIMERS SEQUENCES
Brd101F CTGGTTATGGGGGTACATGG Hdt103R TGCTACATCGAACGCATAGC
Brd101R ATTGCTGGCTTGCTAATGCT Hdt104F TGAAGAGGTTGAACCCGATCT
Brat101F CCCTCCAACCTTCCAGTTTT Hdt104R TATCGCTTGACACTTCATCTTCA
Brat101R CAGAAACCAGCTGAGCAACA DR958383F GATGAGCCGAGCAATGTTAAG
Chb102F TCATGGGACACAAGGAGGAT DR958383R CATGGGTCAGACTACCGTCA
Chb102R GGGACGCCAAATTTAACAGA Srt101F GATGATGCTTTACCCCCTGA
Chc101F GCCAACACAGAGAAGCACAA Srt101R AACAGCTGCAACCATCACTG
Chc101R CACACCAAGCAACATTACGG Hon101F CATCTCCCTCTCCGAAATCA
Chr101F CAAGGTGCAGGAAGAAGAGG Hon101R CACATAACACACCGCCTGAG
Chr101R GGGTGTTCCAGTCAAAAGGA Hon104F AGGAGAAGAAGGCGAGGAAG
Chr106F GAGCGGATCATCAAGAAAGC Hon104R GAGCGCATCACACCAGACTA
Chr106R CATACACGCGGACACAAATC Mbd101F CCCCTCTCAGAGAAAACGAA
Chr110F CTCATGCAAGCAAACCAGAA Mbd101F CGGGTTCGGCCCCCTCTCAG
Chr110R GTGGTCAGCCTCTCTTACGC Mbd109F ACGGGCAAGATCAAGCTCTA
Chr112F ATCGTGCGACCAGAGGATAC Mbd109R CGTCATCAACAAATGCAAGG
Chr112R AGCTCGGCTGATAACCAGAA Nfa101F GCCATGAAAACAAACGAGGT
Chr113F GCGAATGATGCAGCTTACAA Nfa101R CTTGCACAGCCTCACCAGTA
Chr113R TGAAATGTCATCCTCCAATGAG Nfa104F ACACAGCGACGCTAAAAACC
Chr120F GCTGCAAAGCACACTTACCA Nfa104R ATCTTGCACACCATCAGCAA
Chr120R AACTTTGGCAGGTCACCATC Nfc102F TTTGGGATGTTGAAGCACAA
Chr125F CCAGAAGTACCGGAGGTCAA Nfc102R TATGACCAGCGTGCTGAAAG
Chr125R CTGCAGCACACCTCATCAGT Nfc104F AGGGTTATGGCTTGTCATGG
Chr126F TCCAACGTTTTTGCCCTATC Nfc104R CTGGCTGCAGTGAACAAAAA
Chr126R ACAAAAAGGGATGGTTGCAG Nfe101F AGTAAGGCCCAATGCAGATG
Dmt101F TTGGGGATCTGCCTAAAGTG Nfe101R CCCTTATGGCAGAGGTTTCA
Dmt101R CTTGAGCTTCCTCCCAAGTG Sdg101F AGACAGGAGATCGTGGTTGG
Dmt102F GTTGGTTCTTCCGAGCAGAG Sdg101R TGCATGGAACACCTAGGACA
Dmt102R GCTTTCTCATTTCGCACCTC Sdg102F CGAGAATGGGTCAACTGGAT
Dmt103F CTTCTAGTTGTGCCCCCAAA Sdg102R TTCAACCCGTTCTGCTCTCT
Dmt103R CCTTTTGGAGCAAGAGCAAC Sdg107F GGGTGCTGAAGGATGAAGAA
Dmt104F CTATGTGCCTGAGCCTGACA Sdg107R TTCACGTTCTGCAGTCAAGC
Dmt104R TCAACGGATCACAATTCCAA Sdg110F GCGGCCCTCTTTATCTCTCT
Dmt105F TACTTGCCGTTGGTTTTTCC Sdg110R CCAGCCTTTTCATTGTCCAT
Dmt105R GCTTTCTCATTTCGCACCTC Sdg118F AGCAAGAGGCTGAAGCTGAG
Dmt106F AGCTTGAAGGAGGAGGAAGG Sdg118R ACCAGATGGAGGAACAGGTG
Dmt106R ATAGACCATGGATCGCCAAG Sdg122F TCTTGAAGGTGCTCGATGTG
Dmt 107F GACAGATCGCTGCCTACACA Sdg122R ATCTCTCATTGCGTGCCTCT
Dmt 107R ACCTCTGGTGTGATCCTTGG Sdg129 F AGCATGATGACGCTGCTGT
Hxa102F GGAATGTCGTTCTGCTGGAT Sdg129R TGTCACCCTAGGTGCTGATG
Hxa102R GACCGTAGGCAACTCCACAT DT942012F CCACCAAGTGGCAACAGAAT
Hag102F CATGGATGGGAAATGCTTCT DT942012R ATTTCTGCGGCGATAACATC
Hag102R ATCTGCAACAGACGCTTCCT AW288923F GGAGAAGCATGACCAGAAGC
Hac101F TCAATTGATGGATTGCCTGA AW288923R GTAAGCTGCCTCCATTACGC
Hac101R AAGAGGCAAACATCCACACC DR959742F GTGCTATTTGGGAGGTCTGC
AC157319F CTACGGCAACTCCATCTTCC DR959742R GCCTGTCAATGTCATGATCG
AC157319R GCATCAACGAAACAGAAGCA EE168341F GCTCTTAGAGGACGGAGTCG
BT066941F GAGAAGCCTGACTTGGCACT EE168341R TTCCTGGTGGGCATCATT
BT066941R CACCAAGCATTGTTGCCATA EC897242F CCTCACTCGAGACAGACATGG
AY110756F CGAGCCCTATTCCATCTTCA EC897242R ATGGTTAGCCGATTGCAAAA
AY110756R GAAGCATGAACAGAGCGAAA Vef101F CATGCGAGGAGGAAAAAGAG
AY104727F GGAAGGTTTCATGACTGCAA Vef101R GAACCCACAGCATCAACCTT
AY104727R AAGCAAGGTAGAAGCAAGCA WAXYINTRONF CCACCCAAGAAACTGCTCCTTAAG
EY951031F CCGAGGGAGTTTTAGGCAGT OCS3'R CGGCGGTAAGGATCTGAGCTACAC
EY951031R TTGCTTGCAACTTGTCCATT ACT1F CCTGAAGATCACCCTGTGCT
Hda102F TTCTTGCACCGGATAACTCC ACT1R GCAGTCTCCAGCTCCTGTTC
Hda102R AGTTTCAACAGCCCAACACC DRM2F CCCACCTGAGTTTGTGGACT
Hda110F CAATGCCAAATGTGGACAAG DRM2R CACTTCCCCACCACCAATAC
Hda110R GAGGTCTTACCAGCGCAAAG DRM1F GGATGGGAGAACGAAGTTGA
Hdt101F TGGCTACAAGTCCCCTGTTC DRM1R GAATCCTCCGTACTCGTCCA
Hdt101R GATGGACTGCTTTGCAGGAT CMT3F CATGGGAGCCTATTGAAGGA
Hdt103F CAGTTGCTGCACCTTCAAAA CMT3R TGCACCCCATAGAAAGAACC
89

90
CHAPTER 4. Discussion and Perspectives

In this study we showed that some CMEs are essential in regulating sexual development in
maize and, likely, in the phenotypic difference between sexual and apomictic reproduction.
Although transcription profiles in sexual and apomictic ovules at the mature embryo sac stage
showed limited significant differences, a number of CMEs were down regulated, specifically
during apomeiosis. These CMEs are predicted to control both DNA and histone methylation,
and encode essential components of the RdDM pathway in Arabidopsis, a RNA silencing
pathway involved in DNA methylation, imprinting, and plant development. Our results
demonstrate that loss of function of two DNA methyltransferases, DMT102 and DMT103,
gives rise to apomictic features in sexual diploid maize. Loss of function in these genes results
in aposporic-like embryo sac formation and in strong structural defects of both the ovule and
the gametophyte, including the production of extra female gametes into individual embryo
sacs. In maize, only few gametophytic mutations have been characterized (Neuffer et al.,
1997) and, among them, ig1 is of particular interest with regard to our observations: it causes
the formation of multiple female gametes within embryo sacs that arise by extra mitosis
during gametogenesis (Evans, 2007; Huang and Sheridan, 1996). By contrast, extra polar
nuclei formation in dmt103 mutants occurs in the mature embryo sac, most likely from extra
karyokinesis of polar nuclei once the central cell fate has been acquired. Determining
precisely cell identity in some extra nuclei is a difficult task due to the lack of appropriate
markers. However, using this approach in Arabidopsis, Johnston et al. (2008) showed that the
retinoblastoma-related protein RBR is essential for the differentiation of gametophytic cells:
rbr mutations produce numerous unfused polar nuclei, a phenotype, which strongly resembles
that of dmt103. However, rbr mutants neither develop extra embryo sacs nor affect ploidy
levels in pollen and the effects on cellular differentiation are limited to individual embryo sacs
and to the vegetative cell in pollen grains (Ebel et al., 2004; Johnston et al., 2008). Therefore,
despite similarities among the phenotypes displayed by the three mutants, we propose that
DMT103 acts in an independent regulatory pathway of that of IG1 or RBR1.
The fact that both dmt102 and dmt103 are exclusively expressed in and around the germ cells
in the ovule supports the hypothesis for a novel regulatory pathway controlling the
differentiation of reproductive cells. Immunolocalization experiments showed that loss of
function for either genes results in ectopic H3K9 acetylation in the germ cells and the
surrounding tissues and, in alterations in the dimorphic transcriptional patterns observed in
the egg and the central cell in wild type embryo sacs. Interestingly, ectopic transcription was
91

also observed in unfertilized embryo sacs in apomicts suggesting that a loss of transcriptional
quiescence might be implicated in precocious embryo development. Concomitantly, cell
identity and transcriptional silencing marks in developing parthenogenetic proembryos were
altered. The putative function of these genes in transcriptional silencing makes them attractive
candidates to establish a functional relationship with apomixis. Curiously, their homologs in
Arabidopsis and rice are expressed more constitutively (Sharma et al., 2009) and loss-offunction
alleles display no obvious phenotypes. In maize, however, the clear developmental
phenotypes of DMT102 and DMT103 probably define, together with other as yet unknown
proteins, a RdDM-like pathway specific to the germ line. Unexpectedly, the phenotypes
observed in mature ovules of both mutants were related to aposporic development, rather than
to our diplosporic apomictic model, suggesting that natural apomictic systems might share
similar regulatory components of epigenetic nature, which may be crucial for dynamically
(i.e., temporally and spatially) regulating gene expression during development. This might
provide also an explanation for shared common abnormalities between both apomictic
mechanisms (Bicknell and Koltunow, 2004).
The in situ characterization of histone methylation and acetylation residues revealed that
chromatin states strongly differ in the germ cells of apomictic and sexual plants. In particular,
while the egg cell and early embryo of sexual plants appear mostly quiescent, the staining
patterns in apomicts suggest strong transcriptional activity. The extent and role of early
transcription during seed development (which in apomicts will be necessarily maternal) is still
a controversial topic. Several evidences in sexual plants, including expression data (such as
maternal expression of marker genes in seeds during the first few days after fertilization) or
genetic approach, point to a strong maternal control (Grossniklaus et al., 1998; Kinoshita et
al., 1999; Luo et al., 2000; Moore, 2002; Guitton and Berger, 2005; Pagnussat et al., 2005; Ng
et al., 2007; Vielle-Calzada et al., 2000). These conclusions, together with the characterization
of epigenetic histone marks associated with transcriptional competence reported here, provide
a dynamic image regarding genome-wide transcriptional activation during development. We
observed that the developing maize embryo at 4 DAP undergoes a massive increase in
transcriptional activity after a long transcriptional quiescent state in mature egg cell and early
embryogenesis. Interestingly, the central cell and endosperm showed a markedly different
transcriptional profile with sustained activity before and after fertilization. Based on previous
works indicating a predominance of maternal transcripts in the early endosperm (Vielle-
92

Calzada et al., 2000; Grimanelli et al., 2005; Li and Dickinson, 2010) we assume that this
early activity is mostly, if not strictly, maternal.
In mammals, transcriptional quiescence is thought to be important for the establishment and
maintenance of cell fate in the germ lines, by preventing somatic differentiation (Seydoux and
Braun, 2006). When transcription is not repressed, germ cells do not form, due to the
transformation of their precursors to a somatic fate (Schaner and Kelly, 2006). Transcriptional
repression may also be key to establish transcriptional profiles compatible with totipotency
(Seydoux and Braun, 2006). In plants, somatic cells switch programs late during development
to produce reproductive cells. Furthermore, the products of meiosis suffer additional
reprograming, which strongly differs from the gametes in animals, by undergoing additional
cell divisions and differentiation steps to form a multicellular gametophyte containing the
gametes. Despite these differences, some data indicates that animal and plant germ cell
development share mechanistic similarities as illustrated by the importance of Argonaute
proteins members in both kingdoms (Nonomura et al., 2007). Nevertheless, the role in cell
fate determination of the relative transcriptional quiescence detected in egg and early embryo
in sexual plants remains to be examined. Interestingly, we observed here that, in contrast to
sexual embryos, parthenogenetic embryos are transcriptionally active before fertilization.
Indeed, we found that loss of cell fate occurs early in the developmental course of unreduced
egg cells. Therefore, whether both continuous transcription is sufficient to induce
parthenogenesis in apomicts or cell fate definition in sexual egg requires transcriptional
quiescence is unclear.
Here, neither DMT102 nor DMT103 loss of function produces fully functional apomixis,
indicating that it is not sufficient to promote the instalment of the trait in a sexual background.
The similar behaviour of these two genes in two natural apomictic systems, however, supports
the hypothesis that maintenance and/or establishment of DNA methylation might be one of
the mechanisms involved in the switch from sexual to apomictic reproduction. Moreover, we
propose that the major effect of transcriptional repression as a regulator of germ cell
specification and development via a RdDM-like pathway in maize, might be directed to non-
CG sequence sites.
93

This study provides the first evidence that CMEs are likely crucial to apomictic development.
Our analysis remains however extremely preliminary. While the proteins identified here have
clear homologues in Arabidopsis, it is unclear whether they actually define a RdDM, or a
totally different pathway. A detailed characterization of this pathway is consequently required
to determine its role in apomixis, and will constitute an essential follow-up to this project. A
more exhaustive analysis of the complete set of down and up regulated CMEs may provide
additional interesting insights. From that perspective, a crucial test must be performed in an
apomictic background, by overexpressing the downregulated CMEs; this will indicate
whether their down-regulation is indeed necessary for apomixis expression. Such experiments
have been difficult in the past, due to the lack of appropriate promoters to drive gene
expression in reproductive tissues. This work, however, describes a number of extremely
interesting expression profiles that will be used to derive promoter for expression studies.
The key remaining question, clearly, is to define the target genes of this RdDM-like pathway.
How many of these targets genes are involve in the apomictic features? Are their expression
cause or consequence of apomixis?, Can they be used in a future engineer of artificial
apomixis in crops? To answer these questions is therefore the next challenge to evaluate the
full impact of chromatin dynamics in the regulation of epigenome activity and its efficacy in
apomixis.
94

ACKNOWLEDGMENTS
I would like to express my gratitude to the members of APOMIXIS group at IRD for giving
me the opportunity to develop my initial research proposal, and for all the relevant
suggestions to improve my final work.
To:
Daniel Grimanelli, for his patience and unconditional support,
Olivier Leblanc, for all the interesting discussions that lead me toward new questions,
Daphné Autran, for being a nice friend and a very good colleague.
Jerôme, Shalem, Manjit, Marion and Caroline, for created such a nice and collaborative
atmosphere in the laboratory.
Special thanks to the members of my thesis committee: Thomas Dresselhaus, Raphäel
Mercier, Jean-Marc Deragon, Celia Baroux and M. Alain Ghesquiere for dedicating their time
to evaluate my work.
Thank you so much to Olivier Hamant, Celia Baroux and Thierry Lagrange for the excellent
advice during the thesis committee meetings.
I would like to thank Stéphane Jouannic, Therry Joët, and Martine Devic for their helpful
comments and suggestions in the article preparation.
I am also grateful to T Dresselhaus, S Kaeppler, M Sachs, the Maize Chromatin Consortium
and the Maize TILLING project members for sharing materials.
In the same way thank you so much to Martha Hernandez and Pablo Alva for helping me in
the collect of Tripsacum samples. Special thanks to Rigoberto Vicencio Perez for his
encouragement and support.
I am also grateful to the Apomixis Consortium grant from Pionner Hi Bred, Syngenta Seeds,
Group Limagrain and IRD.
Finally, I would like to thank CONACyT (the Mexican National Council for Science and
Technology) for essential economical support.

AGRADECIMIENTOS
Al pueblo de México quien a través del CONACyT hizo posible mis estudios de doctorado en
Francia.
Al Dr. Roberto Miranda Medrano, por recordarme y motivarme a insistir en el logro de mis
sueños y proyectos.
A mi Madre, el soporte firme de mi maravillosa familia, por su confianza, por creer en mi, por
apoyarme, y por comprenderme.
A Rigoberto Vicencio Pérez, Berenice García Ponce de León, Rosalinda Tapia, Dra. Elena
Alvarez-Buylla, y todos los miembros del laboratorio de genética molecular del desarrollo y
evolución de Plantas, IE UNAM, quienes me brindaron un enorme apoyo, moral, económico
y emocional durante los momentos difíciles de este proceso.
Al Dr. Eulogio Pimienta, por su permanente motivación a mi superación profesional.
A Ana Maria Soriano, Leticia Díaz, Martina López y Lucila Méndez cuyo apoyo y
comprensión me sostuvo durante el mas difícil periodo de mi vida. Este logro, es gracias a
Uds. chicas!
A Rocío Guzmán Najera, por todas esas bonitas muestras de amistad incondicional.
A mi poeta, por tu cariño, por tu ternura y por completar mi vida.
A todos mis amiguitos en Francia: Fabienne, Mauricio, Andrea, Harling, Pedro, Deless,
Gustave, Frederic, Marco, Amalia, Mila, Philippe, Julie, Martine, Thom, Edwin, Alexandra,
Helen, Antoin, Laurent and Laurence... por los buenos momentos.

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