Experimental Analysis of the Fertilization Process
Koen Weteringsa,1 and Scott D. Russellb,1
a Department of Experimental Botany, University of Nijmegen, 6525 ED Nijmegen, The Netherlands
b Department of Botany and Microbiology, University of Oklahoma, Norman, Oklahoma 73019
INTRODUCTION
In flowering plants, double fertilization is one of the defining
features of reproductive development (Raghavan, 2003). Double
fertilization was first discovered in 1898 by Nawaschin. It involves
a complex series of interactions between essentially three
plants--the male gametophyte (MG), the female gametophyte
(FG), and the sporophyte--culminating in the fusion of sexual
cells and nuclei and the formation of an embryo and endosperm
through separate, presumably parallel, fusion and activation
steps. In this review, we focus on recent advances toward
understanding the mechanisms that control maturational synchrony,
interaction of the MG with the FG, delivery of the male
gametes, movement of sperm cells to the female gametes, fusion
of the gametes, and cytological changes before and after
fertilization.
The mature MG or pollen is composed of three cells: one
vegetative cell that encloses two sperm cells or male gametes.
The vegetative cell coordinates the delivery of the two male
gametes to the FG. After landing on the stigma of an appropriate
flower, a cascade of events leads to the establishment of polarity
in the MG and the formation of a pollen tube. Interactions
between the sporophytic tissues of the stigma, the style, and the
MG guide the tube--with the two sperm cells inside--to the
ovary. Within the ovary, the tube grows toward a receptive ovule,
enters the ovule through the micropyle, and delivers the sperm
cells within the FG to effect double fertilization. The genetic
programs that govern the development of the MG and its
interactions with the sporophyte are reviewed elsewhere in
this issue (Edlund et al., 2004; McCormick, 2004; Kao and
Tsukamoto, 2004; Sanchez et al., 2004).
The FG, or embryo sac, constitutes the structural setting for
double fertilization. The FG is embedded in several diploid cell
layers of the ovule and typically consists of seven cells: three
antipodal cells at the chalazal end; a large central cell--precursor
of the endosperm; at the micropylar end, two synergids
presumed to govern the chemical attraction and delivery
mechanism; and the egg cell--precursor of the zygote. Other
variants on this pattern also occur (Huang and Russell, 1992).
Together, the egg and synergids form the egg apparatus.
Genetic programs that control the development of the FG are
reviewed elsewhere in this issue (Yadegari and Drews, 2004).
Upon arrival at and penetration into the micropyle, the pollen
tube delivers its two sperm cells into one of the two synergids by
entering near and sometimes penetrating the filiform apparatus
of the receptive synergid and subsequently rupturing. Inside the
synergid, a mixture of cytoplasm of the two former cells (i.e., the
tube and synergid) forms the milieu in which the nonmotile sperm
cells are conveyed to the egg and central cell, where they fuse
and start the development of the zygote and the endosperm,
respectively. Despite a large body of knowledge concerning the
events surrounding fertilization (reviewed recently by Drews and
Yadegari, 2002; Lord and Russell, 2002; Willemse and van
Lammeren, 2002; Higashiyama et al., 2003; Raghavan, 2003),
the cellular mechanisms that control these events are still poorly
understood. However, in recent years, the rediscovery of Torenia
as a model system, improvements in the experimental manipulation
of male and female gametes, advances in microscope
techniques, and new molecular and genetic approaches have
provided the means to obtain deeper knowledge of the underlying
control mechanisms that govern the identity and behavior
of the major participants in the fertilization process.
IN VIVO STUDIES OF FERTILIZATION
IN FLOWERING PLANTS
Cell Cycle and Fertilization
For fertilization to be successful, the cell cycles of the male and
female gametes must be synchronized. Almost all animals
undergo gamete fusion in the G1-phase of the cell cycle (with
a 1C complement of genomic DNA). Seed plants appear to be
an exception in that they fuse in either G1 or G2 in different species.
Additionally, pollen may be disseminated in one of two
conditions, also depending on plant species. Pollen may be shed
in a bicellular condition (containing the vegetative cell and the
generative cell; the latter is an immediate precursor to the sperm
cells) or in a tricellular condition (containing the vegetative cell
and the two sperm cells). In each condition, male germ cells may
be in G1-, G2-, or S-phase at dissemination (Friedman, 1999). In
the case of bicellular pollen, sperm cells are formed within the
pollen tube. Because gamete maturation occurs during pollen
tube elongation in the style and ovary, long-distance signaling
likely occurs such that the male and female gametes achieve
synchrony at the time of fusion. In addition to long-distance
signaling, there probably also is a signal indicating that sperm
and female target cells are at the same receptive cell cycle
phase. Maize (Zea mays) and other members of the grass family
tend to fuse in G1, but others may be disseminated in S-phase
1 To whom correspondence should be addressed. E-mail koenw@sci.
kun.nl; fax: 31-24-3652490 or e-mail srussell@ou.edu; fax: 1-405-325-
7619.
Article, publication date, and citation information can be found at
www.plantcell.org/cgi/doi/10.1105/tpc.016873.
The Plant Cell, Vol. 16, S107­S118, Supplement 2004, www.plantcell.org  2004 American Society of Plant Biologists
and complete DNA synthesis in the pollen tube to fuse in G2, in
synchrony with the female target cells (Friedman, 1999). In
tobacco (Nicotiana tabacum), bicellular pollen is shed with the
generative cell possessing a 2C complement of DNA (G2). At 8 to
10 h after pollination, the generative cell divides and the resulting
sperm cell nuclei remain in a 1C condition (G1) as they approach
the ovary. Within the degenerated synergid, the sperm cells
complete the S-phase and appear to fuse only when they enter
G2 (H.Q. Tian, T. Yuan, and S.D. Russell, unpublished data).
Long-Distance Cues Promoting Gamete
Maturation and Synchrony
Because plant sexual reproduction is based on the interplay of
cells from a variety of different sources, it seems likely that
mechanisms are present to ensure synchrony in receptivity as
a prelude to successful gamete delivery and fusion. A clear
example of long-distance maturational signaling is provided by
the orchid Phalaenopsis. In this species, ovules do not start to
develop until the stigma is pollinated, and in the absence of
pollination, the flower never develops ovules. The progamic
phase takes 80 days, and the FG is fully mature when the pollen
tube arrives. Auxin and the ethylene precursor 1-aminocyclopropane-1-carboxylic
acid can partly mimic this effect, but other
yet unidentified primary pollination factors are needed to induce
the full postpollination syndrome (Zhang and O'Neill, 1993;
O'Neill, 1997). Also, in other species with shorter progamic
phases, evidence for long-distance communication between
gametophytes has been observed. In a number of species,
synergid receptivity and pollination are correlated (Jensen et al.,
1983; Russell, 1992; Christensen et al., 1997; Tian and Russell,
1997a). For instance, in Nicotiana, one of the two synergids
becomes receptive and starts to accumulate more loosely bound
calcium. Such differences in loosely bound calcium originate
between the two synergids within 1 day after their formation, but
they are accelerated in pollinated flowers. In addition, Tian and
Russell (1997a) found that the immature FG reaches the eightnucleate
stage at 1 day after anthesis without pollination or within
12 to 18 h with successful pollination. Interestingly, at 12 to 18 h
after pollination, a 1-aminocyclopropane-1-carboxylic acid
synthase expression peak is present in the lower part of the
pollinated style (Weterings et al., 2002).
Acceleration of other aspects of egg and embryo sac
maturation also is evident in pollinated flowers. For example,
male and female gametes must be synchronized with respect to
the cell cycle to result in a successful fusion product. In Nicotiana,
the egg cell and sperm cells are formed at G1-phase but need to
progress to G2 before fusion (H.Q. Tian, T. Yuan, and S.D.
Russell, unpublished data). The onset of cell cycle progression
(S-phase) in the egg cell occurs in synchrony with the male
gametes if the flower is pollinated near anthesis. If pollination is
withheld, S-phase in the egg cell is completed at 84 h, which is
more than a 1-day delay; thus, pollination may accelerate cell
cycle progression in the egg cell. In maize, maturation of the
female gamete also is accelerated by several hours with the
presence of pollen tubes in the style (Mó l et al., 2000).
In addition, there also may be long-distance signaling with
respect to the restoration of receptivity in aging flowers. If
Nicotiana flowers are pollinated as late as 5 days after anthesis,
ultrastructural changes relating to senescence may be reversed,
loosely bound calcium distributions will be restored, and some
seed set will occur (Tian and Russell, 1997a). The nature of the
synchronizing signals exchanged between the pollen tube and
the FG is not yet understood.
Short-distance signaling also seems to occur between male
and female gametes, because gametic fusion is delayed in
Arabidopsis thaliana (Friedman, 1999) and tobacco (see Cell
Cycle and Fertilization above) until both reach G2. However, such
short-distance signaling is suspected to involve specific cellular
epitopes, whereas long-distance signaling presumably involves
a diffusible stimulus.
Pollen Tube Attraction
Preceding fertilization, the pollen tube needs to be guided from
the stigma to the embryo sac embedded in the ovule. Previously,
it was suggested that the pollen tube may be guided by only
a single chemical cue emanating from the ovule (Mascarenhas,
1993). However, models have indicated that this distance is far
too long to be governed by a single gradient and may require
several consecutive cues (Lush, 1999). In the transmitting tissue
and along the ovarian wall, the growth of the pollen tube appears
to be governed by extracellular matrix components and
arabinogalactan proteins, respectively (Lennon et al., 1998;
Johnson and Preuss, 2002; Sanchez et al., 2004). Once the
pollen tube has emerged within the ovary, the pollen tube
diverges from the septum tract and starts growing toward the
ovule. Upon reaching the ovule, the tube grows up the stalk of the
funiculus, typically elongating parallel to the files of cells until it
nears the micropyle, where it diverges from this path and enters
the ovule through the micropyle (Figure 1A) (Hu¨ lskamp et al.,
1995; Shimizu and Okada, 2000). Both sporophytic and gametophytic
tissues seem to be involved in attracting the pollen tube
from the septum to the ovary.
Recently, g-aminobutyric acid (GABA) was found to be one of
the possible attractants that is produced by sporophytic tissues.
In Arabidopsis, Palanivelu et al. (2003) found a sharp increase in
the concentration of GABA between the septum and the inner
integument cells near the micropyle of the ovule. In the gynoecium
of pollen-pistil incompatibility2 (pop2) mutants of Arabidopsis,
GABA levels are very high as a consequence of reduced
GABA transaminase expression. However, these plants still have
a steep septum-to-integument GABA gradient. Pollen tubes
from pop2 plants grow at normal speed through the stylar
transmitting tract of both mutant and wild-type plants, but only in
the mutant ovary do they show aberrant growth; there, they fail to
adhere to the ovule and do not enter the micropyle. Wild-type
pollen tubes grow normally in pop2 ovaries (Wilhelmi and Preuss,
1996). Palanivelu et al. (2003) showed that pop2 pollen tubes are
hypersensitive to GABA and suggest that they are unable to grow
toward the high-GABA sources in the pop2 ovary. However, the
pollen tube­attracting activity of GABA could not be reproduced
in vitro, suggesting that, together with GABA, additional
molecules are needed.
Using various gametophytic and sporophytic mutants,
Hu¨ lskamp et al. (1995) and Ray et al. (1997) showed that pollen
S108 The Plant Cell
tubes are not attracted to incompletely formed ovules or those
lacking embryo sacs. These findings demonstrate that the mature,
fully formed FG is necessary to guide the pollen tube from
the placental tissue to the ovule, the distance of attraction not
exceeding 200 mm. Additionally, in the recently described
female gametophytic magamata mutants, the pollen tubes grow
normally along the funiculus but lose their way when they are
within 100 mm of the micropyle (Figure 1B) (Shimizu and Okada,
2000). Together, these data suggest that guidance by the FG can
be subdivided into funicular and micropylar guidance.
A new opportunity to dissect the origin and nature of the FG
guidance signal was provided in the model plant Torenia
fournieri. In this plant, part of the embryo sac protrudes through
the micropyle, and the egg and synergids may be manipulated
directly (Erdelská , 1974; Higashiyama et al., 1997). Isolated
mature ovules can be cultured in vitro and attract pollen tubes
from a distance of 100 to 200 mm (Figure 2). Pollen tubes from
Vandellia angustifolia do not respond to this attraction signal.
Remarkably, pollen tubes can grow toward the ovules only when
they have grown through the style, and this ability to capacitate
the pollen tube is independent of the species of the style
(Higashiyama et al., 1998, 2001, 2003; T. Higashiyama, unpublished
data). Laser ablation of individual or various combinations
of egg, synergid, and central cells showed that the
attraction of the embryo sac is lost only when both synergids are
ablated. Together, these finds show that (1) the synergids are the
source of a pollen tube attraction signal, and (2) the competence
of pollen tubes to respond to this directional signal requires
growth in gynoecial tissues. In addition, it shows that in Torenia
the attraction works directly via a species-distinct, diffusible
signal rather than indirectly via changes induced in the extracellular
matrix.
The identity of the synergid-derived attractant is not yet
known. Calcium has been shown to accumulate in the synergids
upon embryo sac maturation (Russell, 1996; Tian and Russell,
1997a; Kristó f et al., 1999), and pollen tubes of some species
show a tropism toward calcium gradients (Reger et al., 1992a).
However, this simple and general signaling molecule does not
seem to be universally active in all plants. In Torenia, for example,
the presence of 2 mM Ca21 in the medium does not disturb
pollen tube attraction (Higashiyama et al., 2001). Reliance upon
Ca21 alone as a signal does not explain why pollen tubes of
different Torenia species fail to find ovules of other Torenia
species (Higashiyama, 2002). Therefore, although calcium as
a synergid-derived signal might attract pollen tubes, other
molecules, such as sugars or peptides, also could play a role
(Reger et al., 1992b; Russell et al., 1996; Palanivelu and Preuss,
2000; Willemse and van Lammeren, 2002). The role of Ca21 as
a second messenger in signal transduction mechanisms also
may provide amplification of more specific attraction systems.
Although numerous pollen tubes elongate in the stigma and
style of multiovulate flowers, the density of pollen tubes
decreases as they enter the ovary and typically reaches a 1:1
ratio as pollen tubes reach the proximity of the ovule. The
tendency of only one pollen tube to penetrate the ovule appears
to be an important mechanism for reducing polyspermy and
preventing heterofertilization (fertilization of the egg and central
cells by sperm cells of different pollen tubes). The reason that
only one pollen tube typically enters an ovule could be the result
of (1) loss of the pollen tube attractive signal (Higashiyama et al.,
2001) or (2) active pollen tube repulsion (Shimizu and Okada,
2000). Cessation of the attractive signal upon pollen tube arrival
is believed to be relatively rapid. Although multiple pollen tubes
elongating on the funiculus often do so on opposite sides, this
fact does not resolve whether they use up an attractive or nutrient
signal or produce a repulsive signal. In addition, Huck et al. (2003)
found that multiple tubes can enter embryo sacs in Arabidopsis
mutant feronia (fer) FGs, which argues against active pollen tube
repulsion. Instead, the authors argue in favor of a rapid loss of the
pollen tube attractive signal, which prevents multiple penetrations.
It was also noted in fer FGs that the activity of other
synergid genes is unchanged in the remaining synergid after
Figure 1. Scanning Electron Micrographs of Pollen Tubes Growing
toward the Micropyle of Arabidopsis Ovules.
(A) Self-pollination of the wild-type ovule. The pollen tube elongates
almost straight along the funiculus, then turns toward the micropyle.
(B) Self-pollination of the magamata FG mutant ovule. The pollen tube
elongates almost precisely axially along the funiculus but loses its way
near the micropyle.
Pollen tubes are indicated with arrowheads. f, funiculus; mi, micropyle.
Bars  20 mm. Adapted from Shimizu and Okada (2000) and reproduced
with the permission of The Company of Biologists, Ltd.
Figure 2. Attraction of the Pollen Tube to the Naked Embryo Sac of
a Torenia fournieri Ovule in Vitro.
Images are from continuous interference contrast microscope video
observations. The time after the start of video recording (from 10 h after
the start of cultivation) is indicated at top right of each image. The
growing tube always remains directed toward the micropylar end of the
embryo sac, suggesting that the tube is attracted to this region. mi,
micropylar end of the embryo sac; ptt, pollen tube tip. Bar  30 mm.
Adapted from Higashiyama et al. (2001) and reproduced with the
permission of the American Association for the Advancement of Science.
Experimental Analysis of Fertilization S109
pollen tube entry. Apparently, in the fer FG, the capacity to
produce attractant is not lost immediately, which would lead to
the attraction of multiple pollen tubes. More data are required
to confirm this idea and to explain why multiple pollen tubes
enter the same ovule so infrequently.
Although a number of pollen-expressed receptors have been
identified (Muschietti et al., 1998; Kim et al., 2002), it is not yet
clear which are involved in perceiving the directional cues. It is
becoming clear, however, that a family of Rho small GTPases
(Rop) may be involved in the ensuing change of growth direction
(Yang, 2002). Studies of both pollen tube and root hair formation
and growth have shown that Rop is involved in establishing and
maintaining polar growth. At its localization site, Rop regulates
a tip-focused cytosolic Ca21 gradient, promoting and regulating
the formation and dynamics of tip-localized F-actin (Li et al.,
1999; Fu et al., 2001; Molendijk et al., 2001). Both are important
factors for polar tip growth (Hepler et al., 2001; Moutinho et al.,
2001). How Rop is localized to the future site of germination or tip
growth is not yet known, but results from studies of root hair
formation suggest that Rop is recruited to the future site of pollen
tube germination by ADP-ribosylation-factor­GTPase-dependent,
actin-independent vesicle trafficking (Molendijk et al.,
2001). Together, these studies suggest that in response to
directional cues from the ovule and synergid, Rop localization
and activation are reoriented in the direction of this signal,
changing the direction of pollen tube growth toward the micropyle
(Zheng and Yang, 2000). Whether this model is true, and
if it is, how the signal is perceived and processed to reorient
Rop, remains to be determined. Recently, it was shown that the
Actin related proteins 2 and 3 (Arp2/3) complex controls polar
cell expansion of trichomes and leaf pavement cells by regulating
the assembly of diffuse cortical F-actin. It is not yet clear whether
Arp2/3 also controls pollen tube growth and reorientation--
which are similarly dependent on cortical F-actin--because
mutations in various genes that code for Arp2/3 complex proteins
in Arabidopsis have not shown any effect on pollen tube growth
(Le et al., 2003; Li et al., 2003; Mathur et al., 2003a, 2003b).
Nuclear Migration and Cellular Communication in the
Embryo Sac
Nuclear migration within the central cell precedes fertilization. In
Torenia and Arabidopsis, the polar nucleus is located originally in
a midchalazal position, but it moves to the center of the embryo
sac and closely approaches the egg apparatus at the time of
pollen tube arrival and cellular fusion (Christensen et al., 1997;
Higashiyama et al., 1997; Faure et al., 2002). In unpollinated
Torenia, the movement of the fused polar nuclei to the fertilization
site also occurs but is delayed for 24 h compared with pollinated
flowers. After the sperm nucleus enters the central cell, it adheres
to the polar fusion nucleus, and, consistent with prior microcinematographic
observations (Erdelská , 1974), the sperm
nucleolus enlarges to the size of the polar nucleoli and fuses
during migration. The fused nucleus migrates to a far chalazal
position, where division of the primary endosperm nucleus
occurs at 15 h after pollination (Higashiyama et al., 1997;
Wallwork and Sedgley, 2000). By contrast, in Arabidopsis, the
fused nuclei do not migrate. Soon after pollen tube arrival, the
egg nucleus instead moves away from the egg­central cell
boundary (Christensen et al., 1997; Faure et al., 2002).
Upon maturation and fertilization, symplastic cell-to-cell
communication appears to become more restrictive. During
maturation, rhodamine-labeled dextran of 10 kD passes freely
between the central cell and egg apparatus cells. At fertilization,
the size exclusion limit of the cell-to-cell communication is
decreased to 3 kD. If pollination is withheld, the exclusion barrier
also decreases to 3 kD, but this occurs 2 days after anthesis (Han
et al., 2000). In either case, barriers are established that may
decrease communication and therefore increase the autonomy
of the fertilized egg before embryo development.
Synergid Degeneration and Gamete Delivery
Pollen tubes enter the ovule through the micropyle, traverse the
nucellus (if present), and elongate within the ovule until they
encounter the synergids within the embryo sac. The tube enters
near the filiform apparatus, elongates into the cytoplasm of the
synergid, and discharges, usually forming an aperture at the tip
(Russell, 1992). In Arabidopsis, it has been determined that
pollen tube discharge occurs within 2 to 3 min after entry into
the synergid (Rotman et al., 2003, online video). The receptive
synergid usually degenerates, and this occurs (1) before
pollination, (2) as a response to pollination, or (3) upon the entry
of the pollen tube (Russell, 1992, 1996). The degenerating
synergid appears to assist fertilization by reducing resistance
during pollen tube entry (Higashiyama et al., 2000, online video).
The other, persistent synergid retains its turgor and thus may
avoid entry. In addition, the degenerating synergid reorganizes
its cytoskeleton to aid gamete migration (see Male Gamete
Migration within the Embryo Sac below). Also, as discussed
in Pollen Tube Attraction above, in a number of species the
degenerating synergid accumulates Ca21 (Russell, 1996; Tian
and Russell, 1997a; Kristó f et al., 1999) that might act as a pollen
tube attractant (Reger et al., 1992a) or might help in establishing
a favorable microenvironment for gamete fusion (Antoine et al.,
2000; Faure, 2001). Little is known about the mechanism that
regulates synergid cell death. Some synergid-specific marker
genes have been identified (Cordts et al., 2001; Huck et al.,
2003), but none of them specifically labels the degenerating
synergid.
In the recently described FG mutant gametophytic factor2
in Arabidopsis, the synergids do not degenerate. The gene has
been cloned and encodes a DnaJ chaperonin that is expressed
throughout the plant and is localized to the mitochondria. This
indicates that synergid cell death probably depends on some
involvement of mitochondrial function (Christensen et al., 2002).
In addition, this mutant and the sire`ne (srn) mutant (in which
the synergids also remain intact) still are capable of attracting
pollen tubes (Rotman et al., 2003), indicating that--at least in
Arabidopsis--synergid cell death is not required for pollen tube
attraction. In fact, experimental ablation studies suggest that
only intact synergids attract pollen tubes to the ovule, underscoring
the importance of both synergids in pollen tube receipt:
the persistent synergid for attraction and the other, typically
degenerated receptive synergid for entry of the pollen tube
(Higashiyama et al., 2001).
S110 The Plant Cell
In Torenia and Arabidopsis, it has been shown that soon after
entry in the embryo sac, the pollen tube discharges. The pollen
tube contents then flow into the space previously occupied by
the receptive synergid (Higashiyama et al., 2000; Rotman et al.,
2003, online videos) and thus into the narrow intercellular space
located between the chalazal end of the egg cell and the
micropylar end of the central cell. This area has been suggested
to be the region where the sperm cells fuse with the egg and
central cell (Russell, 1992).
Some conflicting data exist concerning the duration that
sperm cells reside in the synergid in Torenia. Higashiyama et al.
(1997) estimated that they are present only briefly, whereas
Wallwork and Sedgley (2000) report that the sperm cells remain
in the synergid for more than 1 h. For only a short period after
gamete delivery are four nuclei detectable in the receptive
synergid, representing the nuclei of the synergid, the vegetative
cell, and the two sperm cells (Higashiyama et al., 1997).
Presumably, soon after pollen tube discharge, the DNA in the
exposed synergid and vegetative nuclei breaks down in the
nucleases of the embryo sac (Heslop-Harrison et al., 1999).
Female Control of Fertilization
Although male and female gametes ultimately may share equally
in their genetic contributions, there is accumulating evidence
that the FG may regulate the delivery of the male gametes. In the
Arabidopsis FG mutants fer and srn, embryo sac development
is not affected and synergid differentiation and specification
appear to be normal. However, after the pollen tube has entered
the embryo sac, the tip of the pollen tube appears to either
proliferate or grow the length of the central cell and abort, failing
to discharge and deposit its sperm cells into the synergid
(Figure 3) (Huck et al., 2003; Rotman et al., 2003). Similar
overgrowth of the pollen tube within the embryo sac is observed
after incongruous interspecific crosses in Rhododendron
(Williams et al., 1986). Together, these data suggest that the
arrest of pollen tube growth and rupture are controlled directly
or indirectly by a compound from the synergid. Currently, the
nature of this compound is unknown, but further analysis of
the FER and SRN genes, as well as other genes that disrupt
FG function without otherwise affecting their development
(Christensen et al., 2002; Drews and Yadegari, 2002), will help
in dissecting the mechanisms that regulate gamete delivery. To
date, no evidence has been found for the existence of a specific
male mechanism that controls pollen tube arrest or discharge.
Gametic fusion also may be conditioned by the female. For
example, in barley, the male cytoplasm is excluded from the egg,
as is evident from the cytoplasmic sheath that remains within the
degenerated synergid at the site of fusion with the egg. Exclusion
of the sperm cytoplasm, however, does not occur during fusion
with the central cell (Mogensen, 1988). Whether any inheritance
of paternal plastids occurs in the offspring is determined largely
by the quantity of plastids in the sperm-lineage cells, which often
are excluded or eliminated progressively during development
(Nagata et al., 1999). In an unusual case in alfalfa (Medicago
sativa), there are lineages in which paternal plastids are characteristically
abundant or sparse. However, the abundance of
paternal plastids in the offspring of these plants is inherited
only through the maternal lineage. Pollen with either sparse or
abundant plastids had no predictive value in controlling the
abundance of paternal plastids in the offspring. The maternal
genotype predicted the outcome of the cross (Shi et al., 1991;
Zhu et al., 1991).
Preferential transmission of hyperploid maize sperm cells
based on B-chromosome abundance is similarly conditioned by
the female (Chiavarino et al., 1998) (see Preferential Fertilization
below). The ratio of B-chromosomes in the progeny is influenced
strongly by the female plant. Plants displaying either high or low
transmission of B-chromosomes are correlated with the maternal
plant rather than the paternal plant. High or low transmission
is conditioned by the maternal plant, indicating that the maternal
parent controls fusion rates.
Interestingly, mate choice in plants appears to occur with similar
consequences as in animals; female choice conditions the
mate decisions made, whereas the male triggers this process.
Male Gamete Migration within the Embryo Sac
The complex pollen tube cytoskeletal arrays that maintain the
subapical position of male gametes during tube elongation are
depolymerized rapidly after pollen tube discharge, thus releasing
the sperm cells from the tube, from their physical association,
and from the enclosing membranes of the pollen. Presumably,
this is when the cotransmission of the sperm cells ends and their
developmental fates diverge. Two actin ``coronas'' are formed
from the middle of the degenerated synergid, one terminating
near the egg nucleus and the other ending near the central cell.
The actin coronas, together with the myosin that is acquired on
the surface of the sperm cells (Zhang et al., 1999; Zhang and
Russell, 1999), appear to cooperate in the actomyosin-mediated
transport of one sperm cell along the actin corona pathway to
a position opposite the egg nucleus and the other sperm cell to
a position near the central cell nucleus (Huang and Russell, 1994;
Huang and Sheridan, 1998; Huang et al., 1999; Fu et al., 2000;
Figure 3. Female Control of Fertilization in Arabidopsis.
Fluorescence microscope images of aniline blue­labeled ovules penetrated
by pollen tubes.
(A) Wild-type ovule. The pollen tube terminates in the micropylar area of
the embryo sac.
(B) feronia ovule. The pollen tube enters the central cell after coiling in the
micropylar area.
pt, pollen tube; mi, micropylar area. Bars  30 mm. Adapted from Huck
et al. (2003) and reproduced with the permission of The Company of
Biologists, Ltd.
Experimental Analysis of Fertilization S111
Ye et al., 2002). Cellular fusions at these sites transmit the sperm
nuclei to their respective target cells.
Microinjected Alexa 488­phalloidin in the living embryo sac
of Torenia reveals a dramatic reorganization of F-actin in the
synergid and egg cells during pollen tube approach, arrival, and
discharge. An actin cap located at the base of the synergid
accumulates in patches along the cortex of the egg cell and
develops into a linear corona track. When the sperm cells arrive
within the embryo sac, they appear to follow this track to the
female gametes. Soon after fusion, the actin corona disappears
(Fu et al., 2000). A critical role for actin coronas during fertilization
also can be concluded from a study of the fertilization of
supernumerary eggs in the maize FG mutant indeterminate
gametophyte1. Only fertilized eggs are associated with actin
coronas; unfertilized supernumerary egg cells lack an actin
corona (Huang and Sheridan, 1998). In addition, in the egg and
central cells of nun orchids (Phaius tankervilliae), cellular
indentations complementary to the sperm cell shape have been
noted as target sites near the termination of coronas (Ye et al.,
2002). Despite the important role for actin coronas in fertilization,
it is not yet known how the reorganization of actin microfilaments
into linear corona tracks is regulated. A template for the coronas
is present before pollen tube entry (Fu et al., 2000), but most of
the corona appears to assemble after the pollen tube discharges.
The completion of the corona at a time when the synergid is
highly degenerated suggests the incorporation of pollen tube
actin. Likewise, free myosin may associate with sperm cells to
form an actomyosin transport mechanism (Zhang and Russell,
1999).
Preferential Fertilization
Preferential fertilization--in which one sperm cell has a greater
likelihood of fusing with the egg than the other--may theoretically
occur in any of the plants in which the sperm cells differ, but it
may not be visible without a specific cellular marker. The data
in Table 1 suggest that sperm cells are frequently dissimilar. In
Plumbago zeylanica, fusion between the egg and the plastid-rich
sperm cell occurs in >95% of the cases examined (Russell, 1985,
and subsequent unpublished observations), selectively transmitting
male plastids into the zygote.
Preferential fertilization also occurs in maize, in which the
sperm cell containing one extra set of B-chromosomes fuses
with the egg up to 75% of the time (Roman, 1948; Carlson, 1969,
1986), but the result is interestingly variable. Rather than simply
providing a marker for the sperm cell destined to fuse with the
egg, B-chromosomes appear to confer a selective advantage to
the sperm cell that contains them (Carlson, 1969). Not every
division of the generative cell results in sperm cells with
aneuploidy. In lines of maize in which B-chromosomes occur,
the products of division vary, and fusion results differ (Carlson,
1969; Rusche et al., 1997, 2001). Preferential fertilization in maize
is important as a populational phenomenon, but each experiment
seems to result in differing degrees of sperm preferentiality
(Roman, 1948; Carlson, 1969, 1986; Faure et al., 2003), and
demonstrating preferentiality in individual fusions has not been
possible. For example, it has not been possible to obtain
preferential fertilization using the maize in vitro fertilization
system (Faure et al., 2003). Nevertheless, preferential fertilization
remains an intrinsically interesting phenomenon of flowering
plants that could provide considerable insight into the control of
gamete maturation and identity.
IN VITRO STUDIES OF FERTILIZATION
IN FLOWERING PLANTS
In the late 1980s, technical advances in isolating living, viable
gametes led to the in vitro combination of male and female
gametes. Initially, electrofusion (Kranz et al., 1991) and chemical
induction studies (Kranz and Lo¨ rz, 1994) were introduced as
reliable means of producing fusion, but a calcium-based fusion
model that seems largely to mimic in vivo fertilization has
attracted considerable interest (Faure et al., 1994). However,
plant regeneration has not been achieved using direct products
of the calcium-based fusion method. Regeneration of fertile
plants and in vitro formation of endosperm have been achieved
from electrofused gametes (Kranz and Lo¨ rz, 1993; Kranz et al.,
1998). Electrofusions of gametes of the grasses Coix, Sorghum,
Hordeum, and Triticum also have been achieved (reviewed by
Kranz and Kumlehn, 1999). Attempts to combine dicot gametes
chemically have met with some success in Nicotiana tabacum
(Sun et al., 1995, 2000, 2001; Tian and Russell, 1997b), but there
have been no successes at regenerating plants.
The in vitro fusion model that most closely mimics in vivo fusion
uses isolated maize gametes (Faure et al., 1994) in a medium
containing Ca21 at a concentration of 5 mM, which appears to
facilitate but not to coerce fusion. Fusion of female gametic cells
is most efficient with sperm cells compared with other cells.
Polyspermy is reportedly blocked successfully, and chemical
signals, including a calcium influx, have been recorded (see
Calcium Waves, Site of Fusion, and Egg Activation below). The
model seems to work best on slightly immature egg cells (Mó l
et al., 2000) and is a bit technically demanding because
receptivity may decline quickly after isolation. However, this
model system does not seem to predict all types of gametic
behavior. For example, Faure et al. (2003) reported recently that
they were unable to mimic gamete preference, although in the
same article they confirmed that the same gametes underwent
B-chromosome­based preferential fertilization in vivo in the plant
cultivar that they studied.
Some insightful in vitro fertilization experiments performed on
tobacco sperm cells include comparing the fusion efficiency of
somatic cells with that of a series of cells of different diameters
and conducting multiple fusions. Objects of the same size class
as sperm cells provided the greatest fusion efficiency among
somatic cells (Sun et al., 2001). Sun et al. (1995, 2000) also tested
the ability of the egg to accept multiple fusions. They found that
after the first fusion, a fusion product is inhibited from fusing with
another cell or organelle for a refractory period of at least 30 min.
This inhibition of fusion applies to various combinations of
gametes, somatic cells, and gametic-somatic combinations.
Until the fused cells complete cytoplasmic reorganization,
further fusion appears to be inhibited (Sun et al., 1995). This
would prevent rapid serial fusions with more than one cell in the
medium at a specific time. Potentially, this could inhibit
S112 The Plant Cell
polyspermy as well. That such a nonspecific mechanism may
inhibit multiple fusions suggests a need for further studies to
corroborate claims that there is a specific plant mechanism to
inhibit polyspermy.
Obtaining distant hybrids and genetically modified artificial
zygotes has been a frequently cited goal of in vitro fertilization, so
it is interesting that transgenes in gametes and in in vitro­fertilized
zygotes have now been achieved (Scholten and Kranz, 2001).
The use of promoters specific to the germline (e.g., the promoter
of the generative cell-specific gene lilium generative cell 1
(LGC1), which is activated in the generative and sperm cells
[Singh et al., 2003]) could alter gene expression in the germline to
achieve novel expression in seeds or in the next generation.
Sperm Maturation
The competence of flowering plant gametes to respond appropriately
to gametes of the opposite sex is acquired after
gamete formation and likely involves modification of the gametic
cell surface. In tobacco, for example, immediately after their
formation, sperm cells placed in apposition often fuse spontaneously
(Tian and Russell, 1998). In vivo, a rapidly deposited
callose wall forms between the sperm cells, separating them and
impeding such indiscriminate fusions (Yu and Russell, 1993).
Within minutes of formation, however, sperm cells lose this
tendency, and at 20 h after sperm formation, fusion can be induced
only by modifying the cell surface. Exposure of the cells
to dilute solutions containing both cellulase and pectinase was
sufficient to trigger fusion in tobacco (Tian and Russell, 1998).
Presumably, sperm maturation involves multiple carbohydrate
determinants on the cell surface that reduce the likelihood of
random fusions but that do not impede sexual gamete fusion.
Such surface changes are anticipated to include compounds
such as arabinogalactan proteins (Southworth and Kwiatkowski,
1996) and other potentially immunogenic compounds
(Southworth et al., 1999). In Plumbago, the polarity of the male
germ unit is determined by the position of the vegetative
nucleus and appears to be determined in the generative cell
(Russell et al., 1996).
Table 1. Summary of Heteromorphism in Flowering Plant Sperm Cells Examined to Date
Pollen Type and Family Species Method Morphology Sample Size Reference
Tricellular pollen
Monocotyledons
Poaceae Hordeum vulgaris 3-D I 5 Mogensen and Rusche, 1985
Secale cereale 3-D I 4 Mogensen and Rusche, 2000
Zea mays 3-D SD 1 McConchie et al., 1987a
3-D SD 1 Rusche, 1988
3-D SD 1 Rusche and Mogensen, 1988
Dicotyledons
Carifoliaceae Abelia spathulata DAPI OD  Saito et al., 2002
Brassicaceae Brassica campestris 3-D MD 5 McConchie et al., 1987b
Brassica oleracea 3-D MD 3 McConchie et al., 1987b
Euphorbiaceae Euphorbia dulcis 3-D MD 1 Murgia and Wilms, 1988
Asteraceae Gerbera jamesonii 3-D MD 1 Provoost et al., 1988
Plumbaginaceae Limonium sinuatum DAPI OD  Saito et al., 2002
Plumbago auriculata DAPI OD  Saito et al., 2001
Plumbago zeylanica 3-D MPD 11 Russell, 1984
Chenopodiaceae Spinacia oleracea 3-D MD 7 Wilms, 1986
Bicellular pollen
Monocotyledons
Iridaceae Gladiolus gandavensis ImAn SD 6 Shivanna et al., 1988
Dicotyledons
Bignoniaceae Campsis grandiflora DAPI OD  Saito et al., 2002
Tecoma capensis DAPI OD  Saito et al., 2002
Ericaceae Rhododendron spp ImAn SD 7 Shivanna et al., 1988
Fabaceae Erythrina crista-galli DAPI OD  Saito et al., 2000
Erythrina variegata DAPI OD  Saito et al., 2002
Solanaceae Nicotiana tabacum 3-D I 9 Yu et al., 1992
(to 30 h after pollination) 3-D I 19 Yu and Russell, 1994
(near pollen tube discharge) ImAn SD 25 Tian et al., 2001
Petunia hybrida 3-D I 1 Wagner and Mogensen, 1988
Methods are as follows: 3-D, serial ultrathin sectioning and three-dimensional analysis; DAPI, fluorochromatic localization on sections embedded in
Technovitt; ImAn, image analysis of whole-cell profiles and/or nuclear fluorochromatic patterns. Morphology types are as follows: I, isomorphic; MD,
mitochondrial cytoplasmic dimorphism; MPD, mitochondrial and plastid cytoplasmic dimorphism; OD, organelle dimorphism in which DNA nucleoids
were visualized but could represent plastids, mitochondria, or both; SD, structural dimorphism (condition of cytoplasmic organellar DNA unknown).
Sample size represents the number of male sperm pairs examined, unless unavailable ().
Experimental Analysis of Fertilization S113
Egg Maturation
For female gametes, the best understood model is maize, in
which egg maturation is categorized into three classes (Mó l et al.,
2000). Small, densely cytoplasmic egg cells with a centrally
positioned nucleus indicating a nonpolarized condition are
classified as type A. The next stage is formed by larger egg cells
that have numerous small vacuoles in the central perinuclear
cytoplasm, designated type B. Mature egg cells, classified as
type C, are large and highly polarized, with a prominent apical
vacuole and a chalazally placed nucleus with perinuclear cytoplasm.
As the silks emerge from their husks, type A egg cells are
prevalent in the ovules (88%), but this decreases to nearly half
(58%) at optimum silk length for pollination. Types B and C
become more common near the time of fertilization.
In the brown alga Fucus and in many animal systems, the site
of sperm entry forms the focal point of a postfertilization calcium
influx, and electrical stimulation may determine zygote polarity.
In animals, pricking the zygote can displace such a focal point
and alter zygotic polarity. In Fucus, light also may reorient polarity
(Goodner and Quatrano, 1993). In angiosperms, there is no such
evidence of plasticity, nor is there any particular region on the
female gamete that seems to be more receptive to fusion than
any other region, at least according to in vitro studies (Antoine
et al., 2001a). In vivo, however, position within the embryo sac
appears to be an important factor in establishing both cellular
identity and polarity. This polarity and orientation seem to remain
unchanged during later development.
Calcium Waves, Site of Fusion, and Egg Activation
In all animal systems studied to date, gamete fusion leads to,
among other events, an increase in cytoplasmic calcium
([Ca21]cyt). This increase in [Ca21]cyt is believed to be essential
to egg activation and occurs at the very onset of embryo
development (Stricker, 1999). In maize, combining the use of
a calcium-specific vibrating probe and the single-wavelength
calcium fluorochrome Fluo-3, Antoine et al. (2001b) demonstrated
that gamete fusion always triggers a calcium influx, which
is then followed by an increase in cytoplasmic calcium (Figures
4A and 4B). The site of the calcium influx is initiated at the fusion
site and within a few minutes moves in a wave-like pattern over
the rest of the membrane, resulting in a homogeneous influx
(Antoine et al., 2000). Thus, the initial pattern of polarity that is
created at the fusion site is no longer evident after the first several
minutes. In vitro, the first visible sign of egg activation is egg
contraction. This seems to coincide with the increase in calcium
influx and is followed by a later smoothing of the cell surface and
a turgid appearance in the cytoplasm (Figure 4C). Within 20 min
after fusion, the cell wall begins to form, as detected using
Calcofluor white (Figure 4D).
When the Ca21 channels are inhibited by Gd31, the sperm cell
still can fuse with the egg cell but the sperm cell cytoplasm is not
incorporated into the egg cell. Despite the blocked Ca21 influx,
however, the cytoplasmic calcium increase still occurs, presumably
because of calcium release from intracellular stores.
Together with the cytoplasmic calcium increase, egg contraction
and cell wall deposition are observed in Gd31-treated cells
(Antoine et al., 2001b), indicating that calcium influx is not
needed to create an increased [Ca21]cyt. The observed calcium
influx instead may be needed for sperm incorporation and
subsequent karyogamy. These observations allow flowering
plants to be included in the general model of multicellular
Figure 4. Calcium Flux, Cytoplasmic Calcium Levels, and Egg Activation
during Maize in Vitro Fertilization.
(A) Graph representing Ca21 influx (blue line) and changes in cytoplasmic
Ca21 levels ([Ca21]cyt; red line). Ca21 influx was measured using
a calcium-specific vibrating probe, and changes in [Ca21]cyt were
measured using the calcium fluorochrome Fluo-3 in combination with
fluorescence microscopy. Time 0 was set at gamete adherence. Gamete
fusion (first vertical line) is followed by the onset of a Ca21 influx and by
a transient increase in [Ca21]cyt. The synchrony of the onset of the
[Ca21]cyt increase with the stabilization of the Ca21 influx (single arrow)
and the synchrony of the [Ca21]cyt peak and the Ca21 influx peak (double
arrow) coincide in this experiment. BPL, basal prefertilization level.
DF/F0  (Ft  F0)/F0, where Ft is fluorescence of the egg cell or zygote as
time  t and F0 is fluorescence of time  0.
(B) Sequence of fluorescence microscope images from which the
changes in [Ca21]cyt were calculated. Changes in [Ca21]cyt inside the egg
cell are reflected by changes in the color spectrum, indicated by the
vertical bar at right. Fusion (Fu) and the time after fusion are indicated at
bottom left. In the first image, the adhered sperm (arrow) is visible
because of Fluo-3 diffusion.
(C) Bright-field microscope images of different cytological events during
maize in vitro fertilization. (a) Gamete adhesion (arrowhead shows the
sperm). (b) Fusion. The sperm can be seen penetrating the egg cell. (c)
Mild contraction of the egg cell. (d) Reshaping of the cell. Bar  20 mm.
(D) Fluorescence microscope image of a 2-h zygote stained with
Calcofluor white to detect cell wall deposition. Bar  20 mm.
Adapted from Antoine et al. (2001b) and reproduced with the permission
of the Nature Publishing Group.
S114 The Plant Cell
organisms in which increased cytoplasmic calcium is necessary
and sufficient to induce egg activation.
Han et al. (2002) studied whether the increase in cytoplasmic
calcium level induced during fertilization is caused by the sperm
fusion event or by a factor present in the sperm cytoplasm. They
injected T. fournieri sperm cell extract mixed with calcium green
dextran into the central cell and within minutes measured
increased [Ca21]cyt that lasted for >40 min. Injection of maize
sperm extract into T. fournieri did not have the same effect,
indicating that the sperm cell factors may vary in different
phylogenetic groups. Injection and activation of caged inositol
1,4,5-triphosphate showed that the signaling cascade triggered
by the sperm factor(s) depends on inositol 1,4,5-triphosphate
for calcium release. These experiments were performed in the
central cell because, as was found in the maize in vitro system, it
was impossible to inject the egg cell (Antoine et al., 2001b; Y. Han
and B.Q. Huang, unpublished data). Although it remains to be
determined whether sperm cell factors are the main cause of
cytoplasmic calcium increase in the egg cell, this seems to be
supported by the fact that sperm extract from flowering plants
can cause calcium oscillations in mouse eggs (Li et al., 2001).
CONCLUSIONS AND PROSPECTS
Fertilization in flowering plants is a very subtle and accurate
process. Compared with animals, in which hundreds of millions
of male gametes may target one, often enormous, female
gamete, in plants the two nonmotile male gametes target two
female gametes with micrometer-level precision. Clearly, fertilization
is a complex process that we are gradually beginning
to understand. In recent years, we have seen advances in
understanding the mechanisms that govern the interactions
between the MG and FG and their gametes at cytological,
physiological, biochemical, and molecular genetic levels. Using
the Torenia system and FG mutants, we are probably very close
to identifying the synergid-derived pollen tube attractants and
compounds that regulate gamete delivery. Surely, studies of
fertilization seem poised for more significant advances as both
molecular understanding and experimental systems improve.
The unique characteristics are currently being explored by
genome- and transcriptome-level examination (McCormick,
2004), and powerful, high-throughput means exist to characterize
the metabolic status of MG and FG cells. The incorporation of
experimental methods, including microdissection, ablation, and
fluorescence bleaching using lasers, in addition to increasingly
precise genetic and molecular labels, represents an opportunity
to understand gamete biology. Given the high number of genes
found in the genomes of Arabidopsis and rice compared with
mammals, including humans, it would by surprising if gametophytic
biology does not occupy a significant role in the genetic
complexity of plants.
ACKNOWLEDGMENTS
We thank T. Higashiyama for allowing us to use data before publication
and T. Higashiyama and V. Raghavan for critically reading the
manuscript. K.W. is supported by Grant 811.36.003 from the Netherlands
Organization for Scientific Research. Work by S.D.R. was
supported by U.S. Department of Agriculture National Research
Initiative Competitive Grants Program Grant 99-37304-8097 and the
University of Oklahoma.
Received September 2, 2003; accepted January 12, 2004.
REFERENCES
Antoine, A.F., Dumas, C., Faure, J.E., Feijó , J.A., and Rougier, M.
(2001a). Egg activation in flowering plants. Sex. Plant Reprod. 14,
21­26.
Antoine, A.F., Faure, J.E., Cordeiro, S., Dumas, C., Rougier, M., and
Feijó , J.A. (2000). A calcium influx is triggered and propagates in the
zygote as a wavefront during in vitro fertilization of flowering plants.
Proc. Natl. Acad. Sci. USA 97, 10643­10648.
Antoine, A.F., Faure, J.E., Dumas, C., and Feijó , J.A. (2001b).
Differential contribution of cytoplasmic Ca21 and Ca21 influx to
gamete fusion and egg activation in maize. Nat. Cell Biol. 3, 1120­
1123.
Carlson, W.R. (1969). Factors affecting preferential fertilization in maize.
Genetics 62, 543­554.
Carlson, W.R. (1986). The B chromosome of maize. CRC Crit. Rev.
Plant Sci. 3, 201­226.
Chiavarino, A.M., Rosato, M., Rosi, P., Poggio, L., and Naranjo, C.A.
(1998). Localization of the genes controlling B chromosome transmission
rate in maize (Zea mays ssp. mays, Poaceae). Am. J. Bot. 85,
1581­1585.
Christensen, C.A., Gorsich, S.W., Brown, R.H., Jones, L.G., Brown,
J., Shaw, J.M., and Drews, G.N. (2002). Mitochondrial GFA2 is
required for synergid cell death in Arabidopsis. Plant Cell 14, 2215­
2232.
Christensen, C.A., King, E.J., Jordan, J.R., and Drews, G.N. (1997).
Megagametogenesis in Arabidopsis wild type and the Gf mutant. Sex.
Plant Reprod. 10, 49­64.
Cordts, S., Bantin, J., Wittich, P.E., Kranz, E., Lo¨ rz, H., and
Dresselhaus, T. (2001). ZmES genes encode peptides with structural
homology to defensins and are specifically expressed in the female
gametophyte of maize. Plant J. 25, 103­114.
Drews, G.N., and Yadegari, R. (2002). Development and function of the
angiosperm female gametophyte. Annu. Rev. Genet. 36, 99­124.
Edlund, A.F., Swanson, R., and Preuss, D. (2004). Pollen and stigma
structure and function: The role of diversity in pollination. Plant Cell 16
(suppl.), S84­S97.
Erdelská , O. (1974). Contribution to the study of fertilization in the living
embryo sac. In Fertilization in Higher Plants, H.F. Linskens, ed
(Amsterdam: North Holland Publishing Company), pp. 191­195.
Faure, J.E. (2001). Double fertilization in flowering plants: Discovery,
study methods and mechanisms. C. R. Acad. Sci. III 324, 551­558.
Faure, J.E., Digonnet, C., and Dumas, C. (1994). An in vitro system for
adhesion and fusion of maize gametes. Science 263, 1598­1600.
Faure, J.E., Rotman, N., Fortune, P., and Dumas, C. (2002).
Fertilization in Arabidopsis thaliana wild type: Developmental stages
and time course. Plant J. 30, 481­488.
Faure, J.-E., Rusche, M.L., Thomas, A., Keim, P., Dumas, C.,
Mogensen, H.L., Rougier, M., and Chaboud, A. (2003). Double
fertilization in maize: The two male gametes from a pollen grain have
the ability to fuse with egg cells. Plant J. 33, 1051­1062.
Friedman, W.E. (1999). Expression of the cell cycle in sperm of
Arabidopsis: Implications for understanding patterns of gametogenesis
and fertilization in plants and other eukaryotes. Development 126,
1065­1075.
Experimental Analysis of Fertilization S115
Fu, Y., Wu, G., and Yang, Z. (2001). Rop GTPase-dependent dynamics
of tip-localized F-actin controls tip growth in pollen tubes. J. Cell Biol.
152, 1019­1032.
Fu, Y., Yuan, M., Huang, B.Q., Yang, H.Y., Zee, S.Y., and O'Brien,
T.P. (2000). Changes in actin organization in the living egg apparatus
of Torenia fournieri during fertilization. Sex. Plant Reprod. 12,
315­322.
Goodner, B., and Quatrano, R. (1993). Fucus embryogenesis: A model
to study the establishment of polarity. Plant Cell 5, 1471­1481.
Han, Y.Z., Huang, B.Q., Guo, F.L., Zee, S.Y., and Gu, H.K. (2002).
Sperm extract and inositol 1,4,5-triphosphate induce cytosolic
calcium rise in the central cell of Torenia fournieri. Sex. Plant Reprod.
15, 187­193.
Han, Y.Z., Huang, B.Q., Zee, S.Y., and Yuan, M. (2000). Symplastic
communication between the central cell and the egg apparatus cells
in the embryo sac of Torenia fournieri Lind. before and during
fertilization. Planta 211, 158­162.
Hepler, P.K., Vidali, L., and Cheung, A.Y. (2001). Polarized cell growth
in higher plants. Annu. Rev. Cell Dev. Biol. 17, 159­187.
Heslop-Harrison, J., Heslop-Harrison, J.S., and Heslop-Harrison, Y.
(1999). The structure and prophylactic role of the angiosperm embryo
sac and its associated tissues: Zea mays as a model. Protoplasma
209, 256­272.
Higashiyama, T. (2002). The synergid cell: Attractor and acceptor of the
pollen tube for double fertilization. J. Plant Res. 115, 149­160.
Higashiyama, T., Kuroiwa, H., Kawano, S., and Kuroiwa, T. (1997).
Kinetics of double fertilization in Torenia fournieri based on direct
observations of the naked embryo sac. Planta 203, 101­110.
Higashiyama, T., Kuroiwa, H., Kawano, S., and Kuroiwa, T. (1998).
Guidance in vitro of the pollen tube to the naked embryo sac of
Torenia fournieri. Plant Cell 10, 2019­2031.
Higashiyama, T., Kuroiwa, H., Kawano, S., and Kuroiwa, T. (2000).
Explosive discharge of pollen tube contents in Torenia fournieri. Plant
Physiol. 122, 11­13.
Higashiyama, T., Kuroiwa, H., and Kuroiwa, T. (2003). Pollen-tube
guidance: Beacons from the female gametophyte. Curr. Opin. Plant
Biol. 6, 36­41.
Higashiyama, T., Yabe, S., Sasaki, N., Nishimura, Y., Miyagishima,
S.-Y., Kuroiwa, H., and Kuroiwa, T. (2001). Pollen tube attraction by
the synergid cell. Science 293, 1480­1483.
Huang, B.Q., Fu, Y., Zee, S.Y., and Hepler, P.K. (1999). Threedimensional
organization and dynamic changes of the actin cytoskeleton
in embryo sacs of Zea mays and Torenia fournieri. Protoplasma
209, 105­119.
Huang, B.-Q., and Russell, S.D. (1992). Female germ unit: Organization,
isolation and function. Int. Rev. Cytol. 140, 233­296.
Huang, B.Q., and Russell, S.D. (1994). Fertilization in Nicotiana
tabacum: Cytoskeletal modifications in the embryo sac during
synergid degeneration. A hypothesis for short-distance transport of
sperm cells prior to gamete fusion. Planta 194, 200­214.
Huang, B.Q., and Sheridan, W.F. (1998). Actin coronas in normal and
indeterminate gametophyte1 embryo sacs of maize. Sex. Plant
Reprod. 11, 257­264.
Huck, N., Moore, J.M., Federer, M., and Grossniklaus, U. (2003). The
Arabidopsis mutant feronia disrupts the female gametophytic control
of pollen tube reception. Development 130, 2149­2159.
Hu¨ lskamp, M., Schneitz, K., and Pruitt, R.E. (1995). Genetic evidence
for a long-range activity that directs pollen tube guidance in
Arabidopsis. Plant Cell 7, 57­64.
Jensen, W.A., Ashton, M.E., and Beasley, C.A. (1983). Pollen tubeembryo
sac interaction in cotton. In Pollen: Biology and Implications
for Plant Breeding, D.L. Mulcahy and E. Ottaviano, eds (New York:
Elsevier Science Publishing), pp. 67­72.
Johnson, M.A., and Preuss, D. (2002). Plotting a course: Multiple
signals guide pollen tubes to their targets. Dev. Cell 2, 273­281.
Kao, T.-H., and Tsukamoto, T. (2004). The molecular and genetic
bases of S-RNase-based self incompatibility. Plant Cell 16 (suppl.),
S72­S83.
Kim, H.U., Cotter, R., Johnson, S., Senda, M., Dodds, P.,
Kulikauskas, R., Tang, W.H., Ezcurra, I., Herzmark, P., and
McCormick, S. (2002). New pollen-specific receptor kinases identified
in tomato, maize and Arabidopsis: The tomato kinases show
overlapping but distinct localization patterns on pollen tubes. Plant
Mol. Biol. 50, 1­16.
Kranz, E., Bautor, J., and Lo¨ rz, H. (1991). In vitro fertilization of single,
isolated gametes of maize mediated by electrofusion. Sex. Plant
Reprod. 4, 12­16.
Kranz, E., and Kumlehn, J. (1999). Angiosperm fertilisation, embryo
and endosperm development in vitro. Plant Sci. 142, 183­197.
Kranz, E., and Lo¨ rz, H. (1993). In vitro fertilization with isolated, single
gametes results in zygotic embryogenesis and fertile maize plants.
Plant Cell 5, 739­746.
Kranz, E., and Lo¨ rz, H. (1994). In vitro fertilisation of maize by single
egg and sperm cell protoplast fusion mediated by high calcium and
high pH. Zygote 2, 125­128.
Kranz, E., von Wiegen, P., Quader, H., and Lo¨ rz, H. (1998).
Endosperm development after fusion of isolated, single maize sperm
and central cells in vitro. Plant Cell 10, 511­524.
Kristó f, Z., Timar, O., and Imre, K. (1999). Changes of calcium
distribution in ovules of Torenia fournieri during pollination and
fertilization. Protoplasma 208, 149­155.
Le, J., El-Assal, S.E.-D., Basu, D., Saad, M.E., and Szymanski, D.B.
(2003). Requirements for Arabidopsis ATARP2 and ATARP3 during
epidermal development. Curr. Biol. 13, 1341­1347.
Lennon, K.A., Roy, S., Hepler, P.K., and Lord, E.M. (1998). The
structure of the transmitting tissue of Arabidopsis thaliana (L.) and the
path of pollen tube growth. Sex. Plant Reprod. 11, 49­59.
Li, H., Lin, Y., Heath, R.M., Zhu, M.X., and Yang, Z. (1999). Control of
pollen tube tip growth by a Rop GTPase-dependent pathway that
leads to tip-localized calcium influx. Plant Cell 11, 1731­1742.
Li, S., Blanchoin, L., Yang, Z., and Lord, E.M. (2003). The putative
Arabidopsis Arp2/3 complex controls leaf cell morphogenesis. Plant
Physiol. 132, 2034­2044.
Li, S.T., Huang, X.Y., and Sun, F.Z. (2001). Flowering plant sperm contains
a cytosolic soluble protein factor which can trigger calcium oscillations
in mouse eggs. Biochem. Biophys. Res. Commun. 287, 56­59.
Lord, E.M., and Russell, S.D. (2002). The mechanisms of pollination
and fertilization in plants. Annu. Rev. Cell Dev. Biol. 18, 81­105.
Lush, W.M. (1999). Whither chemotropism and pollen tube guidance?
Trends Plant Sci. 4, 413­418.
Mascarenhas, J.P. (1993). Molecular mechanisms of pollen tube
growth and differentiation. Plant Cell 5, 1303­1314.
Mathur, J., Mathur, N., Kernebeck, B., and Hu¨ lskamp, M. (2003a).
Mutations in actin-related proteins 2 and 3 affect cell shape
development in Arabidopsis. Plant Cell 15, 1632­1645.
Mathur, J., Mathur, N., Kirik, V., Kernebeck, B., Srinivas, B.P., and
Hu¨ lskamp, M. (2003b). Arabidopsis CROOKED encodes for the
smallest subunit of the ARP2/3 complex and controls cell shape by
region specific fine F-actin formation. Development 130, 3137­3146.
McConchie, C.A., Hough, T., and Knox, R.B. (1987a). Ultrastructural
analysis of the sperm cells of mature pollen of maize, Zea mays.
Protoplasma 139, 9­19.
McConchie, C.A., Russell, S.D., Dumas, C., Touhy, M., and Knox,
R.B. (1987b). Quantitative cytology of the mature sperm of Brassica
campestris and B. oleracea. Planta 170, 446­452.
S116 The Plant Cell
McCormick, S. (2004). Control of male gametophyte development.
Plant Cell 16 (suppl.), S142­S153.
Mogensen, H.L. (1988). Exclusion of male mitochondria and plastids
during syngamy in barley as a basis for maternal inheritance. Proc.
Natl. Acad. Sci. USA 85, 2594­2597.
Mogensen, H.L., and Rusche, M.L. (1985). Quantitative ultrastructural
analysis of barley sperm. I. Occurrence and mechanism of cytoplasm
and organelle reduction and the question of sperm dimorphism.
Protoplasma 128, 1­14.
Mogensen, H.L., and Rusche, M.L. (2000). Occurrence of plastids in
rye (Poaceae) sperm cells. Am. J. Bot. 87, 1189­1192.
Mó l, R., Idzikowska, K., Dumas, C., and Matthys-Rochon, E. (2000).
Late steps of egg cell differentiation are accelerated by pollination in
Zea mays L. Planta 210, 749­757.
Molendijk, A.J., Bischoff, F., Rajendrakumar, C.S.V., Friml, J.,
Braun, M., Gilroy, S., and Palme, K. (2001). Arabidopsis thaliana
Rop GTPases are localized to tips of root hairs and control polar
growth. EMBO J. 20, 2779­2788.
Moutinho, A., Hussey, P.J., Trewavas, A.J., and Malhó , R. (2001).
cAMP acts as a second messenger in pollen tube growth and
reorientation. Proc. Natl. Acad. Sci. USA 98, 10481­10486.
Murgia, M., and Wilms, H.J. (1988). Three-dimensional image and
mitochondrial distribution in sperm cells of Euphorbia dulcis. In Plant
Sperm Cells as Tools for Biotechnology, H.J. Wilms and C.J. Keijzer,
eds (Wageningen, The Netherlands: Centre for Agricultural Publishing
and Documentation), pp. 75­80.
Muschietti, J., Eyal, Y., and McCormick, S. (1998). Pollen tube
localization implies a role in pollen-pistil interactions for the tomato
receptor-like protein kinases LePRK1 and LePRK2. Plant Cell 10,
319­330.
Nagata, N., Saito, C., Sakai, A., Kuroiwa, H., and Kuroiwa, T. (1999).
The selective increase or decrease of organellar DNA in generative
cells just after pollen mitosis one controls cytoplasmic inheritance.
Planta 209, 53­65.
O'Neill, S.D. (1997). Pollination regulation of flower development. Annu.
Rev. Plant Physiol. Plant Mol. Biol. 48, 547­574.
Palanivelu, R., Brass, L., Edlund, A.F., and Preuss, D. (2003). Pollen
tube growth and guidance is regulated by POP2, an Arabidopsis gene
that controls GABA levels. Cell 114, 47­59.
Palanivelu, R., and Preuss, D. (2000). Pollen tube targeting and axon
guidance: Parallels in tip growth mechanisms. Trends Cell Biol. 10,
517­524.
Provoost, E., Southworth, D., and Knox, R.B. (1988). Threedimensional
reconstruction of sperm cells and vegetative nucleus
in pollen of Gerbera jamesonii. In Plant Sperm Cells as Tools for
Biotechnology, H.J. Wilms and C.J. Keijzer, eds (Wageningen, The
Netherlands: Centre for Agricultural Publishing and Documentation),
pp. 69­74.
Raghavan, V. (2003). Some reflections on double fertilization, from its
discovery to the present. New Phytol. 159, 565­583.
Ray, S., Park, S., and Ray, A. (1997). Pollen tube guidance by the
female gametophyte. Development 124, 2489­2498.
Reger, B.J., Chaubal, R., and Pressey, R. (1992a). Chemotropic
responses by pearl millet pollen tubes. Sex. Plant Reprod. 5, 47­56.
Reger, B.J., Pressey, R., and Chaubal, R. (1992b). In vitro chemotropism
of pearl millet pollen tubes to stigma tissue: A response to
glucose produced in the medium by tissue-bound invertase. Sex.
Plant Reprod. 5, 201­205.
Roman, H. (1948). Directed fertilization in maize. Proc. Natl. Acad. Sci.
USA 34, 36­42.
Rotman, N., Rozier, F., Boavida, L., Dumas, C., Berger, F., and
Faure, J.-E. (2003). Female control of male gamete delivery during
fertilization in Arabidopsis thaliana. Curr. Biol. 13, 432­436.
Rusche, M.L. (1988). Three-dimensional reconstruction of Zea mays
in the mature pollen grain. In Plant Sperm Cells as Tools for
Biotechnology, H.J. Wilms and C.J. Keijzer, eds (Wageningen, The
Netherlands: Centre for Agricultural Publishing and Documentation),
pp. 61­68.
Rusche, M.L., and Mogensen, H.L. (1988). The male germ unit of Zea
mays: Quantitative ultrastructure and three-dimensional analysis.
In Sexual Reproduction in Higher Plants, M. Cresti, P. Gori, and
E. Pacini, eds (Heidelberg, Germany: Springer-Verlag), pp. 221­226.
Rusche, M.L., Mogensen, H.L., Chaboud, A., Faure, J.E., Rougier,
M., Keim, P., and Dumas, C. (2001). B chromosomes of maize (Zea
mays L.) are positioned nonrandomly within sperm nuclei. Sex. Plant
Reprod. 13, 231­234.
Rusche, M.L., Mogensen, H.L., Shi, L., Keim, P., Rougier, M.,
Chaboud, A., and Dumas, C. (1997). B chromosome behavior in
maize pollen as determined by a molecular probe. Genetics 147,
1915­1921.
Russell, S.D. (1984). Ultrastructure of the sperms of Plumbago
zeylanica. 2. Quantitative cytology and three-dimensional organization.
Planta 162, 385­391.
Russell, S.D. (1985). Preferential fertilization in Plumbago: Ultrastructural
evidence for gamete-level recognition in an angiosperm. Proc.
Natl. Acad. Sci. USA 82, 6129­6133.
Russell, S.D. (1992). Double fertilization. Int. Rev. Cytol. 140, 357­388.
Russell, S.D. (1996). Attraction and transport of male gametes for
fertilization. Sex. Plant Reprod. 9, 337­342.
Russell, S.D., Strout, G.W., Stramski, A.K., Mislan, T.W., Thompson,
R.A., and Schoemann, L.M. (1996). Microgametogenesis in Plumbago
zeylanica (Plumbaginaceae). 1. Descriptive cytology and threedimensional
organization. Am. J. Bot. 83, 1435­1453.
Saito, C., Nagata, N., Sakai, A., Kuroiwa, H., and Kuroiwa, T. (2001).
Behavior of plastid nucleoids during male gametogenesis in Plumbago
auriculata. Protoplasma 216, 143­154.
Saito, C., Nagata, N., Sakai, A., Mori, K., Kuroiwa, H., and Kuroiwa,
T. (2000). Unequal distribution of DNA-containing organelles in
generative and sperm cells of Erythrina crista-galli (Fabaceae). Sex.
Plant Reprod. 12, 296­301.
Saito, C., Nagata, N., Sakai, A., Mori, K., Kuroiwa, H., and Kuroiwa,
T. (2002). Angiosperm species that produce sperm cell pairs or
generative cells with polarized distribution of DNA-containing organelles.
Sex. Plant Reprod. 15, 167­178.
Sanchez, A.M., Bosch, M., Bots, M., Nieuwland, J., Feron, R., and
Mariani, C. (2004). Pistil factors controlling pollination. Plant Cell 16
(suppl.), S98­S106.
Scholten, S., and Kranz, E. (2001). In vitro fertilization and expression
of transgenes in gametes and zygotes. Sex. Plant Reprod. 14, 35­40.
Shi, L., Zhu, T., Mogensen, H.L., and Smith, S.E. (1991). Paternal
plastid inheritance in alfalfa: Plastic nucleoid number within generative
cells correlates poorly with plastid number and male plastid transmission
strength. Curr. Genet. 19, 399­402.
Shimizu, K.K., and Okada, K. (2000). Attractive and repulsive
interactions between female and male gametophytes in Arabidopsis
pollen tube guidance. Development 127, 4511­4518.
Shivanna, K.R., Xu, H., Taylor, P., and Knox, R.B. (1988). Isolation of
sperms from the pollen tubes of flowering plants during fertilization.
Plant Physiol. 87, 647­650.
Singh, M., Bhalla, P.L., Xu, H., and Singh, M.B. (2003). Isolation and
characterization of a flowering plant male gametic cell-specific
promoter. FEBS Lett. 542, 47­52.
Southworth, D., and Kwiatkowski, S. (1996). Arabinogalactan proteins
at the cell surface of Brassica sperm and Lilium sperm and generative
cells. Sex. Plant Reprod. 9, 269­272.
Experimental Analysis of Fertilization S117
Southworth, D., Kwiatkowski, S., Smith, A.R., Sharpless, H.,
Merwin, J., and Marusich, M.F. (1999). Antibodies to floweringplant
sperm. Protoplasma 208, 115­122.
Stricker, S.A. (1999). Comparative biology of calcium signaling
during fertilization and egg activation in animals. Dev. Biol. 211,
157­176.
Sun, M.X., Moscatelli, A., Yang, H.Y., and Cresti, M. (2000). In vitro
double fertilization in Nicotiana tabacum (L.): Polygamy compared
with selected single pair somatic protoplast and chloroplast fusions.
Sex. Plant Reprod. 13, 113­117.
Sun, M.X., Moscatelli, A., Yang, H.Y., and Cresti, M. (2001). In vitro
double fertilization in Nicotiana tabacum (L.): The role of cell volume in
cell fusion. Sex. Plant Reprod. 13, 225­229.
Sun, M.X., Yang, H.Y., and Zhou, C. (1995). Single-pair fusion of
various combinations between female gametoplasts and other
protoplasts in Nicotiana tabacum. Acta Bot. Sin. 36, 489­493.
Tian, H.Q., and Russell, S.D. (1997a). Calcium distribution in fertilized
and unfertilized ovules and embryo sacs of Nicotiana tabacum L.
Planta 202, 93­105.
Tian, H.Q., and Russell, S.D. (1997b). Micromanipulation of male and
female gametes of Nicotiana tabacum. I. Isolation of gametes. Plant
Cell Rep. 16, 555­560.
Tian, H.Q., and Russell, S.D. (1998). The fusion of sperm cells and the
function of male germ unit (MGU) of tobacco (Nicotiana tabacum L.).
Sex. Plant Reprod. 11, 171­176.
Tian, H.Q., Zhang, Z.J., and Russell, S.D. (2001). Sperm dimorphism in
Nicotiana tabacum L. Sex. Plant Reprod. 14, 123­125.
Wagner, V.T., and Mogensen, H.L. (1988). The male germ unit in
the pollen and pollen tubes of Petunia hybrida: Ultrastructural,
quantitative and three-dimensional features. Protoplasma 143,
101­110.
Wallwork, M.A.B., and Sedgley, M. (2000). Early events in the
penetration of the embryo sac in Torenia fournieri (Lind.). Ann. Bot.
85, 447­454.
Weterings, K., Pezzotti, M., Cornelissen, M., and Mariani, C. (2002).
Dynamic 1-aminocyclopropane-1-carboxylate-synthase and -oxidase
transcript accumulation patterns during pollen tube growth in tobacco
styles. Plant Physiol. 130, 1190­1200.
Wilhelmi, L.K., and Preuss, D. (1996). Self-sterility in Arabidopsis due
to defective pollen tube guidance. Science 274, 1535­1537.
Willemse, M., and van Lammeren, A. (2002). Fertilization, from
attraction to embryo. Biologia (Bratisl.) 57, 13­22.
Williams, E.G., Kaul, V., Rouse, J.L., and Palser, B.F. (1986).
Overgrowth of pollen tubes in embryo sacs of Rhododendron
following interspecific pollinations. Aust. J. Bot. 34, 413­423.
Wilms, H.J. (1986). Dimorphic sperm cells in the pollen grains of
Spinacia. In Biology of Reproduction and Cell Motility in Plants and
Animals, M. Cresti and R. Dallai, eds (Heidelberg, Germany: SpringerVerlag),
pp. 193­198.
Yadegari, R., and Drews, G.N. (2004). Female gametophyte development.
Plant Cell 16, (suppl.), S133­S141.
Yang, Z. (2002). Small GTPases: Versatile signaling switches in plants.
Plant Cell 14 (suppl.), S375­S388.
Ye, X.L., Yeung, E.C., and Zee, S.Y. (2002). Sperm movement during
double fertilization of a flowering plant, Phaius tankervilliae. Planta
215, 60­66.
Yu, H.S., Hu, S.Y., and Russell, S.D. (1992). Sperm cells in pollen tubes of
Nicotiana tabacum L.: Three-dimensional reconstruction, cytoplasmic
diminution and quantitative cytology. Protoplasma 168, 172­183.
Yu, H.S., and Russell, S.D. (1993). Three-dimensional ultrastructure
of generative cell mitosis in the pollen tube of Nicotiana tabacum. Eur.
J. Cell Biol. 61, 338­348.
Yu, H.S., and Russell, S.D. (1994). Populations of plastids and
mitochondria during male reproductive cell maturation in Nicotiana
tabacum L.: A cytological basis for occasional biparental inheritance.
Planta 193, 115­122.
Zhang, X.S., and O'Neill, S.D. (1993). Ovary and gametophyte
development are coordinately regulated by auxin and ethylene
following pollination. Plant Cell 5, 403­418.
Zhang, Z., and Russell, S.D. (1999). Sperm cell surface characteristics
of Plumbago zeylanica L. in relation to transport in the embryo sac.
Planta 208, 539­544.
Zhang, Z., Tian, H.Q., and Russell, S.D. (1999). Localization of myosin
on sperm-cell-associated membranes of tobacco (Nicotiana tabacum
L.). Protoplasma 208, 123­128.
Zheng, Z.L., and Yang, Z.B. (2000). The Rop GTPase switch turns on
polar growth in pollen. Trends Plant Sci. 5, 298­303.
Zhu, T., Mogensen, H.L., and Smith, S.E. (1991). Quantitative cytology
of the alfalfa generative cell and its relation to male plastid inheritance
patterns in three genotypes. Theor. Appl. Genet. 81, 21­26.
S118 The Plant Cell