American Journal of Botany 92(4): 576–583. 2005.
POST-MEIOTIC
CYTOKINESIS AND POLLEN APERTURE
PATTERN ONTOGENY: COMPARISON OF DEVELOPMENT
IN FOUR SPECIES DIFFERING IN APERTURE PATTERN1
ADRIENNE RESSAYRE,2 LEANNE DREYER,3 SARAH TRIKI-TEURTROY,4
ARLETTE FORCHIONI,4 AND SOPHIE NADOT4
UMR de Genetique Vegetale, Ferme du Moulon, 91190 Gif-sur-Yvette, France; 3Botany Department, University of Stellenbosch,
Private Bag X1, Matieland 7602, South Africa; and 4Laboratoire Ecologie Systématique et Evolution, CNRS UMR 8079, Université
Paris-Sud, 91405 Orsay cedex, France
2
Pollen aperture patterns vary widely in angiosperms. An increasing number of studies indicate that aperture pattern ontogeny is
correlated with the way in which cytokinesis that follows male meiosis is completed. The formation of the intersporal callose walls
that isolate the microspores after meiosis was studied in four species with different aperture patterns (two monocots, Phormium tenax
and Asphodelus albus, and two eudicots, Helleborus foetidus and Protea lepidocarpodendron). The way in which post-meiotic cytokinesis is performed differs between all four species, and variation in callose deposition appears to be linked to aperture pattern
definition.
Key words:
aperture pattern ontogeny; callose; cytokinesis; eudicot; meiosis, monocot; pollen.
Angiosperm pollen grains are composed of two or three
cells enclosed within a complex multilayered wall. Apertures
are well-defined areas of the pollen surface where the external
part of the wall, the exine, is reduced or absent. They accommodate variation in pollen volume, the passage of water during
pollen rehydration, and the exit of the pollen tube during pollen germination. Aperture shape, structure, number and distribution constitute the aperture pattern of a pollen grain. Angiosperms are widely diverse in their aperture patterns (Walker
and Doyle, 1975). This diversity is structured in two morphological groups according to the main taxonomic divisions
(Walker and Doyle, 1975; Blackmore and Crane, 1998). Eudicots, also known as the tricolpate clade, usually produce pollen with three equatorial apertures, but species producing pollen with two or four to six equatorial apertures are commonly
observed. Basal angiosperms and monocots usually produce
pollen with a single distal polar aperture (monosulcate pollen)
or a set of morphologies with a few equatorial (or nearly equatorial) apertures.
In angiosperms, microsporogenesis is surprisingly variable
(Sampson, 1975; Longly and Waterkeyn, 1979a, b; Bandhari,
1984; Brown, 1991). Microsporogenesis of species differ in
three ways: (1) in the timing of the nuclear divisions relatively
to the cytoplasmic divisions (cytokinesis can be successive,
simultaneous or intermediate), (2) in the orientation of the
meiotic axes (tetragonal, rhomboidal, tetrahedral, decussate,
and intermediate tetrad types result from such variation), (3)
in the way callose is deposited to form the cleavage walls
during cytokinesis. In 1935, Wodehouse proposed that cytokinesis following meiosis could be involved in aperture pattern
ontogeny. Since then, an increasing number of studies have
linked aperture pattern ontogeny to meiosis (Blackmore and
Crane, 1998; Ressayre et al., 2002; Furness and Rudall, 2004).
Recently, a developmental model based on the cytological
events that occur during meiosis was proposed. The model
suggests that the combination of the different variable elements during meiosis is sufficient to account for the most
widespread patterns, opening the opportunity to study both
proximal and distal causes of such a diversification (Ressayre
et al., 2002).
However, several aspects of the model still need to be investigated and its generality remains to be demonstrated. The
model is based on the hypothesis that for angiosperms species
displaying six or less apertures, the progress in cytokinesis
defines the places where apertures will meet within tetrads.
More precisely, the model predicts that apertures will meet in
the regions where cytokinesis is completed, a hypothesis initially proposed by Wodehouse (1935). In the model, this mechanism is supposed to apply both to polar and nonpolar patterns, polar apertures being additionally defined by the position
of the spindle pole in the second meiosis (Ressayre et al.,
2002). While there is experimental evidence to show that the
definition of polar apertures is determined by the distribution
of the second meiotic poles (Dover, 1972; Sheldon and Dickinson, 1983, 1986), there is no evidence supporting the idea
that the relationships between apertures in polar species are
determined by cytokinesis.
The developmental model permits us to test this hypothesis.
If relationships between apertures are indeed determined by
cytokinesis, convergence in cytokinesis can be expected between polar and nonpolar species displaying the same relationships between apertures within tetrads. Reciprocally, species displaying different distribution of apertures within tetrads
should differ in terms of cytokinesis. To test these aspects of
the model, we studied the pattern of callose deposition in four
selected species (Asphodelus albus Miller; Phormium tenax
J.R. Forst. and G. Forst.; Protea lepicarpodendron (L.) L.;
Helleborus foetidus L.) that, on the one hand, display common
developmental features, including simultaneous cytokinesis
(Fig. 1A), multiplanar tetrads and a two-step production of the
intersporal callose walls, but on the other hand, differ in aperture patterns (Fig. 1B). As a result, the only differences expected would pertain to the direction of callose deposits during
Manuscript received 3 June 2004; revision accepted 9 December 2004.
A. R. received a financial support from the Société de Secours des Amis
des Sciences and the Singer-Polignac Foundation.
2
Author for correspondence (e-mail: [email protected]) phone:
01.69.33.23.359, fax: 01.69.33.23.40.
1
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Fig. 1. Diagrams of simultaneous cytokinesis and aperture pattern within tetrads in angiosperms. (A) Simultaneous cytokinesis. (a) Before meiosis, the
microsporocytes enclose themselves in a thick wall of callose. (b–c) Both nuclear divisions take place within the cytoplasm of the dividing cell leading to four
haploid nuclei. (d) Microspores are separated by the simultaneous production of callose walls and remain within tetrahedral tetrads until they produce a primary
exine wall. (B) Aperture distribution within tetrad in the four studied species. Microspores are shaded in gray, while apertures are indicated in black. (a)
Monosulcate pollen. Pairs of microspores can generally be recognized within a tetrad (microspores of the same color). The apertures are placed orthogonally
to the cleavage walls separating microspores belonging to the same pair. (b) Trichotomosulcate pollen. Each microspore has a single, polar, distal furrow divided
into three branches. The extremities of the branched furrows meet three by three in four positions of the tetrad (Garside’s distribution of apertures). (c) Triporate
pollen. Each microspore has three equatorial pores. The pores are joined three by three in four positions of the tetrad (Garside’s distribution of apertures). (d)
Tricolpate pollen. Each microspore has three equatorial apertures. The apertures are joined in pairs in six regions of the tetrad (Fisher’s distribution of apertures).
Abbreviations: N, nucleus; cy, cytoplasm; ca, callose.
the formation of the intersporal walls. Asphodelus albus produces monosulcate pollen (Huynh, 1976), that is, pollen that
displays a single distal furrow (Fig. 1B-a). Phormium tenax
produces trichotomosulcate pollen, that is, pollen that displays
a single furrow placed at the distal pole of the grain and is
divided into three branches (Rudall et al., 1997). Protea lepicarpodendron produces triporate pollen (Ertdman, 1952) and
H. foetidus L. produces tricolpate pollen (Echlin and Godwin,
1968). The first two species are monocots and belong to the
order Asparagales (families Asphodelaceae and Phormiaceae,
respectively). The last two species are basal eudicots that belong to the Proteaceae and the Ranunculaceae respectively.
The two eudicot species differ in the arrangement of apertures
within their tetrads. The apertures of H. foetidus are joined in
pairs within the tetrads, as is the common condition in eudicots
(Fig. 1Bd), an arrangement known as the Fisher’s arrangement
of apertures (Fisher, 1890). The apertures of P. lepicarpodendron are joined three by three in the tetrads (Fig. 1Bc). This
is known as the Garside’s arrangement of apertures (Garside,
1946). The extremities of the trichotomosulcus of monocots
are recorded in the literature to follow Garside’s arrangement
(Fig. 1Bb; Huynh, 1971). Differences in cytokinesis are expected within both the monocot and eudicot clades as the species differ in aperture distribution within tetrads. In contrast,
P. tenax (monocot) and P. lepicarpodendron (dicot) display
the same relationships of apertures within the tetrads (Garside’s arrangement). As a result, convergence in cytokinesis is
expected in both species.
MATERIAL AND METHODS
Plant material—Fresh flower buds of each species were collected. Material
of A. albus was collected on the campus of the University of Paris-Sud, Orsay,
France. Material of P. tenax was collected at the Museum National d’Histoire
Naturelle, Paris, France. Material of H. foetidus was collected in the wild in
mountains in the south of France (Col du Coq and Col de Porte, Chartreuse,
France), while material of P. lepicarpodendron was collected in the Jan S.
Marais Reserve (Stellenbosch, South Africa).
Staining—Fresh buds were collected and dissected immediately. The sporogenous cells were extracted from one anther and mounted in acetocarmine
to determine the meiotic stage of the bud. When buds were found to be
undergoing meiosis, the remaining anthers were squashed in aniline blue
(modified from Arens, 1949) to observe callose wall formation using epifluorescence and a Zeiss Axiophot microscope with filter set 01 (excitation 345,
emission 425 nm long pass). When buds were at the tetrad stage, some anthers
were mounted in aniline blue to observe callose, while the rest was mounted
in congo red (Stainier et al., 1967) to observe aperture pattern within the
tetrads. Acetocarmine and congo red preparations were observed with a Zeiss
Axiophot microscope (light microscopy). In A. albus, aperture pattern within
tetrads was also observed using epifluorescence and a Zeiss Axiophot microscope with filter set 07 (excitation 495, emission 520 nm long pass).
RESULTS
In all species studied, cleavage wall formation appears to
be a two-step process. First, the cytoplasm of the future microspores is separated by the formation of callosic cell plates.
Second, additional callose deposition takes place on the cell
plates. Cell plate formation was identical within the monocots
and within the eudicots, but differed between the two groups
(clades), while patterns of additional callose deposition differed among all four species.
Cell plate formation—In both monocot species, cytokinesis
began with the formation of small discs of callose in the cy-
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Fig. 2. Fluorescent light micrographs of cell plate formation after staining with aniline blue. (a–d) Asphodelus albus. (e–h) Phormium tenax. (i–l) Protea
lepicarpodendron. (m–p) Helleborus foetidus. (a–h) Centrifugal formation. Cell plates formation begin with the formation of six small discs of callose (a and
e) that expand centrifugally and rapidly fuse in the center of the cell (b and f) before reaching the microsporocyte cell wall (c and g). Small ingrowths of
callose indicate the places where the cleavage walls will join the surrounding wall of the microsporocyte during cell plate formation in A. albus (arrows in Fig.
2b and c). (i–p) Centripetal formation. In P. lepicarpodendron, cell plate formation begins at the border of the cleavage plane (i) and progress centripetally
towards the center of the plane (j–k). In H. foetidus, cleavage wall formation began with loose callose deposits next to the callose wall surrounding the dividing
microsporocytes in four places (m, arrows). Cell plates then expand centripetally toward the middle of the division plane (n–o). In all species, six naked cell
plates can be seen in the tetrahedral tetrads at the end of cell plate formation (d, h, l, and p). The cell plates are identical in all species, except in A. albus, in
which two of the cleavage walls are wider than the other four (this can be seen in d, where the upper wall is large and the two lateral ones are smaller).
toplasm of the dividing cells (Fig. 2a, e). These discs rapidly
joined in the center of the dividing cells, and six cleavage
walls separating the four future microspores became visible
(Fig. 2b–d and f–h). The progression of cell plates continued
toward the surrounding wall of the dividing cell. Cell plate
formation then appeared to progress centrifugally. In A. albus
small ingrowths of callose were also seen during the extension
of the cell plates (Fig. 2b, c). These ingrowths appear to predict the places were cell plates will meet with the outer wall
surrounding the dividing microsporocyte.
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Unlike the case in monocots, cytokinesis in both eudicots
studied began with the formation of cell plates that progressed
inward from the outer wall that surrounds the dividing cells
(Fig. 2i, m, n). In H. foetidus, it began with the formation of
loose deposits of callose against the dividing microsporocyte
wall (arrows, Fig. 2i). Callose deposition is thus centripetal.
Tetrads—In the four species, almost all tetrads are multiplanar. In these multiplanar tetrads, six cleavage walls intersect
in the center of the dividing cells (Fig. 2d, h, l, p). In both
eudicots and in P. tenax, all cleavage walls appeared to be
identical and orientated at 1208 angles relatively to each other
(Fig. 2h, l, p). Two different shapes of cleavage walls were
detected in A. albus. In at least some tetrads of this species,
two of the cleavage walls are large and crossed the other walls
at angles of close to 1808, while the four others are smaller
and crossed the other walls at angles of close to 908 (Fig. 2d).
A few uniplanar tetrads were also observed in all the species
except in P. lepicarpodendron, where all the tetrads are tetrahedral.
Additional callose deposits and aperture pattern—Asphodelus albus—Additional callose deposits are observed near the
callose wall surrounding the developing tetrad, at the intersection of the cleavage walls (Fig. 3a). In multiplanar tetrads, they
are apparently more abundant on the two largest cell plates
and very faint on the four others (Fig. 3b). Callose deposition
progressed from the intersections of the cell plates toward the
middle of the cleavage walls. As a result, in tetrahedral tetrads,
additional callose deposits led to the formation of two patches
of callose on the two largest cleavage walls (Fig. 3b). The
sulcus formed orthogonally to these cleavage walls (Fig. 4a–
f). In tetragonal tetrads, callose deposits were concentrated
near the callose wall surrounding the tetrad at the intersection
of the cleavage walls, leading to the formation of two patches
on both sides of the tetrads (Fig. 3c, d). The sulcus also formed
orthogonally relative to the cleavage walls (Fig. 4g, h).
In all other species, callose deposits appeared to be identical
on all cell plates. In P. tenax (Fig. 3e–j), they progressed from
the middle of the cell plates toward the intersections of the
cell plates. At the end of cytokinesis, the outlines of the microspores can be seen (Fig. 3j). Microspores appeared as
smooth triangular volumes, the extremities of which met three
by three at the intersections of the cell plates (Fig. 3j). As a
result, the callose deposition following cell plate formation
was completed in the four regions of the tetrads corresponding
to the intersections of three cleavage walls (Fig. 3j). The three
branches of the trichotomosulcus met three by three in the
same regions of the tetrads (Fig. 4i). In P. lepicarpodendron,
additional callose was deposited on the cell plates in a similar
way to that described for P. tenax, although callose appears
to be more evenly deposited (Fig. 3m–t). Callose deposition
started in the middle of the cell plates and progressed centrifugally toward the intersection of the cell plates. Additional
callose deposition was completed at the intersections of three
cleavage walls, and apertures formed at these intersection. In
H. foetidus, callose deposits were produced both at the intersections of the cell plates and of the callose wall surrounding
the developing tetrad and in the center of the tetrad (Fig. 3k,
l). Both deposits appeared to progress toward the middle of
the cleavage walls. Apertures formed in pairs at these places.
579
DISCUSSION
Cytoplasmic division following meiosis appears to be as
variable as aperture pattern itself in the four species studied.
As expected if it is involved in aperture pattern ontogeny, cytokinesis takes place in a different way in each of the four
species. In addition, aperture distribution within tetrads appears to correlate with the places where cytokinesis is completed, both in polar and nonpolar species. All the data provided by the comparison among the four species support the
hypothesis that both in polar and nonpolar species, cytokinesis
is involved in aperture pattern ontogeny. Our results thus
strongly support the theoretical model of pollen aperture development proposed by Ressayre et al. (2002).
The present study also contributes new information on two
aspects. First, we show that cell plate formation progresses
differently in monocots and eudicots, although cytokinesis following male meiosis is a two-step process (cell plate formation
followed by additional callose deposition) in both clades (Waterkeyn, 1962; Longly and Waterkeyn, 1979a). Each step is
apparently independent of the other. Second, we found that
within the species where cell plate formation and additional
callose deposits do not progress in the same way, aperture sites
coincide with the last points of callose deposition and not with
the last points of contact between the cytoplasm of the dividing cells as stated by Wodehouse (1935).
Cell plate formation: one way for monocots, another for
eudicots—In both eudicot species, cell plate formation begins
at the edges of the cleavage planes and progresses toward the
center of the dividing cell. Although centripetal progression of
isolated cell plates has, to our knowledge, never before been
recorded, such a progression appears consistent with other reports of cleavage wall formation in core eudicots (Longly and
Waterkeyn, 1979a, b; Brown, 1991; Ressayre et al., 2001). In
these species, cell plate formation is synchronized with additional callose deposits and both progress centripetally. Formation of isolated cell plates has been described in H. foetidus,
although no indication of cell plate progression (centripetal or
centrifugal) was provided (Waterkeyn, 1962). Centrifugal cell
plate formation has been described in two Proteaceae species
(Blackmore and Barnes, 1995). This discrepancy between Proteaceae species is difficult to explain and further studies are
required to investigate whether these species indeed differ in
cell plate formation. If this was confirmed to be the case, the
distribution of both types of cell plate formation should be
determined. Centripetal progression is markedly different from
the known progression of the cell plate formation during plant
mitosis in general (Heese et al., 1998) and from the progression of cell plates during male meiosis in monocots in particular (Longly and Waterkeyn, 1979a; Brown, 1991). During
plant mitosis, cytokinesis is initiated in late anaphase with the
formation of a phragmoplast, a complex array of microtubules
and actin microfilaments. The phragmoplast is composed of
short, overlapping microtubules of opposite polarity formed at
the center of the cell. From here, they progress centripetally
towards the cell walls. Callose vesicles move along the microtubules of the phragmoplast and coalesce to form a cell
plate that expands centifugally along with the phragmoplast.
Cell plate formation during plant meiosis has never been described in such detail (Otegui and Staehelin, 2000), but has
been recorded to progress centrifugally in a similar fashion to
what we observed in the two monocot species in the present
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Fig. 3. Fluorescent light micrographs of additional callose deposits after staining with aniline blue. (a–d) Asphodelus albus. (e–j) Phormium tenax. (k–l)
Helleborus foetidus. (m–t) Protea lepicarpodendron. (a–b) A. albus multiplanar tetrads. Additional callose is deposited on two of the cell plates in A. albus.
Callose accumulates at the intersection of the cell plates close to the callose wall surrounding the tetrad (arrows in slide a). In b, six cell plates can be seen;
only two have additional callose deposits. (c–d) Tetragonal tetrads. (c) Naked cell plates. (d) The additional callose deposits are concentrated at the intersection
of the four cell plates close to the callose wall surrounding the tetrad (arrows). (e–j) P. tenax. (e–f) Upper views of two different tetrads at different stages of
callose deposition. Callose appears to be deposited on the cell plates in the middle part of the wall (arrow in e) and to progress toward the intersection of the
walls (compare e, depicting the beginning of the process and f, depicting the end of the process). (g–j) Middle and upper views of two different tetrads illustrating
the beginning of callose deposition (g–h) and the fully developed tetrad (i–j), confirming another orientation of callose progression in tetrads. (k–l) Two different
tetrads of H. foetidus. (k) Additional callose seems to be deposited simultaneously at the center of the tetrad and at its border (arrows). (l) End of tetrad
formation. Deposits converge toward the middle of the cell plates. (m–t) P. lepidocarpodendron. (m) Beginning of the process. Cell plates first thicken.
(n) Center of a tetrad. Cell plates start to be embedded in non-uniform callose deposits that begin to accumulate at the distal poles of the microspores.
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Fig. 4. Fluorescent light micrographs of aperture distribution within tetrads. (a–h) Asphodelus albus. After staining with aniline blue. (i) Phormium tenax.
After staining with congo red. (a), (c), (e), and (h): callose walls, using filter set 01. (b), (d), (f), and (g): microspores using filter set 07. (a–f) Upper middle
and lower view of an irregular tetrahedral tetrad of A. albus. The distal furrows, seen on most microspores, appear in between the additional callose deposits,
orthogonal to the largest cleavage walls. (g) Tetragonal tetrad, using filter set 07. (h) The same tetrad, using filter set 01. The microspores are all monosulcate,
and the sulcus is also orthogonal to the cleavage walls. (i) P. tenax. The ends of the trichotomosulcus are joined three by three.
study (Longly and Waterkeyn, 1979a; Brown, 1991; Brown
and Lemmon, 1996). Other studies have indicated that plant
meiosis may or may not resemble mitosis in terms of the formation of a phragmoplast, depending on the specific plant species studied (Brown and Lemmon, 1996). Because we did not
label microtubules, we do not know whether a phragmoplast
is produced in the two monocot species studied.
Additional callose deposits: a key role in aperture pattern
determination?—The species studied differed markedly in
terms of additional callose deposits on cell plates. The pattern
of callose deposition is hard to follow: callose observed by
epifluorescence is a translucent material, and callose deposits
within tetrads are observed across the callose wall surrounding
the tetrad. In addition, tetrahedral tetrads are complex tridi-
←
Fig. 3. Continued. (q) Same stage in another tetrad. Callose deposits are thicker in the middle of the cleavage wall and at the periphery of the tetrad (arrow).
(r) End of process, side view. Callose deposits increase in the middle of the cleavage wall at the periphery of the tetrad (arrow), then toward the intersections
of the cleavage walls and the wall surrounding the tetrad. (o–s) Middle and upper view of a tetrad at an intermediate stage of callose deposition. Callose is
deposited in the middle of the cleavage wall. (p–t) Middle and upper views of a tetrad after callose deposition is completed. Comparison between o and p
indicates that callose deposition has progressed toward the intersections of the cleavage walls.
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Fig. 5. Diagram of the progression of further deposition of callose on the cell plates. Left, Phormium tenax and Protea lepicarpodendron type. Callose
deposits begin in the middle part of the cell plates (arrows) and progress towards the intersections of the cell plates (black dots). Right, Helleborus foetidus
type. Callose is deposited (arrows) from the edges of each cell plate and progresses towards the middle of the cell plate (black dots).
mensional objects that hide part of the cleavage walls that
isolate the microspores behind the other walls. Observation of
the different areas of a single tetrad is rarely possible. In extreme cases, additional callose deposits are so abundant that
they prevent any detailed observation of the underlying patterns. As a result, one can only form a general idea of the
progression of callose depositition. Two main ways of callose
deposition were observed (Fig. 5; note that the figure only
indicates the progression of the callose deposits and does not
present a realistic image of additional callose deposits). In A.
albus and H. foetidus, additional callose deposition begins at
the intersection of the cleavage walls and progresses centripetally inwards along the cell plates (only on two of the cleavage walls in A. albus). Alternatively, in P. tenax and P. lepicarpodendron, callose deposition begins on the cell plate and
progresses centrifugally towards the intersection of the cleavage walls (this time our observation is consistent with the observations made in the two other Proteaceae species studied
by Blackmore and Barnes [1995]). We thus observed a convergence in patterns of callose deposition between monocots
and eudicots, while cell plate formation remains different between these clades. Interestingly, cell plate formation and additional callose deposition appear to be completely independent. This was already known to be true for monocots and
basal angiosperms, but has never been confirmed for eudicots
(Longly and Waterkeyn, 1979a, b; Bandhari, 1984; Blackmore
and Crane, 1998). This study thus confirms that cleavage wall
formation is the result of two independent variable elements,
namely cell plate formation and additional callose deposition.
Additional callose deposits appear to predict future aperture
sites the best in all four species. The convergence in callose
deposition patterns between P. tenax and P. lepicarpodendron
is correlated with a convergence in aperture relationships within the tetrads (Garside’s arrangement of apertures), despite dramatic differences in aperture pattern (P. tenax displays a single, branched furrow, while P. lepicarpodendron has three
equatorial pores). In contrast, H. foetidus, which differs in
terms of callose deposition, displays different relationships of
apertures within the tetrad (Fisher’s arrangement of apertures).
This correlation between additional callose deposits and areas
where apertures meet is confirmed by aperture distribution
within tetrads, because the places where additional callose deposition is completed coincide with aperture location in all of
the species. This seems also to hold true for A. albus, where
the position of the ends of the furrow is consistent with the
progression of callose deposition. The difference between A.
albus (monosulcate) and H. foetidus (tricolpate) appear to be
driven by other mechanisms. In A. albus, cleavage walls are
not all identical, and this may explain the asymmetric distribution of the additional callose deposits, which occur on two
of the cell plates only and are absent from the center of the
forming tetrad. The effect of differences in tetrad shape is
beyond the scope of this paper. Further studies, focused on
species that display variation in tetrad shape, would be necessary to investigate this aspect of microsporogenesis.
The present study has implications for our understanding of
the genetic control of aperture pattern. It appears to be a highly
complex character despite its apparent simplicity. Both the
model (Ressayre et al., 2002) and the observations reported
here converge to indicate that aperture pattern is determined
by the places where cytokinesis following meiosis is completed. The places themselves are determined by a number of
events (cytokinesis type, orientation of the second meiotic axes
that affects tetrad shape and pattern of callose deposition following cell plate formation) that individually appear to be controlled by at least one gene. The cell progression cycle during
male meiosis is probably controlled by several genes (Magnard et al., 2001). Several non-allelic mutants presenting cytokinesis defects leading to microsporogenesis failure have
been described in potatoes (Mok and Peloquin, 1975) and in
Arabidopsis mutants (Peirson et al., 1996; Hülskamp et al.,
1997; Spielman et al., 1997). A mutant disrupting the normal
orientation of the second meiotic axes has also been described
in potatoes (Mok and Peloquin, 1975). In the same way, callose deposition and degradation is controlled by several different genes (Fei and Sawhney, 1999). Our study indicates that
the genetic system controlling cell plate formation and additional callose deposits is almost certainly different. In addition,
the genetic control of additional callose deposits is plausibly
quantitative because the amount of additional callose deposits
as well as their progression vary strongly between species. For
example in monocots, depending on the species, the additional
callose deposits have been described to be either absent or
inconsistent, present but not oriented, or present and progressing either centripetally or centrifugally (Sampson, 1969; Longly and Waterkeyn, 1979b; Ressayre, 2001). Additional variation can be described; there are, for example, several different
ontogenetic pathways for producing apertures in angiosperms
(Rowley, 1975). As a result, aperture pattern appears to be a
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highly complex character controlled by at least half a dozen
genes, but probably by many more.
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ARENS, K. 1949. Prova de calose por meio da microscopia a luz fluorescente
e aplicações do metodo. Lilloa 18: 71–75.
BANDHARI, N. N. 1984. The microsporangium. In B. M. Johri [ed.], The
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