Department Biological Structure and Function SD, Oregon Health Sciences University, Portland, OR 97201-3097 USA
* Author for correspondence (e-mail: danilchi{at}ohsu.edu)
Accepted 10 October 2002
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Summary |
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Key words: Cleavage furrow, Cytokinesis, D2O, Exocytosis, Fusion pore, Microtubule, Xenopus
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Introduction |
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Amphibian embryos are distinctive among dividing cells for the
comparatively large amount of membrane that is added continuously during the
cleavage process (Selman and Perry,
1970). For example, in Xenopus, as much as 1.4
mm2 of new plasma membrane appears in the cleavage plane during
15 minutes of furrow progression
(Bluemink and de Laat, 1973
).
This new membrane is known to differ qualitatively from that of the original
egg surface (Kalt, 1971
;
Sanders and Singal, 1975
;
Byers and Armstrong, 1986
;
Servetnick et al., 1990
;
Bieliavsky et al., 1992
;
Aimar, 1997
). The main source
of the new membrane appears to be a pool of oogenetically produced vesicles
(Leaf et al., 1990
;
Servetnick et al., 1990
) that
contributes not only membrane lipids, but also glycoproteins, and
extracellular matrix components that ultimately line the surface of the
blastocoel (Kalt, 1971
;
Servetnick et al., 1990
).
How membrane expansion is regulated during cleavage is not understood. In
particular, it is not yet clear where exocytosis actually takes place. It has
been known since Zotin's pioneering studies in sturgeon and amphibian embryos
(Zotin, 1964) that vesicles
accumulate in the spindle midzone beneath the advancing first cleavage furrow,
suggesting that membrane addition might occur at the leading edge of the
furrow, contemporaneously with its advance through the midzone. In support of
this idea, Sawai's tracing of carbon particle motions on the surface of
cleaving newt eggs suggested that membrane expansion occurs from the furrow
base (Sawai, 1987
).
Alternatively, the persistence of a stable original-membrane domain at the
leading edge of the furrow, including membrane glycoproteins
(Byers and Armstrong, 1986
)
and collections of microvilli
(Denis-Donini et al., 1976
),
has suggested that membrane addition occurs elsewhere along the cleavage
plane, for example, at the margin between new and old domains or along its
entire length. This idea has support from electron microscopic studies that
show a variety of putative exocytotic vesicles at various sites along the
cleavage plane, but not necessarily near its leading edge
(Bieliavsky and Geuskens, 1990
;
Singal and Sanders, 1974
).
In Xenopus, vesicles contributing basolaterally targeted
U-cadherin to each early cleavage plane are distributed uniformly in the
peripheral cytoplasm until cleavage (Angres
et al., 1991). Thus, however it occurs, vesicle recruitment to the
site of exocytosis must be locally regulated in the vicinity of each furrow.
The possibility that microtubules might be involved in localized basolateral
vesicle recruitment to the cleavage furrow was raised when Danilchik et al.
(Danilchik et al., 1998
) and
Jesuthasan (Jesuthasan, 1998
)
identified novel microtubule-containing structures at the bases of cleavage
furrows in Xenopus and Brachydanio, respectively. These
structures are now referred to as furrow microtubule arrays (FMAs) to
distinguish them from other microtubule-containing structures in the cleavage
plane (e.g. midbodies and interzonal microtubules). Microtubule-disrupting
experiments indicated that basolateral membrane growth in the cleavage plane
requires microtubules (Danilchik et al.,
1998
; Jesuthasan,
1998
). Similarly, Larkin and Danilchik found that furrow
microtubules are required in cleaving sea urchin embryos to complete the
closure of the cytoplasmic bridge, and proposed that one function of furrow
microtubules is to direct vesicles toward a site of fusion at the cytoplasmic
bridge to accomplish cell separation
(Larkin and Danilchik, 1999
).
More recently, Skop et al. (Skop et al.,
2001
) and Shuster and Burgess
(Shuster and Burgess, 2002
)
have identified microtubule-dependent membrane-addition events in the final
stages of cytokinesis in C. elegans and sea urchin egg cleavage. This
apparently general requirement for microtubules in the localization of
membrane addition to terminate cytokinesis seems to reflect a direct
recruiting of vesicles to the furrow base.
In the present report, scanning electron microscopy (SEM) and live-embryo confocal imaging revealed clusters of exocytotic fusion pores in the immediate vicinity of the expanding furrow base. Nocodazole treatment both randomized and reduced the number of these pores, indicating that microtubules are indeed required for a step in localizing vesicle exocytosis. To test whether microtubules are sufficient to direct this localization process, D2O was used to generate ectopic microtubule monasters. D2O-treated embryos underwent a rapid, nearly uniform expansion of the surface; SEM analysis indicated large numbers of exocytotic fusion pores randomly scattered across the entire surface. This effect of D2O was abolished with nocodazole, confirming that microtubules near the surface, near the end of the cell cycle are sufficient to recruit exocytotic vesicles to the plasma membrane, and supporting the hypothesis that furrow microtubule bundles play a similar role in the furrows of dividing cells.
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Materials and Methods |
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Treatments
Stock solutions of nocodazole, cytochalasin B, cytochalasin D and
latrunculin B (Sigma) in dimethylsulfoxide (DMSO) were diluted to
concentrations of 5 or 10 µg/ml in MMR/3 and applied to embryos at
specified times. Controls consisted of similar dilutions of DMSO in MMR/3.
D2O was diluted to 60% in MMR/3 and applied to embryos at specified
times during early development.
Immunostaining
For examining embryos via wholemount confocal immunocytochemistry, we
adapted Gard's (Gard, 1993)
protocol, as described previously
(Danilchik et al., 1998
).
Embryos were fixed for 2-4 hours at room temperature or overnight at 4°C
in Gard's fixative (3.7% formaldehyde, 0.25% glutaraldehyde, 0.2% Triton
X-100, 80 mM PIPES, pH 6.8, 5 mM EGTA, 1 mM MgCl2). After fixation,
embryos were stored in methanol overnight at -20°C. Pigmented embryos were
bleached in 10% H2O2 in methanol on a light table for
1-4 hours (bleaching time varied, depending on rate of progress of pigment
fading). After bleaching, embryos were rehydrated in three consecutive rinses,
for 10 minutes each, of: 50% MeOH in TBS, 25% MeOH in TBS, 100% TBS (1x
TBS: 155 mM NaCl, 20 mM Tris-Cl, pH 7.4). To decrease background fluorescence
caused by glutaraldehyde and autofluorescence of yolk platelets, embryos were
placed in a reducing solution of 100 mM NaBH4 in TBS for 4 hours
(at room temperature) or overnight (at 4°C). NaBH4 was removed
by rinsing in NTBS five times over the course of 1 hour (NTBS: 1x TBS,
with 0.1% NP-40).
Vitelline envelopes were removed prior to exposing fixed specimens to
antibody. Some embryos were bisected with a razor blade fragment, cutting
parallel to the animal-vegetal axis, either perpendicular to or paralleling
the cleavage plane. Other embryos were processed intact for wholemount
observation. Intact or bisected embryos were incubated with antibodies diluted
in TBS with 10% fetal bovine serum and 5% DMSO overnight at 4°C, with
agitation, with five one-hour washes in NTBS after each incubation. Primary
antibodies included monoclonals against ß-tubulin (Biogenesis;
1:1000),
-tubulin (Sigma; 1:200), VSVG (clone P5D4; Sigma; 1:400) and
sheep polyclonal antibodies against
ß-tubulin (Cytoskeleton;
1:200). Secondary antibodies were Alexa-546- or -488-conjugated goat
anti-mouse or donkey anti-sheep (Molecular Probes, 1:100). After
immunostaining and subsequent washes in NTBS, embryos were dehydrated in two
rinses of MeOH, 30-60 minutes each, and then cleared in two rinses of Murray's
clear (2:1::benzyl benzoate:benzyl alcohol).
Surface labeling, time-lapse recording, confocal microscopy and image
analysis
The growth of new membrane during cell division was visualized in
devitellinated embryos exposed to fluorescent soybean agglutinin for five
minutes before first cleavage (Alexa 488 conjugate; 125 µg/ml; Molecular
Probes). Unbound lectin was removed via several exchanges of MMR/3 prior to
time-lapse recording.
To view membrane expansion, a suspension of activated charcoal particles was pipeted over devitellinated embryos prior to time-lapse recording. Time-lapse sequences were recorded using an NEC color CCD camera mounted on an Olympus SZH stereoscope. Images were captured on a Panasonic LQ-3031 optical disk recorder, digitized via an Xclaim VR 128 graphics card (ATI), and converted to TIFF image stacks for further analysis using Wayne Rasband's NIH Image (v. 1.62; http://rsb.info.nih.gov/nih-image), or to QuickTime (Apple) movies using Quicktime Pro or Adobe Premiere 5.1.
To display carbon particle motion as a kymograph, TIFF time-lapse image stacks were opened in NIH Image, assigned a 1-pixel virtual spacing between slices, and selected regions were then rotated 90° about the Y axis to display elapsed time along the X-axis as 1 pixel per captured frame.
Exocytotic fusion pore distributions in the cleavage furrow were measured in panoramic montages of digitized SEM images across relevant surfaces registered using Adobe PhotoShop. NIH Image's particle analysis package was then used to determine pore positions relative to the base of the furrow (leading edge). Pore counts in adjacent 10 µm x 50 µm regions of interest were determined and their local density was then plotted as a function of distance from the base of the furrow.
To examine membrane dynamics in vivo at high magnification, devitellinated
embryos were bathed in MMR/3 containing 10 µM cytochalasin B and 10 µM
FM1-43 (Molecular Probes) and examined via confocal microscopy (BioRad
Radiance 2100; Nikon E800 upright microscope using a 60x/1.0 NA
CFI60 Fluor dipping objective). Focus on the surface of the animal
hemisphere was readjusted frame by frame manually because the devitellinated
embryos' height rises and falls rapidly throughout the cell cycle
(Hara et al., 1980).
Scanning electron microscopy
For scanning electron microscopy, intact or devitellinated embryos were
fixed in 2.5% glutaraldehyde in 0.1 M Na-cacodylate, pH 7.5 overnight at
4°C. Fixed specimens were then transferred into 0.1 M Na-cacodylate, pH
7.5 at 4°C. If necessary, embryos were manually devitellinated at this
point, and then dehydrated via 15-minute exchanges with 50, 75 and 95%
ethanol, followed by three 30-minute exchanges with 100% ethanol. Specimens
were rinsed twice with hexamethyldisilizane (Polysciences) for 30 minutes.
Specimens were then allowed to dry overnight at room temperature in open
vials. More rapid, vacuum-driven removal of the hexamethyldisilizane was found
to be unsatisfactory, since it often resulted in tissue contraction or surface
rupture. Dried specimens were mounted onto aluminum stubs using silver paste,
sputtercoated with gold-palladium to 50 nm on a Hummer VII (Analect, USA), and
then examined with a JEOL T330A scanning electron microscope.
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Results |
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|
|
Furrow microtubules underlie membrane addition site
As was earlier shown (Bluemink and de
Laat, 1973; Drechsel et al.,
1997
; Danilchik et al.,
1998
), localized membrane addition continues unabated after
disruption of the contractile ring by treatment with cytochalasin B or
function-blocked rho protein, with the result that a broad stripe of
unpigmented surface develops at the presumptive cleavage plane. Here, we show
similar results with another microfilament inhibitor, latrunculin B
(Fig. 2A). The same embryo
fixed, processed, and stained to reveal microtubules via confocal wholemount
microscopy displays prominent bundles of microtubules in close proximity to
the surface (i.e. a few microns) along the entire length of the disrupted
furrow (Fig. 2B). Similar
results were obtained with both cytochalasin B and D.
|
Surface particle motions indicate site of membrane expansion
We used a method developed previously
(Sawai, 1987) to follow the
movement of small carbon particles dropped onto the surface of devitellinated,
cleaving Xenopus embryos treated with cytochalasin B to expose the
growing membrane domain. Video time-lapse recordings of particle motions
(Fig. 3A-D) were reprocessed to
produce kymographs displaying particle movement along the ordinate and elapsed
time on the abscissa (Fig. 3E).
Particle drift away from the furrow base was relatively steady, averaging 35
µm/minute. Particles landing near each other on the same side of the furrow
traveled at nearly the same rate without drifting apart, indicating that
little or no membrane expansion took place between them once they had moved
beyond the furrow base. In contrast, particles landing on opposite sides of
the furrow base traveled away from each other. Some particles, landing
directly over the furrow base, remained there for several minutes before
abruptly commencing to drift toward one side or the other. These results
indicate that most of the membrane expansion occurs from a site or sites
within
50 µm of the furrow base. Similar results (not shown) were
obtained in embryos not treated with cytochalasin.
|
Localized sites of exocytosis: scanning electron microscopy
Scanning electron microscopy was used to examine the new surface of
devitellinated, cleaving embryos for evidence of exocytosis. At low
magnification (Fig. 4A,C), the
new membrane domains appeared as a pair of relatively smooth triangular areas
on either side of the cleavage furrow (Fig.
4A). At higher magnification, surveys revealed large numbers of
circular pits or craters in the otherwise smooth new-membrane domain
(Fig. 4B,D). Pits ranged in
size from 0.5 to 2.0 µm in diameter. Two irregular stripes, roughly 10
µm wide, at either side of the furrow base itself were nearly devoid of
pits (Fig. 4B,D). Similar
analysis of embryos fixed at different times during first cleavage indicated
that generally more than 75% of these pits were concentrated within 50 µm
of the furrow base during the membrane expansion phase. Few pits were found
near the margin between new and old membrane domains. For example,
Fig. 5 (filled circles) plots
pit density as a function of distance from the furrow base. In embryos treated
with nocodazole during furrow formation, both the number and concentration of
surface pits were significantly reduced
(Fig. 5, open circles).
|
Localized sites of exocytosis: confocal time-lapse
The above SEM observations are consistent with the idea that the pits
represent fusion pores at sites of recent or ongoing vesicle exocytosis. As an
independent test of this hypothesis, we searched for evidence of exocytosis in
live embryos in which the plasma membrane was loaded with the fluorescent
styryl dye FM1-43. Although FM1-43 is most commonly used in pulse-chase
experiments to selectively label recycling exocytotic vesicles
(Betz et al., 1996), it has a
relatively high affinity for new membrane in the cleavage plane of
Xenopus embryos (see below), making it useful for identifying sites
of ongoing exocytosis. Devitellinated embryos undergoing first cleavage were
incubated continuously in medium containing both cytochalasin B and FM1-43. As
with latrunculin (Fig. 3), the
cytochalasin disrupts furrow deepening, thereby making the membrane addition
site accessible for viewing by confocal microscopy
(Fig. 6). FM1-43 gradually
accumulated in the plasma membrane, preferentially labeling the new membrane
domain (Fig. 6, asterisk
indicates less-labeled original-membrane of the animal hemisphere surface).
Although some endocytosis has been detected in the new membrane domain of
Xenopus (data not shown), consistent with recent work in zebrafish
(Feng et al., 2002
), the
intense labeling along the margins between new and old membrane domains
(Fig. 6) primarily reflects the
large amount of membrane involved with microvilli and other protrusions in
this region (Denis-Donini et al.,
1976
).
|
As embryos entered second cleavage, a new site of membrane addition
appeared near the disrupted second furrow (arrow,
Fig. 6). This expanding area
was significantly less fluorescent than other regions within the new membrane
domain, suggesting ongoing localized addition of unlabeled membrane.
Time-lapse confocal recordings at high magnification in this region confirmed
this idea. As an example, Movie 2 (see
http://jcs.biologists.org/supplemental)
displays a particularly active site recorded near the arrow in
Fig. 6. This movie segment
shows the abrupt appearance of irregular dark patches, 10-20 µm2
in area, which quickly diffused into the surrounding FM1-43-labeled membrane.
Still frames representing three successive sequences from this time-lapse
recording are presented in Fig.
7. In the sequence shown in panels A1-A4, the arrows indicate a
single site of membrane expansion in which a patch of unlabeled membrane
abruptly grew from 10 to 25 µm2, before fading into the
surrounding labeled surface. Similar single-fusion events are shown
(Fig. 7B1-B4).
|
In some cases, circular structures, apparently stable exocytotic fusion pores, were seen to associate with the membrane for several minutes before abruptly flattening into the plane of the membrane and contributing unlabeled patches to it. For example, in the sequence shown in Fig. 7C1-C4, a 2.5 µm circular fluorescent structure (the same structure had also been present through the previous two sequences) suddenly disappeared to be replaced by an irregular patch of unlabeled surface (cf. arrows in Fig. 7C2-C3). The sudden introduction of the unlabeled patch evidently contributes surface area to the local membrane: by looping the movie sequence back and forth a few frames across this event, small surface particles can be seen to spread a short distance in all directions away from the patch.
A densitometric profile across the structure indicates that the membrane within the intensely labeled edges contains significantly less label than the surrounding plasma membrane (not shown). The brightly labeled edge evidently constitutes a barrier to lateral diffusion between the membrane of the vesicle and that of the cell surface until the abrupt flattening of the vesicle into the plane of the membrane.
VSVG protein exocytosis
To learn the size range of vesicles destined for exocytosis along the
basolateral surfaces, we injected fertilized eggs with capped, synthetic mRNA
encoding the full-length sequence of the basolaterally targeted viral coat
glycoprotein of vesicular stomatitis virus (VSVG; plasmid gift of H.-P. Moore,
Berkeley, CA) and fixed and processed them for wholemount immunocytochemistry
using an anti-VSVG antibody (Sigma). Fig.
8 is a projection of a confocal image stack showing a collection
of VSVG-labeled vesicles near the base of a cleavage furrow. The surface is
also labeled, indicating recent exocytosis of similarly labeled vesicles at or
near this site. Vesicles ranged in diameter from 0.6 µm to 2.0 µm,
which is similar to the size of the presumed exocytotic pits shown above via
SEM.
|
D2O induces ectopic membrane expansion
We reported previously that embryos exposed to high concentrations of
D2O form numerous ectopic monasters
(Danilchik et al., 1998). The
concomitant disturbance in cortical pigmentation had suggested new membrane
addition, consistent with the idea that the presence of polarized microtubules
near the surface is sufficient to provoke new membrane addition. To examine
this possibility further, we repeated our earlier experiment, this time
removing the vitelline envelope prior to incubation in D2O. The
experiment shown in Fig. 9
confirms that D2O provokes a rapid, essentially random expansion of
the embryo surface. Without the support of a vitelline envelope, embryos
normally flatten slightly onto the substratum and assume an oblately
spheroidal morphology (Bluemink and de
Laat, 1973
). In contrast, in the presence of D2O, the
surface of the embryos rapidly expanded at about the time of first cleavage
(Fig. 9A,B). The overall
spreading and thinning of the pigmented animal hemisphere suggests that the
new surface area was introduced randomly across the entire surface. Scanning
EM analysis (Fig. 10) confirms
that exocytotic fusion pores appeared in large numbers in the smooth surface
regions between clusters of microvilli
(Fig. 10B), consistent with
the idea that D2O completely randomizes the site of membrane
addition. Because microvilli were retained in large numbers throughout the
D2O-driven surface expansion, it is unlikely that the observed
increase in surface area is simply due to microvillar shortening.
|
|
D2O-induced expansion is blocked with nocodazole
With the appearance of new surface, the overall height of the embryo
concomitantly collapsed to 250-350 µm
(Fig. 9C1-C6). To estimate the
amount of cell surface area expansion induced by D2O treatment, we
regarded the flattening embryos as a family of progressively more oblate
spheroids, with major (horizontal) and minor (vertical) axes that could be
used to calculate surface areas. Radii measured across the growing horizontal
profiles of the embryos yielded values for the major half-axis, `a'. Since
embryo volume (V) probably changed little during the course of an experiment,
the following relationship,
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|
The D2O-induced membrane expansion appears to be entirely microtubule-dependent, as indicated by the inhibition of expansion by 10 µM nocodazole (Fig. 11). To determine whether the nocodazole treatment actually disrupts microtubule formation in D2O-treated embryos, control and treated embryos were fixed at 105 minutes post-fertilization, and examined via confocal microscopy for the presence of microtubules (Fig. 12). In an untreated embryo, aligned microtubule bundles of the FMA decorated the base of the first cleavage furrow (Fig. 12A). Elsewhere on the animal hemisphere of the same embryo, only a few microtubules were found near the surface (Fig. 12B). Vertically resectioning the same confocal stack (Fig. 12B, inset) revealed the normally low concentration of microtubules at the surface. In contrast, following treatment with D2O, numerous microtubule monasters were found near the cell surface (Fig. 12C and vertical resection, inset). Nocodazole completely abolished microtubules in an embryo incubated in normal medium (Fig. 12D and inset), and effectively depolymerized microtubules near the surface of D2O-treated embryos, even though monasters apparently persisted more deeply in the cytoplasm (Fig. 12E and inset). These results are consistent with a requirement for intact microtubules in membrane expansion; with the results of Figs 9,10,11 they indicate that the presence of microtubules near the surface is sufficient to induce membrane expansion.
|
Polarity of microtubules in D2O-induced ectopic
monasters
D2O-treated embryos were double-stained for both ß- and
-tubulin and examined via confocal microscopy. Monastral microtubules
were seen to extend in all directions from
-tubulin-rich centers
(Fig. 13). Since
-tubulin is a centrosomal protein that associates with the minus ends
of microtubules (Oakley et al.,
1990
; Stearns et al.,
1991
; Stearns and Kirschner,
1994
; Li and Joshi,
1995
), we conclude that most of the microtubules interacting with
the cortex in D2O-treated embryos are oriented with plus ends
facing the surface.
|
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Discussion |
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The present report addressed two related issues in basolateral membrane
formation during Xenopus cleavage: the site of membrane insertion
along the furrow, and the potential role of furrow microtubules in regulating
this site. We found that the bulk of membrane addition proceeds via exocytosis
from a site within 50 µm of the furrow base. This location is
significant, since it overlies the ends of microtubules of the furrow
microtubule array, and is therefore consistent with the idea that the
microtubules mobilize the exocytotic vesicles involved in the membrane
expansion. Also consistent with this hypothesis was the finding that
ectopically placed microtubule plus ends near the cell surface in
D2O-treated embryos are sufficient to provoke ectopic new membrane
addition. Therefore, we suggest that vesicles encountering furrow microtubules
at the advancing furrow tip are transported toward the microtubule plus ends
to become concentrated at either side of the base of the furrow.
Fusion pores and membrane expansion
Exocytotic fusion pores are transitory aqueous channels connecting the
lumen of vesicles with the extracellular space during the process of
exocytosis. As originally defined, pores are small structures, 20 to 100 nm in
diameter (Breckenridge and Almers,
1987). We were initially inclined to dismiss the 2 µm pits seen
via scanning EM as fixation artefacts. However, their nonrandom distribution,
relatively uniform size range, and absence under conditions preventing new
membrane addition (e.g. nocodazole) suggested that they might represent
authentic exocytotic fusion pores. Similar structures in the 2 µm diameter
range, observed via SEM in secretory alveolar type II cells, are accepted as
authentic fusion pores, with channels remaining open for up to several hours
(Haller et al., 2001
).
Similarly, large, stable cortical crypts or pits remain in the surface for
several minutes following cortical granule exocytosis in zebrafish
(Becker and Hart, 1999
).
By independent methods, we visualized 2 µm hollow cavities in the
growing surface of living embryos that resemble those seen in SEM. Time-lapse
confocal microscopy of embryos stained with FM1-43 gave us a glimpse of the
dynamic, transitory nature of these exocytotic fusion pores. Some apparently
stable fusion pores remained at the surface for up to several minutes before
flattening into the plane of the cell surface as a patch of unlabeled
membrane. The area of each patch was approximately 10-20 µm2, an
amount of surface that could have been provided by individual vesicles ranging
in diameter from 2.2 to 2.8 µm. We asked whether the density of fusion
pores seen in cleavage furrows could provide a sufficient number of exocytotic
events to account for the large amount of membrane introduced during cleavage.
Bluemink and de Laat (Bluemink and de Laat,
1973
) estimated that
2.3 mm2 of new membrane is
introduced ectopically to the surface following cytochalasin B treatment. Our
embryos were smaller than those used by Bluemink and de Laat (1.16 mm
diameter); we estimate the total new membrane in our cytochalasin-treated
embryos (Fig. 6) to be
1.85 mm2. Assuming that each fusion pore represents a
potential delivery of 20 µm2, as suggested from the FM1-43
labeling experiment (Fig. 7C),
then
92,500 vesicles would be needed. Since membrane addition is
continuous over a period lasting
15 minutes, only a limited number of
fusion pores should be seen at any time in fixed SEM specimens. The profile of
fusion pore density in Fig. 5
shows 27±3 fusion pores per adjacent 500 µm2 regions
nearest the furrow base. Again, if each pore represents 20 µm2
of potential surface area, this density of exocytotic vesicles could represent
99,000 vesicles. In other words, the fusion pore clusters near the furrow
base are of a density consistent with that capable of providing the entire
1.85 mm2.
Microtubule polarity
D2O-stabilized microtubules appear to be fully capable of
supporting organelle transport in Xenopus embryos, in a reportedly
`randomized' fashion (Rowning et al.,
1997). However, as we have shown here, D2O-stabilized
microtubules are evidently not randomized; instead, they form well-organized,
ectopic monasters (see also Danilchik et
al., 1998
). The presence of centriole-like structures
(van Assel and Brachet, 1966
)
and, as shown in this report,
-tubulin at the foci of these monasters,
indicate that the D2O-stabilized monasters resemble conventional
MTOCs, with microtubule plus ends radiating away from organizing centers. In
effect, the entire cortex becomes enriched with overlapping arrays of
microtubule plus ends, a situation that normally only occurs along the
cleavage plane. Since the effect of D2O on membrane expansion is
blocked with nocodazole, we conclude that the exocytosis of vesicles depends
on a plus-ward transport of vesicles. This conclusion is thus consistent with
that of experiments demonstrating kinesin-dependence in the microtubule-based
mobilization of vesicles toward sites of wound-induced exocytosis
(Bi et al., 1997
).
We still lack direct information about the polarity of microtubules in the
FMA. From the SEM data shown here, we know that the site of membrane addition
is just to either side of the furrow base, making it unlikely that the FMA
transports vesicles toward the midline. Rather, the observed sites of
exocytosis are consistent with vesicles being transported away from the
midline. However, this leads to a surprising result: if exocytotic vesicles
are indeed plus-directed, as suggested by the D2O experiments, then
the FMA bundles should be oriented with minus ends toward the midline. Such a
microtubular arrangement is clearly in conflict with the known organization of
plant-cell phragmoplasts (Staehelin and
Hepler, 1996) and animal-cell midbodies (Eutenauer and McIntosh,
1980). To resolve this issue, we are presently investigating the polarity of
FMA microtubules by following the motion of GFP-tagged EB1 in cleaving
Xenopus embryos (E.E.B. and M.V.D., unpublished).
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Acknowledgments |
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Footnotes |
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References |
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