Development 122, 1187-1194 (1996)
Printed in Great Britain © The Company of Biologists Limited 1996
DEV0050
1187
Coordination of cellular events that precede reproductive onset in
Acetabularia acetabulum: evidence for a ‘loop’ in development
Linda L. Runft† and Dina F. Mandoli*
Department of Botany 355325, University of Washington, Seattle, WA 98195-5325, USA
*Author for correspondence
†Present address: The University of Connecticut Health Center, Department of Physiology, Farmington, CT, USA
SUMMARY
Amputated apices from vegetative wildtype cells of the
uninucleate green alga Acetabularia acetabulum can differentiate a reproductive structure or ‘cap’ in the absence of
the nucleus (Hämmerling, J. (1932) Biologisches Zentralblatt 52, 42-61). To define the limits of the ability of wildtype cells to control reproductive differentiation, we determined when during development apices from wildtype cells
first acquired the ability to make a cap in the absence of
the nucleus and, conversely, when cells with a nucleus lost
the ability to recover from the loss of their apices. To see
when the apex acquired the ability to make a cap without
the nucleus, we removed apices from cells varying either
the developmental age of the cells or the cellular volume
left with the apex. Cells must have attained the adult phase
of development before the enucleate apex could survive
amputation and make a cap. Apices removed from cells
early in adult growth required more cell volume to make a
cap without the nucleus than did apices removed from cells
late in adult growth. To define the limits of the cell to recapitulate development when reproduction falters, we
analyzed development in cells whose caps either had been
amputated or had spontaneously aborted. After loss of the
first cap, cells repeated part of vegetative growth and then
made a second cap. The ability to make a second cap after
amputation of the first one was lost 15-20 days after cap
initiation. Our data suggest that internal cues, cell age and
size, are used to regulate reproductive onset in Acetabularia
acetabulum and add to our understanding of how reproduction is coordinated in this giant cell.
INTRODUCTION
actions within the cell and with its environs, not by signals in
or from multicellular tissues or organs. The cellular dialog
between the nucleus, which is located in the basal rhizoid of
the cell, and the cell apex, which is the site of differentiation,
occurs over a long distance (3-4 cm). Second, the age of a cell
can be assessed because the stalk is decorated with rings of
hairs, or ‘whorls’, that leave scars whose spacing is characteristic of the phase of development (Nishimura et al., 1992b).
Third, it’s giant size and ability to heal wounds (Fester et al.,
1993; Menzel, 1980) facilitates manipulations such as cell
grafting (Berger et al., 1987; Menzel, 1994) and allows the
developmental potential of pieces of the cell to be analyzed.
Finally, the cell can be stably transformed (Neuhaus et al.,
1986), expresses and correctly targets proteins encoded by heterologous or foreign DNA readily (Neuhaus et al., 1984), and
is amenable to biochemical (Berger et al., 1987) and genetic
manipulations (Mandoli and Larsen, 1993). Comprehensive
knowledge of development in wildtype cells is important to
make full use of these features.
Reproductive onset is marked by an explicit shape change
at the cell apex, initiation of a cup-shaped structure or ‘cap’,
and is regulated by both internal and external cues. The internal
cues are the age and size of the cell (Nishimura et al., 1992b),
the presence of a putative population of cap-specific mRNAs
Coordination of cellular events at reproductive onset is
important for the unicellular alga, Acetabularia acetabulum:
since virtually all of the contents of the cell are donated to its
progeny, if the parent cell fails to coordinate nuclear and cytoplasmic events it will die without producing offspring. Thus,
reproductive onset in A. acetabulum may be more tightly
regulated than in organisms whose commitment to reproduction does not entail such drastic consequences for the species.
Accordingly, the cell has evolved strategies both to prevent
premature entry into reproduction and to reiterate reproductive
development. For example, it survives winters by retracting the
cytoplasm into the rhizoid (Berger and Kaever, 1992; Dao,
1962), it can stall development between the phases in vegetative growth (Mandoli, unpublished data), and the cell can
execute a ‘developmental loop’ by aborting reproduction (A.
crenulata: Puiseux-Dao, 1963; A. mediterranea: Decléve et al.,
1972) if environmental conditions are not optimal or the
original apex is lost. The strategies used by the cell to ensure
reproductive success are not well understood.
This organism is attractive for developmental studies for
many reasons. First, in contrast to multicellular organisms,
reproductive onset in A. acetabulum is regulated only by inter-
Key words: Acetabularia acetabulum, competence, differentiation,
reproduction, gametangium
1188 L. L. Runft and D. F. Mandoli
and had 1-5 whorls consisting of clear, ephemeral hairs that branched
0-3 times each. ‘Early adults’ had thin stalks, were 1-3 cm tall and
had more than 6 green whorls comprised of hairs that branched three
times each. ‘Late adults’ had thick robust stalks and persistent green
whorls composed of hairs that branched four times each. The ‘reproductive phase of development’ includes cap initiation and expansion,
gametogenesis, and mating. Cells whose caps had reached their
maximum diameters and contained gametangia were termed ‘mature’
cells or caps.
Measurements of structures, amputation of cells, and data
analysis
Cap diameters were measured on a dissecting microscope fitted with
an ocular micrometer. An isolation-fixation solution containing 4′ 6diamidino-2-phenylindole (DAPI) was used to stain nuclei (De et al.,
1990). Plastic and glassware were sterilized with ethylene oxide
(Zeller et al., 1993). All chemicals were purchased from Sigma (St.
Louis, MO, USA).
Experimental manipulations are depicted by cartoons on the
relevant figures. Before amputation, a ‘pressure-wound’ was made at
the prospective cut site with a dental tool (small excavator spoon, CGAMERICAN Model #UW A 46, GC International Corp., Scottsdale,
AZ, USA) so as to miminize loss of cell contents. Pressure wounds
bisected the cell contents, caused them to retract away from the wound
site, and created a region devoid of both cytoplasm and vacuole
(Fester et al., 1993). The cell wall was cut with a scalpel (type 21,
Bard Parker, Division of Becton, Dickenson, and Company Ruther-
(from above)
cap
expands
nucleus (2n)
divides
early adult
whorl
of hairs
stalk
nucleus (2n)
MATERIALS AND METHODS
Cell strain and culture
Acetabularia acetabulum (L.) Silva (Chlorophyta), strain Aa0005
(Ladenburg #17), was used in all experiments. This laboratory strain
has been axenically propagated for 3 generations using all the
gametangia from >200 individuals per generation to prevent loss of
alleles. Gametangia-bearing caps were decontaminated (Hunt et al.,
1992) and then stored at 10°C in the dark until use. Following mating
(Mandoli et al., 1993), the axenic zygote stock was stored in the dark
at 10°C. Zygotes were grown at 1 cell/3 ml until they made the first
whorl (Zeller et al., 1993). Then, each cell was grown to reproductive
maturity in a polystyrene Petri dish with 20 ml of seawater at 21±1°C
under cool-white fluorescent lights (40-70 µmol photons/m2/second)
on a 14:10 hour photoperiod. One version of a novel artificial seawater
(Hunt et al., 1995) which is based chemically on a derivation of
Müller’s medium (Müller, 1962) as modified by Schweiger et al.
(1977) was used here except in some of the experiments involving
amputation (Figs 5 and 6) in which a slightly different, earlier version
of the recipe was used.
Cell developmental age
The phase of vegetative development was assessed for all cells
(Nishimura et al., 1992b). ‘Juveniles’ were threadlike, 0.1-1.0 cm tall,
=
thousands of nuclei (n) transported
late adult
x
ape tes
initia p
c
a a
juvenile
concentrated at the cell apex (Hämmerling, 1963a), and the
absence of a putative, cytosolic inhibitor of the expression of
these mRNAs (Bannwarth et al., 1991; Beth, 1953; Li-Weber
et al., 1985; Shoeman et al., 1983; Zetsche, 1966). The external
cues include the nutrient environment in which the cell grew
(Hunt et al., 1995) and probably blue light (Clauss, 1970).
Cells reproduce only once all the internal and the external cues
are met.
Vegetative development of Acetabularia acetabulum, which
must be completed prior to reproductive onset, has morphologically distinct juvenile and adult phases (see Materials and
Methods) which are temporally sequential and spatially
stacked (Nishimura et al., 1992b). In Fig. 1, the vegetative
phases are delineated on a mature or ‘late adult’ cell which is
ready to make a cap. After the cap initiates, it expands laterally
(Fig. 1). During cap expansion the diploid nucleus probably
undergoes meiosis (Green, 1973; Koop, 1975) followed by 910 rounds of mitosis (Nishimura et al., 1992a). The resulting
haploid ‘secondary’ nuclei are transported up the stalk along
with most of the parent cell contents into the cap. After nuclear
transport, the junction or ‘septum’ between the cell stalk and
the cap closes (Fig. 1, far right, center cap). The stalk is now
optically clear yet retains its shape. Gametangial cell walls
enclose each haploid nucleus with some parental cell contents
(Fig. 1, far right, bottom cap). Gametangial formation is fairly
independent in the individual partitions or gametophores of the
cap (Berger et al., 1992; Shihira-Ishikawa, 1989). Temporal
coordination between and interdependence of these developmental events in wildtype cells is not known.
Our goal was to analyze the age and size requirements for
cap initiation and to begin to assess the roles played by the
nucleus and apex in regulating reproductive onset in wildtype
cells. Our data define the spatial and temporal limits to reproductive development both in intact cells and in cells regenerating from cap amputation. The ‘developmental loop’
described by these data will be a useful experimental tool in
analyzing how reproductive onset is regulated in Acetabularia.
septum
septum
closes
nuclei (n)
gametangia
form
rhizoid
Fig. 1. A brief life cycle of A. acetabulum. A late adult cell (left)
grows through three phases in development (from bottom to top).
The vertical arrows show the direction of growth and the relative
portions of the cell length for each phase. Vegetative morphological
differences are omitted for reasons of clarity (see Materials and
Methods). The stalk of the cell is decorated with rings of hairs, or
‘whorls’, which leave scars whose spacing is characteristic of the
developmental phase (Nishimura et al., 1992b). Events in the
reproductive portion of the life cycle are, from left to right: at
reproductive onset, a vegetative cell apex differentiates a cap which
then expands and is filled with haploid nuclei (center cell). Haploid
nuclei which are of + or − mating type, are shown as black or white
dots and the diploid nucleus is grey. After packing of the haploid
nuclei in the rays of the cap (repeated image shown from above at
top right), subsequent changes in the cap are: partitioning of the stalk
from the cap when the junction, or ‘septum’ closes, partitioning of
the cytoplasm with each nucleus when ‘gametangia’ form. The
mating type of the gametes in each gametangium is not indicated.
These cartoons are not draw to scale.
Coordinating reproduction in Acetabularia 1189
100
50
?
time
0
0
25
50
75
Time, days from first whorl to amputation
Apices that made a cap, % of population
100
Early adult cells
50
0
100
Late adult cells
50
0
0
0.25
0.50
Ability of the apex to differentiate: dependence on
the nucleus
Although it was shown over 60 years ago that vegetative A.
acetabulum apices can differentiate a cap without a nucleus
(Hämmerling, 1932, 1934, 1963b), the developmental age
when the apex first becomes independent of the nucleus has
never been discerned. Cells used for amputation experiments
by previous researchers (Beth, 1953; Hämmerling, 1932, 1934)
were probably late adults (see Materials and Methods).
To see whether the ability of the apex to initiate a cap
depended on the age of the cell when the rhizoid was removed,
whole cells were bisected at different times after formation of
the first whorl leaving all of the stalk and about 90% of the cell
contents with the apex. Fig. 2 shows that apices whose rhizoids
had been amputated ≤20 days after the intact cells had made
the first whorl of hairs had <50% probability of surviving the
procedure whereas apices amputated ≥20 days after making the
first whorl had an 80-100% chance of surviving. The youngest
apices that could make a cap without the nucleus were
amputated from cells ≥35 days after formation of the first
whorl (Fig. 2, arrow), which corresponds to the early adult
portion of vegetative growth. In contrast to apices from
juvenile cells, apices from adult cells can make caps without
the nucleus as shown in Fig. 3. When about 90% of the cell
0.75
intact
Fraction of cell body left with apex
Fig. 3. Apices that made a cap versus the fraction of the cell left with
the apex. The percentages of cap initiation in 4 populations of intact,
wild-type cells are indicated by the symbols on the right. These control
populations were grown under identical conditions and were pressurewounded but not amputated either during the day (open circles) or
night (solid circles). Cells amputated during the day are indicated by
open symbols connected with a line. Cells amputated during the night
are indicated by solid symbols connected with a line. The mean
number of cells represented by data from several experiments was
14.9±0.1 except for the control cell populations (n = 60 cells total).
whorls post abortion
3.1 ± 0.4 in Aa0006
4.0 ± 0.4 in Aa0005
cap abortion
Caps in aborting cell population, #
Fig. 2. Ability of the apex to survive without the rhizoid depends on
the age of the cell. The ages of the populations (e.g. those at 25 and
20 days) at the time of amputation are approximate as the different
experimental populations were not developmentally synchronous.
Cells whose tips were amputated at 0 to 25 days after time zero were
juveniles, at 25 to 45 days were early adults, and at 45 to 60 days
were late adults (see definitions in Materials and Methods). Apices
amputated from cells at 35 days after formation of the first whorl
were the youngest to subsequently initiate caps (vertical arrow).
Each point represents from 6 to 20 cells with a mean of 15.6±1.6
cells. Intact cells made caps starting 45 days and enucleate apices
55.8±0.8 days after formation of the first whorl. Cartoon to the right
of the question mark indicates the terminal morphology of apices
that survive amputation but fail to make a cap.
RESULTS
40
Aa0005
Aa0006
30
20
10
0
0
Aborted cap dia, % of mature
Survival of apices post amputation,
percent of population
ford, NJ) perpendicular to the long axis of the stalk 5-7 seconds after
pressure-wounding.
StatView® 4.0 was used to analyze the data (Abacus Concepts,
Inc.). Standard errors of the mean are given when possible.
200
150
100
50
0
50
100
150
200
Aborted final cap diameter, % of mature cap
Fig. 4. Comparison of the diameters reached by aborted and mature
caps. Only those cells that eventually bore gametangia were used in
two wild-type, laboratory strains (Aa0005: 85 out of 98 cells,
Aa0006: 68 out of 121 cells). The average cap diameter at abortion
was 58.5± 4.1% for Aa0005 or 50.0±3.3% for Aa0006 of the mature,
gametangia-bearing cap. The average diameter of the mature
gametangia-bearing cap was 5.7±0.2 mm for Aa0005 or 4.7±0.2 mm
for Aa0006. Between successive caps, cells made an average of
4.0±0.4 whorls for Aa0005 or 3.1±0.4 whorls for Aa0006. The inset
shows the same data analyzed in a ‘box plot’ format. The five
horizontal lines in each box plot represent 10, 25, 50, 75, and 90% of
the population. The symbols at the top and bottom of each box plot
show ‘outliers’ or individual cells which are the 10% at the
population extremes.
was left with the apex, apices from late adults were 3 times
more likely to initiate a cap than apices from early adults (Fig.
3). These results suggest that for an apex to initiate a cap in
the absence of the nucleus, the cell from which it was taken
must have reached the early adult phase of developmental (Figs
2 and 3).
To establish how cell volume affected the ability of an
enucleate apex taken from adult cells to initiate a cap, early
and late adult cells were amputated at varying points along the
stalk (Fig. 3). Apices varied in length by increments of 1/8 of
a cell. The rhizoid comprised the lower or distal 1/8 of the total
cell length. Since cell height is genetically determined
(Nishimura et al., 1992b; Mandoli and Hunt, unpublished
data), relative rather than absolute length was used. Two thirds
of the intact controls (40 out of 60 cells) initiated caps (Fig. 3).
The apex itself (upper quarter of the cell) was rarely capable
of making a cap without the nucleus. Clearly, early adult apices
required more cell volume than late adult apices to make a cap
without the nucleus.
To see how cap initiation depended on the number of
chloroplasts in the apex, cell populations were amputated both
during the day, when the chloroplasts stream continously in
the stalk, and during the night, when most of the chloroplasts
stop streaming and ‘rest’ in the rhizoid (Schweiger et al.,
1964; Wollum, 1991). This circadian rhythm in chloroplast
movement results in a gross asymmetry of organelles so that
amputation of cells at night yields apices with few chloroplasts and rhizoids which are highly chloroplast enriched.
Conversely, amputation of cells during the day yields cell
pieces with fairly equal chloroplast densities. Apices
amputated from early adult cells during the day needed more
cell body to make a cap than those amputated at night (Fig. 3,
top) whereas apices amputated from late adult cells needed
less than half of the cell body to initiate a cap regardless of
time of day at which amputation occurred (Fig. 3, bottom).
Hence, the ability of an enucleate apex to make a cap was not
strictly dependent on the number of chloroplasts in the apex
(Fig. 3).
The ability to make more than 1 cap: a ‘loop’ in
reproduction
Cells can repeat the transition from vegetative to reproductive
development: a cell will cease expansion of a cap, produce
more stalk, and then initiate a new cap as diagrammed in Fig.
4. The cell contents are withdrawn from the ‘aborted’ cap back
into the stalk so that the aborted cap eventually appears white.
This ‘loop’ in development is repeated until the youngest (i.e.,
uppermost) cap formed at the cell apex completes gametogenesis. A cell that has aborted resembles a stack of 2 or more
umbrellas which are fused top to bottom (Fig. 4, top right).
Abortion occurs both in the ocean and in the laboratory.
Cells with aborted caps were isolated from wildtype laboratory strains in which they had spontaneously appeared. Cap
abortion occurred in about 5% of a population (6 out of 130
cells). In two cell lines, only 56.2 to 86.7% of the cells that
had aborted a cap made gametangia (see Fig. 4 legend). Cells
that failed to make gametangia died and were excluded from
this analysis. The diameters of aborted caps were measured and
then normalized to the diameter reached by the mature
gametangia-bearing cap eventually made by the same cell in
order to nullify genetic variation in cap diameter (Koop, 1977).
Cap diameter, % of mature final cap
1190 L. L. Runft and D. F. Mandoli
2nd cap post
amputation
150
intact
1st cap
100
50
0
0
25
50
Time, days post cap initiation
Fig. 5. Rate of cap expansion for amputated and intact cells. The
linear regression for days 0-25 was y = 9.3+4.0x (R=0.98) for the
amputated cells (mean cells per datum for days 0-25=11.2±1.2, mean
cells per datum point for days 25-45=2±1.2) and was y=11.9+3.6x
(R=0.99) for the intact cells (mean cells per datum point = 28.8±1.3).
The average diameter of the mature gametangia-bearing cap was
5.6±0.1 mm for intact cells and 6.02±0.2 mm for the second cap of
amputated cells. Data for the amputated population at days 45 and 50
are missing since the cells had formed gametangia and so could not
make a second cap. Other developmental events which occur during
this time frame are shown in Fig. 10.
Note that we could not tell how much the aborted cap expanded
after growth of the stalk recommenced because we can’t
predict when and in which cells cap abortion will occur. The
diameters of the aborted caps varied: the mean diameter of the
aborted caps was about 50% of the mature, gametangia-bearing
caps with 3-4 whorls made between successive caps (details in
Fig. 4 legend). Although cap diameter at abortion varied
widely, all cells of a strain made the same number of whorls
before reproduction was attempted again.
To estimate the time of abortion, the final diameter of the
aborted caps (Fig. 4) was correlated with the cap diameter in
cells that developed normally. For this estimate to be valid,
cap expansion must be linear and similar regardless of
whether a cell is making a first or second cap. First, as Fig. 5
illustrates, caps expanded at a linear rate until they reach their
final mature diameter about 25 days after initiation. Second,
the rate of cap expansion is similar for intact cells and the
second cap of cells whose first cap had been removed (Fig. 5:
intact, 3.6%; amputated, 3.9% of the final cap diameter per
day). Note that although 13.3% (10 out of 75) of the mature
gametangia-bearing caps were smaller in diameter than the
immature caps that had been aborted by the same cell (Fig. 4),
this does not affect the calculation of the rate of cap expansion
since the data were normalized. These data predict that cap
abortion occurred about 15 days after a cap initiated: at 3.6%
expansion of the final cap diameter per day, caps reach 55.4%
of the mature diameter 15.4 days post cap initiation (Fig. 4,
inset box plots).
Triggering the loop in reproduction with cap
amputation
To determine the limits of cells to reiterate reproductive onset,
apices of reproductive cells were amputated until all cells in
50
0
0
5
10
15
20
25
Successive apex amputations, number per cell
Fig. 6. Cells could make a cap again despite repeated amputation of
the reproductive apex. All cells made a cap after the first amputation
(first datum) but none made a cap following the 23rd amputation (n =
30 cells total). Caps were amputated on the day of initiation. Note
that the number of cells represented by each point decreases as the
cells either failed to make a new cap or to heal.
Successive cap initiations, # per cell
the population stopped initiating caps. Apices were amputated
on the day of cap initation and were discarded. All cells
initiated a second and then a third cap following amputation of
the first and second caps respectively (Fig. 6). The probability
that a cell would initiate 4 or more caps was inversely related
to the number of apices that had been previously amputated.
On average, cells made 9.0±1.0 caps after successive apex
amputations before losing the ability to reiterate reproductive
onset. One cell initiated 23 caps in succession, failing to make
a new cap only after the 23rd apex amputation. Fig. 7 shows
that new caps were initiated an average of 11 days after amputation of the previous cap. Although the intervals needed to
reinitiate a cap following amputation of caps 3 through 23
ranged from 7 to 17 days there was no pattern in the time
required: one cell might take 17 days to make the 10th cap but
only 7 days for the 11th suggesting that the amount of time
required to initiate a new cap was not a function of the amputation history of the cell but of that particular amputation.
20
10
R = 0.999
0
0
100
200
Time, cumulative days since first cap initiation
Fig. 7. Rate of cap initiations during repeated amputation of the
reproductive apex. The rate of cap initiation (y=11.0x−4.4) was
derived from the previous experiment (Fig. 5). Caps were amputated
on the day of initiation. Standard errors of the mean time between
successive caps are indicated.
Time, days from amputation to next cap
100
25
15
20
2nd cap
15
10
time
10
5
5
0
whorls
0
10
20
30
Whorls, # made post amputation
Cells that made a new cap, % of
population
Coordinating reproduction in Acetabularia 1191
0
Time, days from cap initiation to amputation
Fig. 8. Time of second cap initiation and number of intervening
whorls made between amputation of the first cap and initiation of the
second. Note that the number of cells represented by each datum point
decreased as the cells failed to make a new cap or failed to heal. For
clarity, only one-sided standard errors for both parameters are drawn.
Cell population size per datum point was 8.14±1.56 for time and
5.14±1.1 for whorls. The interval between the 2nd and 3rd cap from
Figs 5 and 6 is replotted here for comparison (solid triangle).
These data show that A. acetabulum has a remarkable capacity
to recover from wounds and can repeat reproductive onset if
the apex is lost.
We wondered whether the cells were returning to the same
earlier point in development regardless of how the cap was lost.
That is, was the loop in development the same in aborted and
amputated cells? We measured both the time from loss of the
cap to initiation of a new one and the cell structure of the stalk
made during this developmental loop. A quantitative temporal
comparison of the abortion and amputation data (Figs 4 vs. 6)
was not possible since we could not induce cap abortion but
only see that a cap had aborted retrospectively. Data in Fig. 8
show that cells whose apices were removed on the day of cap
initiation made a new cap 5 days post amputation with 1 whorl
between the amputation site and the new cap. Cells whose
apices were amputated 5-20 days post cap initiation took
longer to make a new cap and made more whorls than those
amputated on the day of cap initiation (Fig. 8). From 0-20 days
these increases are nearly linear. When apices were removed
>20 days post cap initiation, most of the cells could not make
a new cap but the few that could did so 8-11 days after amputation. The time required for healing the wounds was not
assessed. These data suggest that a cell can repeat reproductive development at almost any time during cap expansion if
the cell returns to a point in vegetative growth at least 5 days
prior to cap initiation and suggests that the amount of vegetative growth after cap amputation depends strongly on the
amount of cell contents that were removed.
Point of no return: loss of ability to repeat
reproductive onset
If there existed a point when a cell had to finish reproduction
with the last cap made and could no longer repeat reproductive onset, then cells whose apices were amputated after this
would not be able to form a new cap. As shown in Fig. 9, cell
populations were amputated at several times post cap initiation
to see if such a point exists. All cells that were amputated ≤15
cap initiation (Fig. 10) after cap expansion was complete (Fig.
5). All cells finished septum closure about 41 days post cap
initiation (Fig. 10). Gametangia first appeared in cells at 30 days
and 50% of the population of cells contained gametangia at about
40 days post cap initiation. All of the cells completed gametangia formation about 55 days post cap initiation (Fig. 10). Cells
completed cap expansion, septum closure and gametangial
formation about 1 week apart (Figs 5, 10).
100
75
50
25
?
DISCUSSION
time
0
0
20
40
Time, days from first cap initiation to amputation
Fig. 9. Ability to make a new cap declined as time from cap
initiation to apex amputation increased beyond 15 days. Mean cells
per datum point was 15.1± 0.2. The linear regression of the data from
day 15-45 was y=151.9−3.6x (R=0.987).
days after the first cap was initiated were able to make a second
cap. When amputation occurred >15 days after cap initiation
the probability that a cell would initiate a second cap decreased
by 3.6% per day. When amputation occurred ≥40 days after
cap initiation, no cells in the population could initiate a new
cap. Fifteen days post cap initiation corresponds to 54% cap
expansion, roughly the calculated mean time of abortion (Fig.
9 vs. Fig. 5). Also, when the first cap was removed 15 days
after initiation, the time required to make a second cap nearly
doubled (Fig. 8). These data suggest that cells 0-15 days post
cap initiation can enter the loop in reproduction and that loss
of the ability to make a new cap occurs as early as 15 days post
cap initiation.
Nuclear division and post meiotic events during cap
expansion
Nuclei were stained with 4′,6-diamidino-2-phenylindole
(DAPI) at various times during development to determine if
meiosis in the diploid nucleus in the basal rhizoid of the cell
was correlated with a particular point in cap expansion. Unfortunately, meiotic tetrads were not easily seen since the nucleus
seems to reside near the center of the fist-like rhizoid. However,
nuclei being transported could be visualized with DAPI (De et
al., 1990; Shihira-Ishikawa, 1984; Shihira-Ishikawa et al.,
1984). Of the cells stained with DAPI, 0% of the vegetative
cells (n=92) and 24% of reproductive cells (n=296) had visible
nuclei in the stalk. Multiple nuclei were detected in cells at any
time post cap initiation until the end of cap expansion (25 days,
Fig. 5). The highest percentage of cells with nuclei being transported up the stalk occurred 15 days post cap initiation as shown
in Fig. 10 about when the cell population begins to lose the
capacity to enter the loop in reproduction (Fig. 9). These data
argue against the idea that the loss of ability to enter the loop
in development and meiosis are concomitant (Figs 7-9).
The last two developmental events that the cell undergoes prior
to gametogenesis are septum closure and gametangial formation.
Septum closure marks the end of nuclear transport (Fig. 1).
Septum closure first occurred about 24 days after cap initiation
and 50% of the cells completed septum closure at 32 days post
Our data suggest that for A. acetabulum to reproduce it must
be old and/or big enough and establishes the limits of the cell’s
ability to reproduce when the reproductive apex aborts or is
lost. Based primarily on our data, Fig. 11 summarizes the
relative timing of the developmental events or interactions that
surround reproductive onset in A. acetabulum. This is the first
time that events preceding and during reproductive development have been quantified with the amputations feasible in A.
acetabulum.
Regulation of cap initiation: putative mRNAs and a
putative inhibitor
The ability of the apex to make a cap without the nucleus has
been attributed to the presence of a putative population of stable
mRNAs (Berger et al., 1987; Beth, 1953; Hämmerling, 1963a).
This ability is largely confined to the apex since isolated midsections of the cell rarely initiate caps (Hämmerling, 1932,
1934). Furthermore, Beth (1953) reasoned that differentiation
of the apex was inhibited by the nucleus until the cell was ready
to reproduce since the enucleate apex differentiated sooner than
it would have in the presence of the nucleus. This hypothesis
has been supported by comparison of the kinetics of cap
initiation and the activity of developmentally regulated
enzymes between intact and enucleate Acetabularia cells
(Bannwarth et al., 1977, 1982; de Groot and Schweiger, 1983,
1985). The existence of the putative mRNAs and the role and
identity of the putative cytosolic inhibitor of cap initiation
Late events, percent of population
Cells that made a new cap,
percent of population
1192 L. L. Runft and D. F. Mandoli
septum closure
100
75
gametangial
formation
50
>1 nucleus in stalk
25
0
0
20
40
60
Time, days from cap initiation
Fig. 10. Timing of three nuclear events which occur post cap
initiation. Percentage of the population which had closed the septum
or formed gametangia was plotted as a function of the time in days
post cap initiation (mean cells per datum point was 10 and 15.1±0.2
respectively). DAPI-stained nuclei of vegetative cells were early
adults and never had more than one nucleus (black square).
Coordinating reproduction in Acetabularia 1193
mRNA expression?
cap initiation
nuclear ÷ & transport, septum closure
cap expansion
gametogenesis
light?
-10
0
10
20
30 da ys
abortion loop
competence
differentiation
point of no return
Fig. 11. Summary of the reproductive phase of development in
Acetabularia acetabulum. Times indicate when an event was first
seen in a population. Cap initiation, the first identifiable event in the
reproductive phase of development, is defined as day zero on the bar
that shows time. Measurable developmental events in reproduction
are above and conceptual terms are below the time bar. Events
followed by question marks are included for the sake of
completeness but were not measured here.
(Bannwarth et al., 1991; Beth, 1953; Li-Weber et al., 1985;
Shoeman et al., 1983; Zetsche, 1966) remains hypothetical.
Coordinating linear developmental between the
nucleus and apex
Our results show that the isolated apex of a cell was competent
to differentiate a cap without the nucleus only after the cell had
reached the adult phase of development and only if there was
enough of the cell body associated with it (Figs 2, 3). Clearly,
these are not just requirements for a certain number of chloroplasts since apices from adult cells make caps when relatively
achlorotic (Fig. 3). Apices from juvenile cells could fail to
make a cap because (1) they do not survive the physical trauma
of amputation well, (2) they were not yet competent to form a
cap without the nucleus-containing rhizoid (i.e. they lack the
putative cap-specific mRNAs or are not yet independent of the
nucleus for basic vegetative, autotrophic growth), or (3)
because they take too long to make a cap (e.g. exceed the halflife of the putative, cap-specific mRNAs estimated at 20 days;
Kloppstech et al., 1982). Whether competence of the apex to
make a cap without the nucleus, acquired during early adult
growth (Fig. 3), can be correlated with accumulation of specific
mRNAs in the apex will be interesting to determine.
Once initiated, the cap expands radially at a steady rate (A.
acetabulum: Fig. 4 and Zetsche, 1966; A. calyculus: ShihiraIshikawa, 1984). Cap expansion was 3 times slower in A.
acetabulum than in A. calyculus (24 versus 8 days respectively:
Fig. 5 vs. Fig. 18 in Shihira-Ishikawa, 1984) although caps of
the two species had similar final diameters. Cap diameter
depends on both genetic (Koop, 1977) and physiological
factors (Schweiger et al., 1977).
Repetition of reproductive development: abortion
versus amputation
To make a new cap after abortion a cell repeats part of vegetative growth (Figs 4-6) that has been previously defined as
late adult on a morphological basis (Nishimura et al., 1992b).
The portion of vegetative growth which is repeated is invariant
within a given strain. Three possible causes for cap abortion
are: unfavorable environmental conditions (Fester et al., 1993),
an error in coordination between the nucleus and apex, or the
presence of a lethal mutation. Short days may induce cap
abortion (Puiseux-Dao, 1963). Abortion in cells that failed to
bear gametangia (see Fig. 4, legend) suggests the presence of
lethal mutations in some cells. Acetabularia cells that have
aborted superficially resemble mutants in yeast (cdc4 and 34:
Fong et al., 1986; Goebl et al., 1988), Aspergillus nidulans
(brlADb: Miller, 1993) and possibly Drosophila (polycomb:
Lonie et al., 1994): in each case reproductive structures are
reiterated several times, stacked one above another. Whether
spontaneous abortion in A. acetabulum also has a genetic basis
is not known.
Although apex loss via abortion and amputation both trigger
initiation of a new cap, the developmental pattern that ensues
is not the same. During abortion the pattern of development
does not vary, there is no wound to heal, and all the cytoplasm
is recovered from the old cap. In contrast, when a cap is physically removed, cells make an increasing number of whorls as
the interval between cap initiation and cap amputation
increased (Fig. 8) and later amputations remove an increasing
proportion of the cell body as more of the parental cell contents
are moved into the expanding cap. (Note that the apparently
faster cap initiation and fewer whorls made in this experiment
(Fig. 8) should not be overinterpreted because the population
was small by this time.) In sum, although cap abortion or
amputation both trigger cells to repeat reproductive onset, it is
not clear how these phenomena are related.
Point of no return
Cells cannot enter the loop in reproductive development after the
cap reaches a certain age, when it must finish reproduction or die
(Fig. 9). We avoid the term ‘commitment’ because our assay is
a loss of function, inability to make a new cap post amputation,
rather than a gain of function or commitment to a defined event
such as the beginning of cell division, called START (Hartwell
et al., 1989). Chromosome behavior visualized with DAPI
staining in A. calyculus suggests that premeiotic chromosome
condensation started 3 days after cap initiation and finished in the
cell population when the cap was about three quarters of its final
diameter (Shihira-Ishikawa, 1984) but these events have not been
documented well in A. acetabulum. The relationship between
meiosis and the point of no return is not obvious.
These data contribute to our understanding of which cellular
activities characterize the different developmental ages of
Acetabularia acetabulum and describe the limits of the ability
of the organism to control reproductive onset. How the apex
and the nucleus of this organism work together to coordinate
progress through the phases in vegetative development and to
correct errors in the onset of reproduction will be intriquing
puzzles to unravel.
The authors contributed equally to this work. This research was
supported by a Hughes Undergraduate Research Fellowship (L. L.
R.), in part by a Project R/B-8 of the Washington Sea Grant Program
with funding from Grant no. NA89AA-D-SG022 from the National
Oceanic and Atmospheric Administration, U.S. Department of
Commerce (D. F. M.), and in part by the National Science Founda-
1194 L. L. Runft and D. F. Mandoli
tion #IBN-9305473 (D. F. M.). We thank Peter Ray, Paul Green, Lee
Hartwell, and Frank Russo for stimulating discussions and Susan
Singer and Winslow Briggs for comments on the manuscript.
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(Accepted 21 December 1995)