1 Verna and Marrs McLean Department of Biochemistry and Molecular Biology,
Baylor College of Medicine, One Baylor Plaza, Houston, TX 77030, USA
2 Department of Molecular and Human Genetics, Baylor College of Medicine, One
Baylor Plaza, Houston, TX 77030, USA
* Author for correspondence (e-mail: akuspa{at}bcm.tmc.edu)
Accepted 1 March 2004
![]() |
SUMMARY |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Key words: Flow cytometry, Bromo-deoxyuridine, G1 arrest, Differentiation
![]() |
Introduction |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Dictyostelium cells must transition out of the growth phase cell
cycle when they encounter starvation conditions and initiate multicellular
development (Kessin, 2001).
The relationship between cell fate determination and cell cycle regulation in
Dictyostelium has been a topic of intense study over the past 30
years. It has been well established that the cell cycle status of growing
cells impinges on cell fate decisions after development is initiated. Cells
that are in middle, or late, G2 phase at the time of starvation preferentially
become prespore cells, whereas cells in the S, M or early G2 phase
preferentially become prestalk cells (Azhar
et al., 2001
; Gomer and
Ammann, 1996
; Gomer and
Firtel, 1987
; McDonald and
Durston, 1984
; Ohmori and
Maeda, 1987
; Weijer et al.,
1984a
; Zimmerman and Weijer,
1993
). Maeda and colleagues have proposed that cells exit the cell
cycle at a particular point late in the G2 phase called the `putative shift'
(PS) point and those cells that must traverse G2 under starvation conditions
for the longest time in order to arrive at the PS point will have a propensity
to differentiate as prestalk cells (Maeda
et al., 1989
; Maeda,
1993
; Araki et al.,
1997
; Maeda,
1997
). Alternatively, it has been suggested that cells either exit
the cell cycle early in G2, biasing them to differentiate as prestalk cells,
or cells exit the cell cycle late in G2, biasing them to differentiate as
prespore cells (MacWilliams et al.,
2001
). There is general agreement that all cells pause in G2
during the first half of development.
It is important to understand the regulation of the cell cycle during
development so that we can critically assess its role in cell type
specification and terminal differentiation. The cell cycle status of
developing cells after their initial pause in the G2 phase is unclear because
data from different experiments have led to contradictory conclusions. Some
cell division takes place in the first 6 hours, prior to aggregation, and
additional cell divisions occur between 12 and 20 hours, after multicellular
structures are formed (Atryzek,
1976; Zada-Hames and Ashworth,
1978a
). The currently accepted model posits that all cells that
divide during development replicate their chromosomes after mitosis and
undergo terminal differentiation as G2 cells
(Weeks and Weijer, 1994
). Flow
cytometry studies have suggested that all the developmental cells have the
same nuclear DNA content as G2 vegetative cells, implying that developmental
cells are in the G2 phase (Durston et al.,
1984
; Weijer et al.,
1984a
). Some reports of DNA synthesis at the time of cell division
appear to support these claims (Zada-Hames
and Ashworth, 1978a
; Zimmerman
and Weijer, 1993
). However, this model has been challenged by
experiments showing that the bulk amount of DNA synthesis during development
cannot account for complete replication of the chromosomes in more than a few
percent of the cells. In fact, the majority of DNA synthesis during
development appears to be accounted for by mitochondrial DNA (mtDNA) synthesis
(Shaulsky and Loomis, 1995
).
Direct measurements of cellular DNA content also suggests that spores have
about half the amount of DNA as vegetative (mainly G2 phase) cells
(Sharpe and Watts, 1984
).
These results argue that developmental cells remain in G1 phase after
mitosis.
In order to begin to explore the possible relationships between cell cycle regulation, cell-fate determination and terminal differentiation during Dictyostelium development, we need an accurate description of the cell cycle during development. We have revisited the question of cell cycle progression during development in an attempt to clarify basic issues of cell cycle timing in relation to cell differentiation events. We provide evidence for tissue-specific regulation that results in G1 phase cell cycle arrest in all cells prior to terminal differentiation.
![]() |
Materials and methods |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Quantification of cells and nuclei
Two filters of developing cells were processed for each time-point as
described previously (Zada-Hames and
Ashworth, 1978a). The cellulose pad was removed and the filter was
placed in its original petri dish and flooded with 3 ml 50 mM Tricine (pH 7.4)
containing 1 mg/ml Pronase (Protease type VI, Sigma) and 0.25%
2,3-dimercaptopropanol (BAL). The cells were washed off the filter by repeated
pipetting for about 3 minutes and the cell suspension was transferred to a 50
ml polypropylene tube. Additional BAL/pronase solution (3 ml) was used to wash
the filter and dish again and the cell suspension and the filter were added to
the tube. Finally, the Petri dish was washed with 10 ml 50 mM tricine buffer
and this wash was added to the tube. The cell suspension and filter were
vortexed vigorously in the tube for 30 seconds. The suspension was brought to
49.5 ml with saline, 0.5 ml of 0.5 M EDTA (pH 8.0) was added, and the tube was
then shaken for 30 seconds and vortexed vigorously for an additional 60
seconds. Two determinations of 100-200 cells were made for each of four
aliquots from each of two independently prepared tubes. The cells from two
tubes were then sedimented (500 g, 5 minutes), combined and
resuspended in PDF containing 2% sucrose in saline solution.
To quantify nuclei in the cell samples, ethanol-fixed cells (5x105) cells were pelleted and resuspended in 100 µl 2% sucrose, 0.9% NaCl, in PDF. Cells were sedimented onto a microscope slide, 9 µl of Vectashield (H-1000, Vector Laboratories) containing DAPI was applied to the cells and the number of nuclei within individual cells was quantified by fluorescence microscopy.
Flow cytometry
Flow cytometry was carried out as described
(Dien et al., 1994). Cells
(1x107) were resuspended in 1.5 ml 0.9% NaCl, 2% sucrose, 5
mM EDTA, in PDF buffer and fixed by adding 5 ml 90% ice-cold ethanol dropwise,
with gentle vortexing. The cells were then incubated for >30 minutes at
22°C and stored at 4°C. For staining with propidium iodide (PI) just
prior to flow cytometry, cells (1x106) were pelleted from the
fixation solution and resuspended in 1 ml of PI (50 µg/ml in
phosphate-buffered saline, PBS). Thirty minutes before analysis, 1 µl of
100 mg/ml DNase-free RNase (Sigma, St Louis, MO) was added and the suspension
incubated at 37°C. Samples were analyzed on a Beckman-Coulter Epics XL-MCL
apparatus. The parameters were adjusted for the measurement of single cells
using the forward and side scatter plots as guides.
The DNA in developing cells or spore DNA might be less accessible to PI
using staining procedures developed for vegetative cells and this might
distort the flow cytometry profiles. We therefore treated samples with 70%
formamide at 85°C to denature the DNA prior to flow cytometry
(Baerlocher et al., 2002). The
flow cytometry profiles were very similar between these treated cells and
untreated cells.
Separation of cell types to measure DNA content
Slug dissection
ecmA/GFP-marked AX4 cells were washed with DB buffer (5
mM Na2HPO4, 5 mM KH2PO4, 1 mM
CaCl2, 2 mM MgCl2, pH 6.5) and resuspended in water to
109 cells/ml. Cells were streaked on a 6 cm line (100 µl)
on water agar plates made with 2% Noble agar (Difco) and allowed to develop in
a dark chamber with unidirectional illumination. After 24-36 hours, slugs were
dissected with a tungsten wire to harvest GFP-labeled slug anteriors and
unlabeled slug posteriors. At least 50 anterior (or posterior) slug sections
were put into 100 µl 2% Sucrose, 0.9% NaCl, in PDF. Pronase (1 mg/ml) and
BAL (1 µl) were added to dissociate the cells by vortexing for 30 seconds.
The cells were harvested by centrifugation, resuspended in 0.5 ml 2% sucrose
in PDF, an equal volume of the fixation solution was added (4%
paraformaldehyde, 30% picric acid, 10 mM PIPES, pH 6.5) and cells were mixed
gently for 15 minutes. Cells were then harvested and resuspended in 0.2 ml of
2% sucrose in PDF and stored at 4°C. Stored cells were later prepared for
flow cytometry, as described above.
DNA content of cell types during development
Strains AX4[cotB/GFP] and AX4[ecmA/GFP] were grown in
HL-5 and prepared for development on filters. Cells from one-quarter of a
filter (1.25x107 cells) were harvested into a 15 ml
conical tube and dissociated by vortexing in 4 ml of BAL/pronase, as described
above. EDTA was added (0.2 ml, 0.5 M, pH 8) and the tubes were stored on ice
prior to sorting with an Altra cell sorter (Beckman-Coulter) as described
(Gerald et al., 2001
). At
least 5x105 sorted cells from each strain were collected in 3
ml and 300 µl of 20% sucrose, 9% NaCl, in PDF were added, followed by the
addition of 10 ml of 95% ethanol for fixation. Cells were stored at 4°C
prior to measuring their DNA content by flow cytometry, as described
above.
DNA synthesis
Bromo-deoxyuridine (BrdU) incorporation was carried out as follows. Cells
pellets form growth cultures were washed at 22°C in PDF supplemented with
0.5 mM BrdU and resuspended in PDF with 0.5 mM BrdU at
1x108/ml. Cells (0.5x108) were spread on one
filter with the underpad soaked with 1.5 ml 0.5 mM BrdU in PDF. After 36 hours
of development, cells from one filter were harvested and used to make one 150
µl agarose block. Agarose blocks were processed to produce high molecular
weight DNA as described, except that cellulase and hemicellulase were used to
digest the cell walls of stalk cells and spores
(Kuspa et al., 1992). Pulsed
field gel electrophoresis and Southern transfer of the DNA to nylon filters
was carried out as described (Shaulsky and
Loomis, 1995
; Vollrath and
Davis, 1987
). Detection of BrdU was carried out with anti-BrdU
antibodies conjugated to peroxidase using an ECL detection kit (Amersham).
Multiple exposures of each autoradiograph were used to assess linearity of the
signals and quantification was carried out using standard image analysis
software. Values for BrdU incorporation were normalized to the total DNA
content in each lane using the Southern hybridization signal from a
single-copy gene (gdtB) as an estimate the amount of chromosomal DNA
on the blots. The gdtB hybridization signal also showed that <10%
of the chromosomal DNA remained in the wells for any given sample. By
comparison with the signal obtained from growing cells, we estimate that we
could detect the labeling of >1% of any cellular DNA species.
Estimating the proportions of DNA species
Cells collected from each developmental time point (5x107)
were used to make two agarose blocks. The DNA in one block was digested with
SmaI and the DNAs were resolved by pulsed-field gel electrophoresis.
Gels were stained with ethidium bromide and multiple images were acquired at
325 nm at different light intensities and under-saturated images were used for
quantification. The gel was shaken in water for at least 24 hours, and each
side of the gel was exposed to UV light for 60 seconds to break DNA
(Stratalinker UV lamp, Stratagene). DNA was transferred onto nylon membranes
and hybridized with radiolabeled (32P) probes
(Vollrath and Davis, 1987).
The mtDNA probe was a PCR-amplified product from nucleotides 9991 to 13579,
which corresponds to a 3.6 kb EcoRI fragment of the mtDNA produced
with primers Mito_5' (agt tta gac act gct gg) and Mito_3' (cta aaa
cgc aca cct tct c) (Ogawa et al.,
2000
). The rDNA probe was a collection of 12 DNA fragments
encompassing the rRNA-coding region
(Sucgang et al., 2003
). The
probe for chromosomal DNA was derived from the DIRS element subclone pAK162
(Kuspa and Loomis, 1996
). The
32P signal was quantified by phosphoimaging using a Molecular
Dynamics Storm system. The relative amount of rDNA or mtDNA in the gel
(Gr or Gm, respectively) and in the wells (Wr
and Wm) was determined for each sample. The images of ethidium
bromide-stained gels were used to obtain the intensity of rDNA (Ir)
and mitochondrial DNA (Im) bands and the total DNA in each lane
(including the DNA in the wells, It). To estimate the percentage of
rDNA and mtDNA relative to total DNA, the fluorescence intensity of the rDNA
and mtDNA bands in the gel were corrected by the proportion of those DNAs
found in the wells by hybridization. Thus, the percentage of rDNA
(Pr) in a sample was estimated as:
Pr=[Ir(1+Wr/Gr)]/It,
the percentage of mtDNA (Pm) was
Pm=[Im(1+Wm/Gm)]/It and
the percentage of chromosomal DNA (Pc) was estimated by the
difference: Pc=1(Pr+Pm).
Predicting flow cytometry profiles
We predicted the DNA content expected of populations of developing cells
using two models of cell cycle control. We assumed that all cells are in G2 at
6 hours of development, that the mean DNA content is determined by flow
cytometry (F6) and that the proportion of the DNA species is defined as:
Pc6 + Pr6 + Pm6=1. The chromosomal DNA
content at 6 hours would then be: F6xPc6. The proportions of
mtDNA and rDNA were taken from the experiments described above. The proportion
of each DNA was also estimated at time `x' as: Pcx + Prx
+ Pmx=1. If all cells were in G2 at time x, the cellular
chromosomal DNA in mononuclear cells should be the same as that of 6-hour
cells (F6xPc6=FxxPcx), and the relative DNA
content would be predicted as: Fx=(F6xPc6)/Pcx. If
all cells were in G1 phase at time `y', the cellular chromosomal DNA in
mononuclear cells should be one-half that of 6-hour-old cells,
F6xPc6=2xFyPcy. The relative DNA content
would be predicted as: Fy=(F6xPc6)/2Pcy. If the
population was a mixture of G1 and G2 cells at a time `n', assuming all cells
have the same proportion of DNA species, the DNA content of G1 and G2
population was estimated as F1n=(F6xPc6)/2P1cn for
G1 cells and F2n=(F6xPc6)/P2cn for G2 cells. The
average DNA content (Fxavg) was then calculated according to the
percentage of G1 and G2 cells in the population (a1 and a2) given that
a1+a2=1. The average DNA content for the population was predicted to be
Fxavg=a1[(F6xPc6)/2P1cn] +
a2[(F6xPc6)/P2cn]. The results reported are based
on calculations made from the results described in
Fig. 2C and estimates of the
DNA content by extrapolation of the flow cytometry measurements in
Fig. 2D.
|
Fluorescence in situ hybridization (FISH)
FISH was used to assess chromosome copy number. The hybridization, DAPI
staining and image analysis was carried out as described previously
(Sucgang et al., 2003). A 7 kb
genomic fragment of the tagB gene, that hybridizes to tagB
and to two highly similar genes (tagC and tagD) that are
tightly linked within a 20 kb region of chromosome 4 was used as the probe
(Shaulsky et al., 1995
). After
hybridization and DAPI staining, preparations were visualized on a Delta
Vision deconvolution microscope and images were processed with the SoftWoRx
(version 2.5) software package (Applied Precision, Issaquah, WA). Nuclei were
scored as having one or two hybridization signals by manually inspection while
traversing through the three-dimensional images. Within each nucleus that was
scored, fluorescent signals that were approximately fivefold lower in
intensity than the more intense signal were not counted as a chromosomal
locus. Over 100 cells from each of several preparations for each biological
sample were scored and the results were averaged.
![]() |
Results |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
|
Cellular DNA content during development
The mitotic activity within aggregates means that some cells divide and
either arrest in G1 phase or they undergo a round of DNA synthesis and return
to G2 phase. If 50% of the cells divide and arrest in G1, a decrease in the
cellular DNA content should be detectable between 12 and 18 hours of
development. We used flow cytometry of whole cells to monitor changes in
cellular DNA content during development. In principle, it would have been best
to measure the DNA content by flow cytometry of nuclei, but we found this
approach problematic. We carried out flow cytometry analyses on nuclei with
several different nuclear purification and fixation protocols. We could
reproduce published profiles, namely unimodal peaks that do not change during
development, but we observed that nuclear fragility varies at different times
in development for each of the procedures that we evaluated and this caused
erratic and aberrant DNA content profiles (G.C., unpublished). The
side-scatter and the forward-scatter profiles did not show a unique nuclear
population using any preparation procedure. Instead, doublets and triplets of
nuclei and cellular debris were the predominant particles and the samples
invariably clogged the cytometer. As nuclear profiles were unreliable, we
resorted to whole-cell flow cytometry and combined it with estimates of the
changes in the extra-chromosomal DNAs to arrive at the likely cell cycle
status of the cells.
Vegetative Dictyostelium cells displayed a broad peak of DNA content that is probably due to variable mtDNA content and the presence of a small number of G1 cells (Fig. 1C). We also observed a smaller peak with twice the DNA content of the first peak, which we confirmed to be binucleated cells by direct microscopic examination (data not shown). These binucleated cells were no longer detectable as a second peak after 6 hours. For simplicity, we summarize the DNA content profiles using the average DNA fluorescence of mononucleated cells (Fig. 1D). In the first 10 hours of development there was a steady decrease in the average cellular DNA content. As we did not observe mitosis during this time, the DNA decrease is probably due to a decrease in the proportion of extra-chromosomal DNA (see below). The average cellular DNA content appeared to plateau from 10-13 hours of development and then decreased between 13 and 18 hours of development. The final spore and stalk cell DNA content is about 50% that of 6 hour cells (Fig. 1C,D). We consistently observed this pattern of cellular DNA loss during development in over a dozen similar experiments.
The decrease in cellular DNA content after 13 hours might be caused either by a shift of most cells from G2 into G1, or by the loss of extra-chromosomal DNA with chromosomal DNA maintained at constant levels by new synthesis. To distinguish between these possibilities, we estimated changes in the proportions of the three major cellular DNAs at different stages of development. We isolated high molecular weight DNA from cells at different times, separated the rDNA, mtDNA and chromosomal DNA by pulsed-field gel electrophoresis and directly quantified the DNAs based on their ethidium bromide staining (Fig. 2A). We then used Southern analyses to determine the extent of DNA trapping in the sample well for each DNA species, in order to correct these estimates (e.g. Fig. 2B and data not shown). We found that the proportion of rDNA relative to total cellular DNA does not change significantly during development, whereas the proportion of mtDNA decreases by about 50% during the first 13 hours of development (Fig. 2C). Significantly, the ratios of the three cellular DNAs did not change during the time of mitosis in the second half of development (Fig. 2C).
The DNA ratios in Fig. 2C and the cell number increase described in Fig. 1A were used to reconstruct the cellular DNA content of cell populations at each time point during development using two different assumptions of chromosome copy number after 12 hours of development. We assumed that all cells are in G2 at 6 hours of development and that after cells divide they either arrest in G1 phase, or they undergo S phase and arrest in G2 phase (see Materials and methods). Our measurements agree with the model that assumes a G1 arrest of post mitotic cells after 12 hours of development (Fig. 2D).
Quantification of DNA synthesis during development
We monitored the synthesis of chromosomal DNA, rDNA and mtDNA by BrdU
incorporation. Cells were deposited on filters supplemented with BrdU and high
molecular weight DNA was prepared after 28 hours of development. Stalk cells,
spores and non-stalk cells (all cells remaining after stalks cells were
removed, including spores) were also purified after BrdU labeling. We observed
little incorporation of BrdU into the chromosomal DNA or rDNA of developing
cells, but there was substantial incorporation into the mtDNA of spores
(Fig. 3). We estimate that our
limit of BrdU detection corresponds to the synthesis of about 1% of the
chromosomal DNA loaded on the gels, based on direct comparisons with the BrdU
incorporated into the DNA of growing cells during one cell doubling
(Fig. 3A). The small amount of
chromosomal BrdU incorporation observed in the `non-stalk cell' sample which
contains spores (6.3±2% of the growth control) compared with the much
lower amount in the purified spore sample (2.2±0.6%) suggests that most
of the chromosomal DNA synthesis occurs in cells that do not undergo terminal
differentiation into spores or stalk cells. The small amount of BrdU
incorporation into the chromosomes of spores or stalk cells indicates that the
major terminally differentiated cell types do not undergo S phase at anytime
during development.
|
|
FISH analysis of chromosome copy number in amoebae and spores
To verify that spores encapsulate as G1 cells by an independent method, we
directly compared the chromosome copy number of amoebae and spores by
fluorescence in situ hybridization (FISH). A single copy locus can be reliably
detected in Dictyostelium by FISH if the probe is complimentary to
greater than 10 kb of the target locus (G.C., unpublished). For this test, we
used a probe corresponding to the 20 kb tagB/C/D locus on chromosome
4. We examined vegetative cells and 6 hour developing cells that are expected
to be in the G2 phase of the cell cycle. We also inferred the chromosome copy
number in spores by carrying out FISH on amoebae that had just emerged from
germinating spores. By allowing spores to germinate in growth media for 12
hours, we obtained robust FISH signals and we could examine their nuclei well
before any chromosomal DNA replication had occurred. Most vegetative cells and
6 hour developing cells had two tagB/C/D loci
(Fig. 5A and data not shown).
We also examined several independent spore samples and found that more than
90% of the emergent amoebae displayed a single tagB/C/D locus
(Fig. 5B). Although detection
of chromosome copy number by FISH is complicated by variable signal intensity
and the possibility of overlapping signals, these results appear to be in
general agreement with the experiments described above. The FISH indicates
that vegetative cells are mainly in G2, as expected, while spores are in
G1.
|
We first compared the DNA content of prestalk and prespore cells in migrating slugs. We dissected slugs marked with a prestalk ecmA/GFP reporter into two cell populations: slug anterior tips that contain mostly prestalk cells and slug posterior ends that contain mostly prespore cells (Fig. 6A). The posterior cells gave a uniform DNA profile and lacked significant numbers of GFP-positive cells (Fig. 6B,C). The anterior cells had a DNA profile suggestive of a bimodal distribution of DNA content among the population. Although the fixation procedure reduced the GFP fluorescence about 100-fold, the remaining 10-fold difference in green fluorescence between the remaining GFP-positive cells and the GFP-negative cells allowed their separation. The flow cytometry window used to segregate GFP-expressing cells was shown to identify populations that contained >95% GFP-positive cells by visual confirmation of fluorescence in the sorted cell populations (Fig. 6C and data not shown). The GFP-positive prestalk cells had significantly higher DNA content than the GFP-negative prespore cells, suggesting that prespore cells undergo mitosis before the slug stage and that prestalk cells divide after the slug stage (Fig. 6D).
|
|
![]() |
Discussion |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
We have observed two stages of cell division during Dictyostelium
development. In the first stage, between 0 and 6 hours, some cells in the
population appear to be completing their final growth-stage cell cycle, as we
could attribute most of the early cell division to the cytokinesis of
multinucleated cells. Multinucleated cells must result from an imperfect
cytokinesis mechanism, a well-known property of cells grown in nutrient broth.
In the first 12 hours of development, no significant change in the number of
nuclei occurs and cells appear to be arrested in G2 based on their DNA
content. Actual mitosis appears to occur between 12 and 18 hours of
development because about half of the cells carryout karyokinesis and
cytokinesis during this period. These issues have been examined in detail for
other axenic strains and our data are consistent with these reports
(Sharpe et al., 1984;
Zada-Hames and Ashworth,
1978a
; Zada-Hames and
Ashworth, 1978b
). The cell division that we observe for AX4
strains is similar to that reported for AX2 and AX3
(Atryzek, 1976
;
Zada-Hames and Ashworth,
1978a
).
We also observe two time periods where the DNA content of cells decreases
during development. The first decrease, during the first 10 hours of
development, appears to be due to loss of about half of the mtDNA. The second
decrease coincides with the period of mitosis and is almost entirely accounted
for by the reduction of DNA content in prespore cells carrying out mitosis.
Previous reports suggesting that DNA loss after 10 hours of development is due
to loss of mtDNA were based on the differences between whole-cell and nuclear
flow cytometry, or by direct measurements
(Durston et al., 1984;
Leach and Ashworth, 1972
). Our
experiments suggest that mtDNA constitutes
30% of the total DNA in
vegetative cells and this decreases to
15% by 10 hours of development,
after which it remains relatively constant. These results are consistent with
previous reports that mtDNA constitutes 25-42% of vegetative cell DNA
(Firtel and Bonner, 1972
;
Soll et al., 1976
;
Sussman and Rayner, 1971
;
Weijer et al., 1984b
). Because
our data also indicate that only prespore cells divide before 20 hours, the
45% decrease in DNA content in these cells cannot be accounted for simply
by the loss of extrachromosomal DNA relative to chromosomal DNA because that
would predict that spores are devoid of extra-chromosomal DNA. Furthermore,
unchanging proportions of the major DNAs between 12 and 18 hours of
development supports the idea that there is a proportionate reduction of all
the cellular DNAs within prespore cells. The FISH analyses with a single-copy
probe also supports the notion that spores have one copy of the chromosomes.
These results are consistent with a previous report indicating that spores
have 40% less DNA than 10-hour developing cells and supports our assertion
that spores encapsulate as G1-arrested cells
(Sharpe et al., 1984
).
Earlier models suggesting that cells undergo terminal differentiation in
the G2 phase are based on flow cytometry that indicated that the nuclei from
growing cells have the same DNA content as the nuclei from developing cells
(Durston et al., 1984). In
principle, nuclear flow cytometry is the best means of measuring chromosome
content, but technical issues confound this approach in
Dictyostelium. We carried out flow cytometry analyses on nuclei with
several different protocols and we could reproduce published profiles, namely
unimodal peaks that do not change during development (G.C., unpublished).
However, it has never been demonstrated that G1 nuclei are as stable to
purification as G2 nuclei and there is no way of telling if there is
differential recovery of the two types of nuclei using the various protocols.
This theoretical concern was made tenable by our finding that nuclear
fragility varies at different times in development. We believe this fragility,
combined with a large amount of debris in the preparations, caused erratic and
aberrant DNA content profiles. Excessive debris in nuclear preparations has
been reported previously to obscure the detection of G1 nuclei in flow
cytometry profiles (Durston et al.,
1984
). Thus, the sample heterogeneity of nuclear preparations
suggests that flow cytometry of nuclei is subject to artifact and is therefore
unreliable.
Our estimates of chromosomal DNA synthesis further support the notion of
post-mitotic G1-arrest. These BrdU incorporation studies could be misleading
only if BrdU becomes inaccessible for chromosomal DNA synthesis during
development. The robust mtDNA synthesis observed in prespore cells suggests
that cells do have access to sufficient BrdU during development, as noted
previously (Shaulsky and Loomis,
1995). However, we cannot exclude the possibility that mtDNA
synthesis uses a different nucleotide pool and that BrdU is somehow excluded
from the nuclear pool. We consider this possibility highly unlikely as BrdU
can clearly be detected in developing nuclei by immunochemistry. Nuclear DNA
synthesis has been observed in developing BrdU-labeled cells and in cells
labeled with [3H]-thymidine
(Durston and Vork, 1978
;
McDonald and Durston, 1984
;
Zada-Hames and Ashworth,
1978a
; Zimmerman and Weijer,
1993
). These studies have been interpreted as evidence of an S
phase late in development, but these techniques do not allow quantification of
chromosome synthesis. It is likely that the observed labeling is due to DNA
repair. It is also possible that mtDNA synthesis in perinuclear mitochondria
has been mistaken for nuclear DNA synthesis, as pointed out previously
(Durston et al., 1984
;
Shaulsky and Loomis,
1995
).
Our measurements of DNA synthesis suggest that 5% of the cells replicate
all of their chromosomal DNA, all cells repair 5% of their chromosomal DNA, or
a combination of these two extremes. Chromosomal replication may occur in
cells that become neither spore or stalk cells. Our `non-stalk cell' sample
consists of 95% spores and 5% non-spore cells. Six percent of the chromosomal
DNA is synthesized in that sample, whereas only 2% of comes from spore
chromosome synthesis. So it is possible that the non-spore cells complete an S
phase, as they would have to account for roughly 4% of the BrdU incorporation
into the chromosomes of that sample. These cells could be the cells that
remain scattered outside of aggregates that do not participate in development,
cells that form minority cell types, or cells that are in S phase at the start
of development. In support of the latter, we detect much less BrdU
incorporation into chromosomal DNA when we begin labeling cells after 4 hours
of development (A.K. and G.S., unpublished). We can also estimate that
prespore cells synthesize about one-third of the mtDNA synthesized during one
vegetative cell cycle. The prespore expression of the small subunit
ribonucleotide reductase gene, rnrB, is consistent with an increased
nucleotide requirement to support mtDNA synthesis
(Tsang et al., 1996).
It has been shown that cell cycle position at the time of starvation biases
cell fate determination in Dictyostelium. Several studies have
suggested that cells starved late in G2 phase become prespore cells, and cells
starved early in G2, M or S phase become prestalk cells
(McDonald and Durston, 1984;
Weijer et al., 1984a
;
Ohmori and Maeda, 1987
;
Gomer and Firtel, 1987
;
Maeda et al., 1989
;
Zimmerman and Weijer, 1993
;
Wood et al., 1996
). Cells that
exit the cell cycle at different times in could be biased towards a particular
cell fate by known cell-cycle regulated signals such as Ca2+, or
cytosolic pH (Clay et al.,
1995
; Gross et al.,
1983
; Leach et al.,
1973
; Maeda and Maeda,
1974
; Saran et al.,
1994
; Thompson and Kay,
2000
; MacWilliams et al.,
2001
). It is tempting to speculate that some condition imposed by
the initial arrest impinges on the regulation of tissue specific mitosis later
in development. For example, it might be that transit through G2 is merely
slowed by the starvation conditions of early development and that prespore
cells, which arise from cells starved later in G2, are able to achieve the
competence to divide much earlier than prestalk cells. The slow increase in
cdc2/cyclin B kinase activity during aggregation, followed by a decline after
aggregation supports this idea (Luo et
al., 1995
). Distinguishing between this and other models will
require additional molecular tools such as temperature-sensitive mutations in
known cell cycle regulators.
The elucidation of the regulation of developmental mitosis will probably
illuminate our understanding of cell type specification and maintenance. It
will be important to determine whether the late mitotic events and G1-phase
arrest are required for spore cell differentiation or stalk formation.
Certainly, prespore-specific genes and genes that become prespore enriched are
expressed in some cells well before mitosis begins so it is unlikely that G1
arrest is required for all aspects of prespore differentiation
(Fosnaugh and Loomis, 1993;
Good et al., 2003
;
Haberstroh and Firtel, 1990
;
Iranfar et al., 2001
;
VanDriessche et al., 2002
;
Williams et al., 1989
).
Microtubule destabilizing drugs that block mitosis do not block development
suggesting that prespore cells need not divide prior to spore differentiation
(Cappuccinelli and Ashworth,
1976
; Cappuccinelli et al.,
1979
). However, some mutants that are blocked in development prior
to the multicellular phase do not undergo mitosis
(Chen, 2003
;
Zada-Hames and Ashworth,
1978a
). In addition, dominant-negative cyclin B mutants that block
the cell cycle in mitosis also block spore production (Q. Luo, PhD Thesis, The
University of British Columbia, 1996) (Luo
et al., 1994
). However, these mutants also block the production of
prespore cells, as has been pointed out for the cyclin B mutant
(Weeks and Weijer, 1994
).
These genetic tests provide some evidence that mitosis is under control of the
developmental program, but they leave unanswered the question of whether
mitosis is strictly required for terminal cell differentiation.
If G1 arrest is not required for spore differentiation, how does
Dictyostelium benefit by having prespore cells divide in the
post-aggregative stage of development? For Dictyostelium, long-term
survival is determined by the stochastic process of the spores of a fruiting
body arriving at new, nutrient-rich environments
(Bonner, 1982). In this regard,
the selective advantage of doubling the number of spore producing cells within
each multicellular organism is likely to be greater than the potential
disadvantage of spores encapsulating with only one copy of the genome.
![]() |
ACKNOWLEDGMENTS |
---|
![]() |
REFERENCES |
---|
![]() ![]() ![]() ![]() ![]() ![]() ![]() |
---|
Araki, T., Abe, T., Williams, J. G. and Maeda, Y. (1997). Symmetry breaking in Dictyostelium morphogenesis: Evidence that a combination of cell cycle stage and positional information dictates cell fate. Dev. Biol. 192,645 -648.[CrossRef][Medline]
Atryzek, V. (1976). Alteration in timing of cell differentiation resulting from cell interactions during development of the cellular slime mold, Dictyostelium discoideum. Dev. Biol. 50,489 -501.[Medline]
Azhar, M., Kennady, P. K., Pande, G., Espiritu, M., Holloman, W., Brazill, D., Gomer, R. H. and Nanjundiah, V. (2001). Cell cycle phase, cellular Ca2+ and development in Dictyostelium discoideum. Int. J. Dev. Biol. 45,405 -414.[Medline]
Baerlocher, G. M., Mak, J., Tien, T. and Lansdorp, P. M. (2002). Telomere length measurement by fluorescence in situ hybridization and flow cytometry: tips and pitfalls. Cytometry 47,89 -99.[CrossRef][Medline]
Bonner, J. T. (1982). Evolutionary strategies and developmental constraints in the cellular slime molds. Am. Naturalist 119,530 -552.[CrossRef]
Cappuccinelli, P. and Ashworth, J. (1976). The effect of inhibitors of microtubule and microfilament function on the cellular slime mould Dictyostelium discoideum. Exp. Cell Res. 103,387 -393.[Medline]
Cappuccinelli, P., Fighetti, M. and Rubino, S. (1979). Differentiation without mitosis in Dictyostelium discoideum. Cell Differ. 8, 243-252.[CrossRef][Medline]
Chen, G. (2003). The growth-development transition and tissue-specific cell cycle control in Dictyostelium discoideum. In Biochemistry and Molecular Biology, pp. 226. Houston, TX: Baylor College of Medicine.
Chen-Kiang, S. (2003). Cell-cycle control of plasma cell differentiation and tumorigenesis. Immunol. Rev. 194,39 -47.[CrossRef][Medline]
Clay, J. L., Ammann, R. R. and Gomer, R. H. (1995). Initial cell-type choice in a simple eukaryote: cell-autonomous or morphogen-gradient dependent? Dev. Biol. 172,665 -674.[CrossRef][Medline]
Dien, B. S., Peterson, M. S. and Srienc, F. (1994). Cell-cycle analysis of Saccharomyces cerevisiae. Methods Cell Biol. 42,457 -475.[Medline]
Durston, A. J. and Vork, F. (1978). The spatial pattern of DNA synthesis in Dictyostelium discoideum slugs. Exp. Cell Res. 115,454 -457.[Medline]
Durston, A. J., Weijer, C. J., Jongkind, J. F., Verkerk, A., Timmermans, A. and Te Kulve, W. (1984). A flow fluorimetric analysis of the cell cycle during growth and differentiation in Dictyostelium discoideum. W.R. Arch. Dev. Biol. 194, 18-24.
Firtel, R. and Bonner, J. (1972). Characterization of the genome of the cellular slime mold Dictyostelium discoideum. J. Mol. Biol. 66,339 -361.[Medline]
Fosnaugh, K. L. and Loomis, W. F. (1993). Enhancer regions responsible for temporal and cell-type-specific expression of a spore coat gene in Dictyostelium. Dev. Biol. 157, 38-48.[CrossRef][Medline]
Gerald, N. J., Damer, C. K., O'Halloran, T. J. and de Lozanne, A. (2001). Cytokinesis failure in clathrin-minus cells is caused by cleavage furrow instability. Cell Motil. Cytoskel. 48,213 -223.[CrossRef][Medline]
Gomer, R. H. and Ammann, R. R. (1996). A cell-cycle phase-associated cell-type choice mechanism monitors the cell cycle rather than using an independent timer. Dev. Biol. 174, 82-91.[CrossRef][Medline]
Gomer, R. H. and Firtel, R. A. (1987). Cell-autonomous determination of cell-type choice in Dictyostelium development by cell-cycle phase. Science 237,758 -762.[Medline]
Good, J. R., Cabral, M., Sharma, S., Yang, J., van Driessche,
N., Shaw, C. A., Shaulsky, G. and Kuspa, A. (2003).
TagA, a putative serine protease/ABC transporter of Dictyostelium that is
required for cell fate determination at the onset of development.
Development 130,2953
-2965.
Gross, J. D., Bradbury, J., Kay, R. R. and Peacey, M. J. (1983). Intracellular pH and the control of cell differentiation in Dictyostelium discoideum. Nature 303,244 -245.[Medline]
Haberstroh, L. and Firtel, R. A. (1990). A spatial gradient of expresssion of a cAMP-regulated prespore cell type-specific gene in Dictyostelium. Genes Dev. 4, 596-612.[Abstract]
Halevy, O., Novitch, B. G., Spicer, D. B., Skapek, S. X., Rhee, J., Hannon, G. J., Beach, D. and Lassar, A. B. (1995). Correlation of terminal cell cycle arrest of skeletal muscle with induction of p21 by MyoD. Science 267,1018 -1021.[Medline]
Hsieh, F. F., Barnett, L. A., Green, W. F., Freedman, K.,
Matushansky, I., Skoultchi, A. I. and Kelley, L. L.
(2000). Cell cycle exit during terminal erythroid differentiation
is associated with accumulation of p27(Kip1) and inactivation of cdk2 kinase.
Blood 96,2746
-2754.
Iranfar, N., Fuller, D., Sasik, R., Hwa, T., Laub, M. and
Loomis, W. F. (2001). Expression patterns of
cell-type-specific genes in Dictyostelium. Mol. Biol.
Cell 12,2590
-2600.
Kessin, R. H. (2001). Dictyostelium evolution, cell biology, and the development of multicellularity. Cambridge, UK: Cambridge University Press.
Knecht, D. A., Cohen, S. M., Loomis, W. F. and Lodish, H. F. (1986). Developmental regulation of Dictyostelium discoideum actin gene fusions carried on low-copy and high-copy transformation vectors. Mol. Cell Biol. 6,3973 -3983.[Medline]
Kuspa, A. and Loomis, W. F. (1996). Ordered
yeast artificial chromosome clones representing the Dictyostelium discoideum
genome. Proc. Natl. Acad. Sci. USA
93,5562
-5566.
Kuspa, A., Maghakian, D., Bergesch, P. and Loomis, W. F. (1992). Physical mapping of genes to specific chromosomes in Dictyostelium discoideum. Genomics 13, 49-61.[Medline]
Kuspa, A., Dingermann, T. and Nellen, W. (1995). Analysis of gene function in Dictyostelium. Experientia 51,1116 -1123.[Medline]
Leach, C. K. and Ashworth, J. M. (1972). Characterization of DNA from the cellular slime mould Dictyostelium discoideum after growth of the amoebae in different media. J. Mol. Biol. 68,35 -48.[Medline]
Leach, C. K., Ashworth, J. M. and Garrod, D. R. (1973). Cell sorting out during the differentiation of mixtures of metabolically distinct populations of Dictyostelium discoideum. J. Embryol. Exp. Morphol. 29,647 -661.[Medline]
Luo, Q., Michaelis, C. and Weeks, G. (1994).
Overexpression of a truncated cyclin B gene arrests Dictyostelium cell
division during mitosis. J. Cell Sci.
107,3105
-3114.
Luo, Q., Michaelis, C. and Weeks, G. (1995). Cyclin B and Cdc2 expression and Cd2 kinase activity during Dictyostelium differentiation. DNA Cell Biol. 14,901 -908.[Medline]
MacWilliams, H., Gaudet, P., Deichsel, H., Bonfils, C. and Tsang, A. (2001). Biphasic expression of rnrB in Dictyostelium discoideum suggests a direct relationship between cell cycle control and cell differentiation. Differentiation 67, 12-24.[CrossRef][Medline]
Maeda, Y. (1993). Pattern formation in a cell-cycle dependent manner during the development of Dictyostelium discoideum. Dev. Growth Differ. 35,609 -616.
Maeda, Y. (1997). Cellular and molecular mechanisms of the transition from growth to differentiation in Dictyostelium cells. In Dictyostelium A Model System for Cell and Developmental Biology (ed. Y. Maeda K. Inouye and I. Takeuchi), pp. 207-218. Tokyo, Japan: Universal Academy Press.
Maeda, Y. and Maeda, M. (1974). Heterogeneity of the cell population of the cellular slime mold Dictyostelium discoideum before aggregation, and its relation to the subsequent locations of the cells. Exp. Cell Res. 84,88 -94.[Medline]
Maeda, Y., Ohmori, T., Abe, T., Abe, F. and Amagai, A. (1989). Transition of starving Dictyostelium cells to differentiation phase at a particular position of the cell cycle. Differentiation 41,169 -175.[Medline]
Mann, S. K. O., Devreotes, P. N., Eliott, S., Jermyn, K., Kuspa, A., Fechheimer, M., Furukawa, R., Parent, C. A., Segall, J., Shaulsky, G. et al. (1998). Cell biological, molecular genetic, and biochemical methods to examine Dictyostelium. In Cell Biology A Laboratory Handbook (ed. J. E. Celis), pp.431 -466. San Diego, CA: Academic Press.
McDonald, S. A. and Durston, A. J. (1984). The cell cycle and sorting behaviour in Dictyostelium discoideum. J. Cell Sci. 66,195 -204.[Abstract]
Ogawa, S., Yoshino, R., Angata, K., Iwamoto, M., Pi, M., Kuroe, K., Matsuo, K., Morio, T., Urushihara, H., Yanagisawa, K. et al. (2000). The mitochondrial DNA of Dictyostelium discoideum: complete sequence, gene content and genome organization. Mol. Gen. Genet. 263,514 -519.[CrossRef][Medline]
Ohmori, R. and Maeda, Y. (1987). The developmental fate of Dictyostelium discoideum cells depends greatly on the cell-cycle position at the onset of starvation. Cell Differ. 22,11 -18.[CrossRef][Medline]
Saran, S., Azhar, M., Manogaran, P. S., Pande, G. and Nanjundiah, V. (1994). The level of sequestered calcium in vegetative amoebae of Dictyostelium discoideum can predict post-aggregative cell fate. Differentiation 57,163 -169.[CrossRef][Medline]
Sharpe, P. T., Knight, G. M. and Watts, D. J. (1984). Changes in the DNA content of amoebae of Dictyostelium discoideum during growth and development. Biochem. J. 217,839 -843.[Medline]
Sharpe, P. T. and Watts, D. J. (1984). Cell cycle-related changes in the surface properties of amoebae of the cellular slime mould Dictyostelium discoideum. FEBS Lett. 168, 89-92.[CrossRef]
Shaulsky, G. and Loomis, W. F. (1995). Mitochondrial DNA replication but no nuclear DNA replication during development of Dictyostelium. Proc. Natl. Acad. Sci. USA 92,5660 -5663.[Abstract]
Shaulsky, G., Kuspa, A. and Loomis, W. F. (1995). A multidrug resistance transporter/serine protease gene is required for prestalk specialization in Dictyostelium. Genes Dev. 9,1111 -1122.[Abstract]
Soll, D. R., Yarger, J. and Mirick, M. (1976). Stationary phase and the cell cycle of Dictyostelium discoideum in liquid nutrient medium. J. Cell Sci. 20,513 -523.[Abstract]
Sucgang, R., Chen, G., Liu, W., Lindsay, R., Lu, J., Muzny, D.,
Shaulsky, G., Loomis, W., Gibbs, R. and Kuspa, A.
(2003). Sequence and structure of the extrachromosomal palindrome
encoding the ribosomal RNA genes in Dictyostelium. Nucleic Acids
Res. 31,2361
-2368.
Sussman, M. (1987). Cultivation and synchronous morphogenesis of Dictyostelium under controlled experimental conditions. In Methods in Cell Biology, Vol.28 (ed. J. A. Spudich), pp.9 -29. Orlando, FL: Academic Press.[Medline]
Sussman, R. and Rayner, E. P. (1971). Physical characterization of deoxyribonucleic acids in Dictyostelium discoideum. Arch. Biochem. Biophys. 144,127 -137.[Medline]
Thompson, C. R. L. and Kay, R. R. (2000). Cell-fate choice in Dictyostelium: intrinsic biases modulate sensitivity to DIF signaling. Dev. Biol. 227, 56-64.[CrossRef][Medline]
Tsang, A., Bonfils, C., Czaika, G., Shtevi, A. and Grant, C. (1996). A prespore-specific gene of Dictyostelium discoideum encodes the small subunit of ribonucleotide reductase. Biochim. Biophys. Acta 1309,100 -108.[Medline]
VanDriessche, N., Shaw, C., Katoh, M., Morio, T., Sucgang, R.,
Ibarra, M., Kuwayama, H., Saito, T., Urushihara, H., Maeda, M. et
al. (2002). A transcriptional profile of multicellular
development in Dictyostelium discoideum. Development
129,1543
-1552.
Vollrath, D. and Davis, R. W. (1987). Resolution of DNA molecules greater than 5 megabases by contour-clamped homogeneous electric fields. Nucleic Acids Res. 15,7865 -7876.[Abstract]
Weeks, G. and Weijer, C. J. (1994). The Dictyostelium cell cycle and its relationship to differentiation. (Minireview). FEMS Microbiol. Lett. 124,123 -130.[CrossRef][Medline]
Weijer, C. J., Duschl, G. and David, C. N. (1984a). Dependence of cell-type proportioning and sorting on cell cycle phase in Dictyostelium discoideum. J. Cell Sci. 70,133 -145.[Abstract]
Weijer, C. J., Duschl, G. and David, C. N. (1984b). A revision of the Dictyostelium discoideum cell cycle. J. Cell Sci. 70,111 -131.[Abstract]
Williams, J. G., Duffy, K. T., Lane, D. P., McRobbie, S. J., Harwood, A. J., Traynor, D., Kay, R. R. and Jermyn, K. A. (1989). Origins of the prestalk-prespore pattern in Dictyostelium development. Cell 59,1157 -1163.[Medline]
Wood, S. A., Ammann, R. R., Brock, D. A., Li, L., Spann, T. and
Gomer, R. H. (1996). RtoA links initial cell type
choice to the cell cycle in Dictyostelium. Development
122,3677
-3685.
Zada-Hames, I. M. and Ashworth, J. M. (1978a). The cell cycle and its relationship to development in Dictyostelium discoideum. Dev. Biol. 63,307 -320.[Medline]
Zada-Hames, I. M. and Ashworth, J. M. (1978b). The cell cycle during the vegetative stage of Dictyostelium discoideum and its response to temperature change. J. Cell Sci. 32, 1-20.[Abstract]
Zavitz, K. H. and Zipursky, S. L. (1997).
Controlling cell proliferation in differentiating tissues: genetic analysis of
negative regulators of G1S-phase progression. Curr. Opin.
Cell Biol. 9,773
-781.[CrossRef][Medline]
Zhang, P., Wong, C., Liu, D., Finegold, M., Harper, J. W. and
Elledge, S. J. (1999). p21(CIP1) and p57(KIP2) control
muscle differentiation at the myogenin step. Genes
Dev. 13,213
-224.
Zimmerman, W. and Weijer, C. J. (1993). Analysis of cell cycle progression during the development of Dictyostelium and its relationship to differentiation. Dev. Biol. 160,178 -185.[CrossRef][Medline]
Related articles in Development: