1 Department of Biochemistry and Molecular Biology, Baylor College of Medicine,
Houston, Texas, 77030, USA
2 Department of Molecular and Human Genetics, Baylor College of Medicine,
Houston, Texas, 77030, USA
3 Program in Developmental Biology, Baylor College of Medicine, Houston, Texas,
77030, USA
Present address: Science and Technology Division, Institute for Defense
Analyses, 4850 Mark Center Drive, Alexandria, VA 22311-1882, USA
Author for correspondence (e-mail:
akuspa{at}bcm.tmc.edu)
Accepted 27 March 2003
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SUMMARY |
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Key words: differentiation, starvation, cell cycle, cell fate, Dictyostelium
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INTRODUCTION |
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In Dictyostelium discoideum, unique members of the ABCB
transporter subfamily, TagB and TagC, appear responsible for peptide signal
export during development. These Tag proteins are required for cell
differentiation in Dictyostelium and have the potential to carry out
the processing and transport of peptide signals since they possess an
N-terminal serine protease domain and a C-terminal transporter domain. The
available genetic data suggest that the function of the Tag proteins is to
transport signaling peptides that regulate the timing and nature of cell
differentiation. During the development of Dictyostelium, equipotent
cells differentiate into prespore cells and distinct populations of prestalk
cells that later form the various tissues in the terminally differentiated
fruiting body that consists of mature spores held atop a cellular stalk
(Kessin, 2001). The TagB
protein is required for cell-cell signaling that promotes spore encapsulation,
and it is required for the differentiation of prestalk A (PstA) cells in a
cell-autonomous fashion (Shaulsky et al.,
1995
). TagC is structurally similar to TagB and the phenotypic
similarities between tagC null mutants and tagB mutants are
so striking that it is presumed that TagB and TagC function as a heterodimer.
The conserved amino acid residues known to be required for serine protease
catalytic activity and for ABC transporter ATPase activity are required for
TagB function (G.S., unpublished). In addition, TagC-null mutants fail to
release the spore encapsulation-inducing peptide SDF-2
(Anjard et al., 1998
). Thus, it
is likely that Tag proteins work by the proteolytic processing and transport
of signaling peptides, in a manner analogous to that in which STE6 transports
the proteolytically processed, lipid-modified a-factor pheromone peptide of
S. cerevisiae (Taglicht and
Michaelis, 1998
). Such a cell-cell signaling function is the
obvious common feature of all of the transporters characterized as being
involved in developmental signaling. A curious feature of the tagB
mutant phenotype is the cell-autonomous defect in prestalk cell
differentiation experienced by cells that would normally express the
tagB gene in wild-type cell development
(Shaulsky and Loomis, 1996
).
This novel feature suggests that the failure of a transporter to export a
substrate/signal can affect the signal-producing cell as well.
Developing Dictyostelium aggregates contain several prestalk cell
sub-populations that can be distinguished from the prespore cells and can be
tracked with cell-type-specific reporter genes
(Williams, 1997). Near the end
of development, the PstAB cells initiate fruiting body formation by plunging
through the prespore mass, forming a cellulosic stalk tube until they make
contact with the substratum. The PstA cells enter the stalk tube following the
PstAB cells and differentiate into stalk cells. As the prespore cells crawl up
the elongating stalk they differentiate into mature spores. Terminal spore
differentiation proceeds in a wave from top to bottom of the nascent sorus.
This observation is consistent with a number of experiments that suggest a
signal emanating from the PstA cells coordinates terminal spore
differentiation with fruiting body morphogenesis
(Harwood et al., 1993
;
Richardson et al., 1994
;
Shaulsky et al., 1995
). The
TagB and TagC transporters are expressed in PstA cells and so they are in the
position to affect spore differentiation during this process
(Shaulsky and Loomis, 1996
)
(G.S., unpublished data). The most plausible model is that TagC exports the
encapsulation-inducing SDF-2 peptide, initiating the observed wave of spore
differentiation as the PstA cells enter the stalk tube at the top of the
prespore mass (Anjard et al.,
1998
). However, there is also a cell-autonomous requirement for
both TagB and TagC in the formation of PstA-derived stalk cells. Cells that do
not express a TagB or a TagC transporter cannot become stalk cells
(Shaulsky et al., 1995
). This
aspect of Tag transporter function is not understood, but could be explained
if the transport substrate acted as an inhibitor of PstA cell differentiation
when retained inside the cell. Given an ABC transporter's capacity to maintain
a concentration differential of a small molecule across a membrane, ABC
transporter activity could alter cellular physiology in a way that would
promote or inhibit a particular differentiation program. In theory, the flux
of a signal through a single transporter could control the fates of the
signal-producing cells as well as the responding cells. There is some
tentative evidence for cell-autonomous functions of ABC transporters in cell
differentiation. In Dictyostelium, the prespore-specific rhodamine
transporter RhT may be required for the production or maintenance of prespore
cells (Good and Kuspa, 2000
).
In mammalian systems, the ABC transporter encoded by the ABCB1 (MDR1)
gene has been implicated in the maintenance of hematopoietic stem cells and in
the regulation of programmed cell death
(Smyth et al., 1998
;
Johnstone et al., 1999
;
Johnstone et al., 2000a
;
Johnstone et al., 2000b
).
Recently, the ABCG2 gene has also been suggested to have a role in
the maintenance of the undifferentiated state of stem cells
(Zhou et al., 2001
).
We have identified a novel member of the tag gene family, tagA. TagA mRNA is expressed in the first 2 hours following starvation, it becomes specific to prespore cells and is eventually expressed in mature spores. Inactivation of the tagA gene results in developing structures with enlarged or supernumerary prestalk regions with roughly twice the normal number of prestalk cells. Within these structures, the tagA- cells that activate the tagA promoter do not become spores but instead adopt a prestalk cell fate. Our results imply that the earliest cells to differentiate require TagA-mediated function to prevent their adoption of a prestalk cell fate, or for promoting their prespore cell fate.
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MATERIALS AND METHODS |
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Strain construction, cell growth and development
The Dictyostelium strains used in this study are described in
Table 1. Ax4 cells were grown
in HL-5 liquid medium (Sussman,
1987) supplemented with streptomycin (50 µg/ml) and penicillin
(50 U/ml). Neomycin-resistant strains (neor)
Ax4[cotB/GFP], Ax4[ecmA/GFP], Ax4[act15/GFP] and
all respective derivatives, were grown in HL-5 liquid medium supplemented with
20 µg/ml G418 (Geneticin, Gibco). All strains were removed from
drug-containing medium 36 hours prior to assay. Cells were plated for
synchronous development on nitrocellulose filters as described previously
(Sussman, 1987
).
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An expression plasmid with the tagA coding region, under the
control of its native promoter was constructed by substituting a 7.1 kb
XbaI/HindIII tagA fragment, described above, for
the lacZ/act8 cassette in pDdGal16(H+)
(Harwood and Drury, 1990). The
resulting plasmid, ptagA/tagA, was transformed into
tagA- cells by calcium-phosphate precipitation
(Nellen and Firtel, 1985
). The
ptagA/lacZ expression plasmid was made with genomic DNA 2 kb
upstream of the tagA coding region including the 5' end of the
coding sequence up to the first BamHI site. This 2.2 kb
EcoRI-BamHI fragment was inserted into pDdGal16(H+) between
its EcoRI and BglII sites. Staining of developmental
structures and spores for ß-galactosidase was carried out as described
previously (Shaulsky and Loomis,
1993
).
Spore and stalk cell assays
Sporulation was measured by harvesting cells from filters into 20 mM
potassium phosphate buffer, pH 6.2, and treating them with 0.4% non-ionic
detergent NP-40 for 10 minutes at 22°C. Cells were then washed with
potassium phosphate buffer twice, and disaggregated by trituration with an
18-gauge needle. Refractile spores were counted by phase-contrast microscopy
and plated on SM agar plates with bacteria. The number of colonies was used as
an estimate of the number of viable spores in each sample. At least three
independent determinations were carried out for each strain and are reported
as the mean±s.e.m.
Induction of prestalk gene expression was measured in submerged culture
(Harwood et al., 1995) as
modified previously (Wang and Kuspa,
2002
). Vegetative cells were harvested at a density of
1-2x106/ml, washed once in 20 mM sodium phosphate buffer (pH
6.4) and three times in stalk buffer [10 mM Mes, 2 mM NaCl, 10 mM KCl, 1 mM
CaCl2, 50 µg/ml streptomycin, 50 U/ml penicillin (pH 6.2)].
Cells were plated at 2.5x104 cells/cm2 in stalk
buffer supplemented with 5 mM cAMP. After 24 hours, cell cultures were washed
free of cAMP with stalk buffer and the original buffer volume was replaced
with stalk buffer with or without supplements. Cells were assayed for prestalk
gene expression 24 hours later by fluorescence microscopy, via an
ecmA/GFP reporter construct and scored for the production of
stalk-like cells using phase-contrast microscopy. Cellulose deposition by the
stalk-like cells was confirmed by staining with calcafluor
(Harrington and Raper,
1968
).
Protein and RNA expression assays
Antibodies against TagA protein were raised at Bethyl Laboratories, Inc.
(Montgomery, Texas). The peptide LPSNSRNTRNADKLRNRSET, representing amino
acids 1627-1646 of the predicted TagA protein, was synthesized and conjugated
to keyhole limpet hemocyanin via a cysteine residue added at the amino
terminal end of the peptide. The conjugated peptide was used to immunize
rabbits and the resulting polyclonal antibodies were affinity purified on a
column conjugated with the peptide antigen.
Cells were harvested at various times during development and resuspended in 50 mM Tris-HCl (pH 8.0), 5 mM EDTA, 150 mM NaCl, 0.5% NP-40, 1 mM PMSF. Protein concentrations were determined with the BioRad protein determination kit (BioRad Laboratories, Richmond CA) and equal amounts of protein were resolved on 6% polyacrylamide gels. Protein was electrotransferred to a nitrocellulose membrane and detected with the affinity-purified anti-peptide antibody described above, followed by goat anti-rabbit antibody and visualized with the ECL kit according to the manufacturer's protocols (Amersham Life Sciences).
RNA was isolated from the wild type and the transformed strains during
vegetative growth and development. Spores and stalks were purified as
described by Van Driessche et al. (Van
Driessche et al., 2002). Spores and stalks purified by this
procedure were estimated to be >99% pure by direct microscopic observation.
Cells were harvested and suspended in TRIzol Reagent (Gibco BRL). RNA
extraction was performed according the manufacturer's protocol. RNase
protection assays were performed using a RPA III kit (Ambion) according to the
manufacturer's protocol with probes synthesized from various fragments of
tagA cDNA, or genomic DNA, templates using a riboprobe in vitro
transcription system (Promega), as described in the text.
Expression profiling with microarrays
Expression profiling was carried out with DNA microarrays as described
previously (Van Driessche et al.,
2002). Cells (108) developing on two filters were
harvested and processed to produce total RNA for each time point. The raw data
from the microarray hybridizations were processed according to the procedure
described in Van Driessche et al. (Van
Driessche et al., 2002
). Briefly, raw image files were quantified
and the resulting data was subjected to a single array normalization procedure
to remove spatial and intensity artifacts and to put the data on a common
measurement scale (Yang et al.,
2002
). Replicate arrays from the same biological preparation were
averaged, and the averages for two biological preparations were then averaged
to yield the final data set. We compared the time patterns of gene expression
in the tagA mutant cells and our previous wild-type data
(Van Driessche et al., 2002
).
The 2,021 genes whose expression levels were altered dramatically during
development were determined and ordered in the `blue-yellow' plots as
described previously (Van Driessche et
al., 2002
). Each gene's studentized score was compared against the
linear contrast function y=x/121 to evaluate how strongly and
consistently the gene appeared to change in relative expression across
developmental time. The same gene order is used in all plots shown.
To compare microarray data for all genes from each time point of
tagA mutant development with each time point of wild-type development
we determined the time point for the wild type at which the expression pattern
was most similar to that found in the tagA- mutant by
using the Pearson correlation distance to make comparison. To measure the
distance we used data from all of the genes that were not excluded from the
analysis for quality control reasons (6,000 genes).
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RESULTS |
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The propensity of tagA mutants to differentiate as ecmA-positive prestalk cells was examined in chimeras with wild-type cells to explore the possibility that this phenotype results from a perturbation of intercellular signaling. Chimeras were made with wild-type and tagA mutant cells at various ratios in which one of the strains was marked with the ecmA/GFP reporter to follow the prestalk population of each strain. Prestalk cells derived from the tagA- cells were consistently overrepresented in the prestalk cell population of the chimeras (Table 2). For instance, when only 10% of the cells were tagA- 18% of the prestalk cells came from the mutant population. We also carried out an analogous set of experiments with the prespore reporter (cotB/GFP) and found that the tagA mutants were slightly underrepresented in the prespore population, as expected (data not shown). Finally, we demonstrated that tagA mutant cells participate in development of the chimeras by examining mixtures of control strains that expressed GFP under the control of the actin15 promoter. Wild-type (Ax4[act15/GFP]) or mutant (tagA-[act15/GFP]) strains were mixed with unmarked cells to determine their representation within aggregates and both strains contributed at the expected percentage to the developing populations within each type of cell mixture described in Table 2 (data not shown). Since tagA mutant cells are able to enter aggregates as well as wild-type cells, these results suggest that the additional prestalk cells result from a cell-autonomous defect in cell differentiation.
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PstB cells defined by the ecmB gene are scattered throughout the
finger and slug structures prior to terminal differentiation
(Williams, 1997). PstB cells
at the bottom of the prespore region of the slug can contribute to the basal
disk. In the absence of slug migration, a population of `rearguard' cells will
form the basal disc proper while the PstB cells will form the outer basal disc
and the lower cup at the base of the sorus. Expression of the ecmB
gene appears to be precocious and elevated in the tagA mutants
(Fig. 4A). Additional
ecmB-positive cells are also apparent in tagA mutant slugs
but they do not appear to expand the volume of ecmB-positive tissues
in the final fruiting body as we observed with ecmA expression
(Fig. 4B).
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|
Cell type specificity of gene expression is compromised in tagA
mutants
Tissue-specific expression of the tagA gene was determined by
expressing lacZ under the control of the tagA promoter in
wild-type and mutant cells. This promoter is likely to be complete as it
rescued TagA protein expression and it corrected the development of
tagA mutant cells (Fig.
2B). Histochemical staining of the developing
Ax4[tagA/lacZ] cells for ß-galactosidase showed expression of
the tagA gene in the prespore region of developing fingers and in the
sori of fruiting bodies (Fig.
6A and data not shown). We isolated spores from mature fruiting
bodies after 36 hours of development and stained them for ß-galactosidase
activity. We found that 82±8.0% of wild-type spores appeared to express
the tagA/lacZ reporter construct as judged by their blue color in
bright-field microscopy. In all developing structures observed, the tips of
fingers and anterior (PstA) region of the slug showed no detectable
lacZ expression. In the tagA-[tagA/lacZ]
strain, very faint staining could be observed in the prespore region during
development, but only after 24 hours of staining. Upon culmination, weak
ß-galactosidase expression was evident only in cells of the outer basal
disc and in the lower cup of the fruiting body
(Fig. 6A). These
tagA-expressing cells occupied the same position in the mutant
fruiting bodies as the extra ecmA-positive cells described above.
Interestingly, there was no detectable staining in the tagA mutant
sori. Very few spores isolated from these fruiting bodies displayed any
ß-galactosidase activity as determined by staining for 48 hours
(0.58±0.62%). To confirm this result we attempted an RNase protection
assay on spore and stalk RNA with a probe that lies between the promoter and
the insertion mutation in the tagA gene. The samples were harvested
at a time that the tagA mRNA levels are predicted to be extremely low
in wild-type cells (24 hours; Fig.
2) and less than ten percent of the tagA mutant cells are
expected to express the gene (Fig.
6A). In spite of this, the assay consistently revealed an
enrichment of tagA mRNA in the spores of the wild type, as expected,
and a consistently higher signal in the stalk tissue relative to the spores
produced by tagA mutant cells
(Fig. 6B).
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These results suggest that a fate change occurs in tagA-expressing
cells within the tagA- cell population so that cells that
would have become prespore cells develop into a cell type similar in character
to PstB cells. Since the expression profiling suggested that the majority of
tagA- cells are affected by the loss of TagA we examined
the potential for additional alterations of cell specification by surveying
the expression of archetypal cell-specific genes. The spore coat protein gene
cotB is a reliable marker of prespore and spore cells and is
coordinately regulated with several other spore coat protein genes
(Fosnaugh and Loomis, 1991).
The cotB/lacZ reporter construct used to visualize cotB
expression showed a normal staining pattern in wild-type cells, with all the
spores staining blue with X-gal and no staining of the stalk cells
(Fig. 6C). TagA mutant spores
stained as expected, but many of the vacuolated stalk cells were also stained,
revealing that these cells had expressed the cotB gene at some time
in development (Fig. 6C). The
spiA gene is normally expressed exclusively within encapsulating
prespore cells and is required for the long-term stability of dormant spores
(Richardson and Loomis, 1992
;
Richardson et al., 1994
). The
spiA mRNA displayed slightly lower spore/stalk enrichment in the
tagA mutant samples compared to the wild type
(Fig. 6D). Intriguingly, the
ecmB mRNA appears to be present in the spore RNA purified from
tagA mutants in a much higher proportion than in the wild type. This
contrasts with the expression pattern observed with the ecmB/lacZ
reporter gene that showed little expression in tagA mutant prespore
cells or spores (Fig. 4). This
difference suggests that the promoter present in the ecmB/lacZ
construct is not active in tagA mutant prespore cells or that the
native ecmB gene has additional promoter elements that are missing in
the artificial lacZ construct. Alternatively, the native
ecmB mRNA may be more stable than lacZ mRNA in tagA
mutant prespore cells. Nevertheless, the unexpected patterns of cotB,
ecmB and tagA expression in tagA mutants indicate that
tagA mutants produce terminally differentiated cells in spite of
substantial mis-expression of cell-specific genes.
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DISCUSSION |
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The analysis of the tagA gene suggests that overt prespore cell
differentiation may occur as early as 2 hours of development. TagA mRNA and
protein begin to accumulate to significant levels within the first 2 hours of
development. The results obtained with the tagA/lacZ reporter
construct indicate that TagA expression becomes spore specific. It is possible
that the tagA promoter is active in all cells early, the
ß-galactosidase is turned over during aggregation and tagA
expression becomes spore specific only later, but the more than 8-hour
half-life of the ß-galactosidase produced from this construct argues
against this possibility (Detterbeck et
al., 1994). Thus, it is possible that some cells require the
expression of tagA in the first 2 hours of development in order to
differentiate as prespore cells. In this regard, it is interesting that the
first significant increase in the curve that describes the increase in
ecmA-expressing cells in the tagA mutant extrapolates back
to about 6 hours of development (Fig.
3). In addition, the global gene expression profiling of the
tagA mutant revealed a delay in the developmental program beginning
between 2 and 4 hours of development. Thus, the cell fate and gene expression
changes that we observed in the mutant support the idea that the first
critical time for TagA function is prior to 6 hours of development.
It has generally been accepted that cell-type-specific gene expression
begins at about eight hours, as the mound forms during aggregation
(Williams et al., 1989;
Haberstroh and Firtel, 1990
;
Fosnaugh and Loomis, 1993
).
However, recent reports suggest that some form of cell-type divergence may
occur much earlier. Iranfar and co-workers uncovered six genes with cell-type
specific expression at the slug stage that initiate expression between 2 and 5
hours of development (Iranfar et al.,
2001
). Van Driessche and co-workers identified dozens of
cell-type-enriched mRNAs that reach their highest expression levels from 2 to
6 hours of development (Van Driessche et
al., 2002
). More detailed analyses may reveal that these mRNAs
become cell-type enriched much later in development through, for instance,
their differential stability within the different cell types. It is also
possible that these transcripts reflect an early divergence in the
physiological state of cells within the starving population that influences
cell-type specification.
Physiological differences, such as prior growth conditions and cytosolic
pH, between vegetative cells have been observed to influence cell type
divergence later in development (e.g.,
Leach et al., 1973;
Maeda and Maeda, 1974
;
Gross et al., 1983
). It is
well documented that cell cycle phase at the time of starvation influences
cell differentiation later in development (reviewed by
Maeda, 1997
). In fact, this
influence can be observed as cell-autonomous cell-type specification in low
cell density cultures and is under regulation by the RtoA protein
(Gomer and Firtel, 1987
;
Wood et al., 1996
). It is
likely that the influence of the cell cycle is to provide a bias in cell fate
determination that is realized by later signaling through, for example, cAMP,
DIF or calcium (e.g. Clay et al.,
1995
; Thompson and Kay,
2000
; Azhar et al.,
2001
). A prespore-specific transporter has been characterized,
RhT, that appears to be involved in prespore cell differentiation and whose
activity can be detected prior to the expression of the spore coat protein
gene, cotB, as the prestalk and prespore regions are coalescing in
the early mound (Good and Kuspa,
2000
). All of these physiological differences amongst cells can
predispose a particular cell to one cell fate or another, but it is generally
accepted that these biases do not determine cell fate and are reversible in
different experimental contexts in vivo. The tagA gene is an example
of a prespore-specific gene that is expressed at the onset of development and
is also required for the sporulation of those cells that express it. Thus,
tagA provides genetic evidence that some prespore cell
differentiation occurs well before the aggregation stage and may provide a
link between the physiological status of growing cells and the cell fate
determination that occurs in the first few hours of development.
We used transcriptional profiling as a way of obtaining a global view of the physiological state of the tagA mutants during development. Monitoring global changes in gene expression allows the detection of mutation-induced deviations from an otherwise robust transcriptional program. The early pause in the transcriptional program that we observed between 2 and 6 hours of development in the tagA mutant coincides well with the onset of TagA RNA and protein expression in the wild type. The ablation of the normal transcriptional program precisely when TagA is first expressed reinforces the notion that tagA plays an important role in early development. It is important to note that we would not have observed this pause unless the majority of cells in the population had experienced a 4-hour delay in the transcriptional program. At least 70% of wild-type cells express tagA at some time during development, but only about ten percent of the cells express detectable levels of tagA in the mutant as judged by ß-galactosidase staining. These facts together with the delay in the transcriptional program and the inappropriate expression of the cotB and ecmB genes suggests that most cells are affected by the loss of tagA, but most of them compensate for the loss and go on to make spores and stalk while a small percentage of cells are directed to an anomalous PstB-like state. The fact that the early delay in global gene expression in tagA mutants lasts for 4 hours suggests that the defect in the unicellular to multicellular transition stems from a failure to make an initial population of TagA-expressing cells in a timely fashion. The later delay in the transcriptional program in the tagA mutant, between 16 and 24 hours, suggests that TagA functions late or that the mutants lose synchrony as development proceeds. This fits with the observed morphological asynchrony, the 12-hour delay in the completion of development and low viability of tagA mutant spores.
Although we have yet to develop an assay for the biological function that
is mediated by TagA, we were able to obtain indirect evidence of a putative
TagA signaling event by monitoring deviation from the wild type in the
expression of thousands of genes early in development. This is important given
that the mutant organism can compensate for the lack of TagA, making it
difficult to explore TagA function using cellular or morphological criteria.
TagA is most similar in its predicted structure to TagC and TagB. TagC has
been implicated in the cellular export of the peptide signal, SDF-2, thought
to stimulate the terminal differentiation of prespore cells; there is also
genetic evidence that TagB is required for this signaling event
(Shaulsky et al., 1995;
Anjard et al., 1998
). The
protease/transporter homology of TagA and the cell-autonomous phenotype of
tagA mutants suggest that TagA exports a peptide that must be cleaved
and removed from the cell for prespore cell differentiation to occur prior to
aggregation. The export of a differentiation inhibitor was also proposed to
explain the cell-autonomous specification of PstA cells by TagB and the
maintenance of the undifferentiated state of stem cells in mammals
(Shaulsky et al., 1995
;
Zhou et al., 2001
).
It will be important to determine the regulatory pathways that
tagA impinges on within the prespore cells that are most affected by
loss of tagA function. There are two other genes whose inactivation
results in the cell-autonomous production of PstB-like cells: one encodes the
Dictyostelium homolog of glycogen synthase kinase 3 (GSK-3),
gskA, and the other, stkA, encodes the Stalky protein that
resembles a GATA family transcription factor
(Harwood et al., 1995;
Chang et al., 1996
). When
gskA mutant cells are co-developed with wild-type cells they produce
PstB cells that occupy the lower cup and outer basal disk and they
over-express the ecmB gene
(Harwood et al., 1995
). Both
of these phenotypes are reminiscent of what we have observed in tagA
mutants. Thus, one possibility is that TagA is needed to maintain active GSK-3
in a small cohort of prespore cells early in development. Stalky mutants also
overproduce stalk cells, but stkA appears to function much later in
development than either tagA or gskA. The function of
stkA also appears to be independent of gskA
(Chang et al., 1996
). Future
work will focus on identifying the signaling pathways controlled by TagA and
the identification of the TagA substrate.
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ACKNOWLEDGMENTS |
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Footnotes |
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REFERENCES |
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Adachi, H., Hasebe, T., Yoshinaga, K., Ohta, T. and Sutoh, K. (1994). Isolation of Dictyostelium discoideum cytokinesis mutants by restriction enzyme-mediated integration of the blasticidin s resistance marker. Biochem. Biophys. Res. Commun. 205,1808 -1814.[CrossRef][Medline]
Ambudkar, S. V. and Gottesman, M. M., eds. (1998). ABC transporters: biochemical, cellular, and molecular aspects. Methods Enzymol. 292, 1-853.
Anjard, C., Zeng, C., Loomis, W. F. and Nellen, W. (1998). Signal transduction pathways leading to spore differentiation in Dictyostelium discoideum. Dev. Biol. 193,146 -155.[CrossRef][Medline]
Anjard, C., the Dictyostelium Sequencing Consortium and
Loomis, W. F. (2002). Evolution of the ABC transporters of
Dictyostelium. Eukaryotic Cell
1, 643-652.
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]
Bai, C. and Elledge, S. J. (1997). Gene identification using the yeast two-hybrid system. Meth. Enzymol. 283,141 -156.[Medline]
Chang, W. T., Newell, P. C. and Gross, J. D. (1996). Identification of the cell fate gene stalky in Dictyostelium. Cell 87,471 -481.[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]
Detmers, F. J., Lanfermeijer, F. C. and Poolman, B. (2001). Peptides and ATP binding cassette peptide transporters. Res Microbiol. 152,245 -258.[CrossRef][Medline]
Detterbeck, S., Morandini, P., Wetterauer, B., Bachmair, A.,
Fischer, K. and MacWilliams, H. K. (1994). The `prespore-like
cells' of Dictyostelium have ceased to express a prespore gene:
analysis using short-lived beta-galactosidases as reporters.
Development 120,2847
-2855.
Early, A. E., Gaskell, M. J., Traynor, D. and Williams, J.
G. (1993). Two distinct populations of prestalk cells within
the tip of the migratory Dictyostelium slug with differing fates at
culmination. Development
118,353
-362.
Fosnaugh, K. L. and Loomis, W. F. (1991). Coordinate regulation of the spore coat genes in Dictyostelium discoideum.Dev. Genet. 12,123 -132.[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]
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. and Kuspa, A. (2000). Evidence that a cell-type-specific efflux pump regulates cell differentiation in Dictyostelium. Dev. Biol. 220, 53-61.[CrossRef][Medline]
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]
Harrington, B. J. and Raper, K. B. (1968). Use of a fluorescent brightener to demonstrate cellulose in the cellular slime mold. J. Appl. Microbiol. 16,106 -113.
Harwood, A. J. and Drury, L. (1990). New vectors for expression of the E. coli lacZ gene in Dictyostelium.Nucleic Acids Res. 18,4292 .[Medline]
Harwood, A. J., Early, A. and Williams, J. G.
(1993). A repressor controls the timing and spatial localisation
of stalk cell-specific gene expression in Dictyostelium.Development 118,1041
-1048.
Harwood, A. J., Plyte, S. E., Woodgett, J., Strutt, H. and Kay, R. R. (1995). Glycogen synthase kinase 3 regulates cell fate in Dictyostelium. Cell 80,139 -148.[Medline]
Hughes, T. R., Marton, M. J., Jones, A. R., Roberts, C. J., Stoughton, R., Armour, C. D., Bennett, H. A., Coffey, E., Dai, H., He, Y. D. et al. (2000). Functional discovery via a compendium of expression profiles. Cell 102,109 -126.[Medline]
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.
Johnstone, R. W., Cretney, E. and Smyth, M. J.
(1999). P-glycoprotein protects leukemia cells against
caspase-dependent, but not caspase-independent, cell death.
Blood 93,1075
-1085.
Johnstone, R. W., Ruefli, A. A. and Smyth, M. J. (2000a). Multiple physiological functions for multidrug transporter P-glycoprotein? Trends Biochem. Sci. 25, 1-6.[CrossRef][Medline]
Johnstone, R. W., Ruefli, A. A., Tainton, K. M. and Smyth, M. J. (2000b). A role for P-glycoprotein in regulating cell death. Leuk. Lymphoma 38, 1-11.[Medline]
Kay, R. R. (1998). The biosynthesis of
differentiation-inducing factor, a chlorinated signal molecule regulating
Dictyostelium development. J. Biol. Chem.
273,2669
-2675.
Kessin, R. H. (2001). Dictyostelium: Evolution, cell biology, and the development of multicellularity. New York: Cambridge University Press.
Kim, S. K., Lund, J., Kiraly, M., Duke, K., Jiang, M., Stuart,
J. M., Eizinger, A., Wylie, B. N. and Davidson, G. S. (2001).
A gene expression map for Caenorhabditis elegans.Science 293,2087
-2092.
Knecht, D. A., Cohen, S. M. and Loomis, W. 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]
Kuchler, K. and Thorner, J. (1992). Secretion of peptides and proteins lacking hydrophobic signal sequences: the role of adenosine triphosphate-driven membrane translocators. Endocrinol. Rev. 13,499 -514.[Abstract]
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]
Lindsey, K., Casson, S. and Chilley, P. (2002). Peptides: new signalling molecules in plants. Trends Plant Sci. 7,78 -83.[CrossRef][Medline]
Ma, Y., Erkner, A., Gong, R., Yao, S., Taipale, J., Basler, K. and Beachy, P. (2002). Hedgehog-mediated patterning of the mammalian embryo requires transporter-like function of Dispatched. Cell 111,63 .[Medline]
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, 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]
Manstein, D. J. and Hunt, D. M. (1995). Overexpression of myosin motor domains in Dictyostelium: Screening of transformants and purification of the affinity tagged protein. J. Muscle Res. Cell Motil. 16,325 -332.[Medline]
Nellen, W. and Firtel, R. A. (1985). High-copy-number transformants and co-transformation in Dictyostelium. Gene 39,155 -163.[CrossRef][Medline]
Richardson, D. L. and Loomis, W. F. (1992). Disruption of the sporulation-specific gene spia in Dictyostelium discoideum leads to spore instability. Genes Dev. 6,1058 -1070.[Abstract]
Richardson, D. L., Loomis, W. F. and Kimmel, A. R.
(1994). Progression of an inductive signal activates sporulation
in Dictyostelium discoideum. Development
120,2891
-2900.
Sharom, F. J., Lu, P., Liu, R. and Yu, X. (1998). Linear and cyclic peptides as substrates and modulators of P-glycoprotein: peptide binding and effects on drug transport and accumulation. Biochem. J. 333,621 -630.[Medline]
Shaulsky, G. and Loomis, W. F. (1993). Cell type regulation in response to expression of ricin-A in Dictyostelium.Dev. Biol. 160,85 -98.[CrossRef][Medline]
Shaulsky, G. and Loomis, W. F. (1996). Initial cell type divergence in Dictyostelium is independent of DIF-1. Dev. Biol. 174,214 -220.[CrossRef][Medline]
Shaulsky, G., Kuspa, A. and Loomis, W. F. (1995). A multidrug resistance transporter serine protease gene is required for prestalk specialization in Dictyostelium. Genes Devel. 9,1111 -1122.[Abstract]
Smyth, M. J., Krasovskis, E., Sutton, V. R., Johnstone, R.
W. (1998). The drug efflux protein, P-glycoprotein,
additionally protects drug-resistant tumor cells from multiple forms of
caspase-dependent apoptosis. Proc. Natl. Acad.
Sci.,USA 95,7024
-7029.
Solomon, J. M., Lazazzera, B. A. and Grossman, A. D. (1996). Purification and characterization of an extracellular peptide factor that affects two different developmental pathways in Bacillus subtilis. Genes Dev. 10,2014 -2024.[Abstract]
Souza, G. M., Lu, S. and Kuspa, A. (1998).
YakA, a protein kinase required for the transition from growth to development
in Dictyostelium. Development
125,2291
-2302.
Stacey, G., Koh, S., Granger, C. and Becker, J. M. (2002). Peptide transport in plants. Trends Plant Sci. 7,257 -263.[CrossRef][Medline]
Sussman, M. (1987). Cultivation and synchronous morphogenesis of Dictyostelium under controlled experimental conditions. Methods Cell Biol. 28, 9-29.[Medline]
Taglicht, D. and Michaelis, S. (1998). Saccharomyces cerevisiae ABC proteins and their relevance to human health and disease. Methods Enzymol. 292,130 -162.[Medline]
Thompson, C. R. and Kay, R. R. (2000). Cell fate choice in Dictyostelium: intrinsic biases modulate sensitivity to DIF signailing. Dev. Biol. 227, 56-64.[CrossRef][Medline]
Van Driessche, N., Shaw, C., Katoh, M., Morio, T., Sucgang, R.,
Ibarra, M., Kuwayama, H., Saito, T., Urushihara, H., Maeda, M., Takeuchi, I.,
Ochiai, H., Eaton, W., Tollett, J., Halter, J., Kuspa, A., Tanaka, Y.,
Shaulsky, G. (2002). A transcriptional profile of
multicellular development in Dictyostelium discoideum.Development 129,1543
-1552.
Wang, B. and Kuspa, A. (2002). CulB, a putative
ubiquitin ligase subunit, regulates prestalk cell differentiation and
morphogenesis in Dictyostelium. Eukaryotic Cell
1, 126-136.
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]
Williams, J. G. (1997). Prestalk and stalk heterogeneity in Dictyostelium. In Dictyostelium A Model System for Cell and Developmental Biology, pp.293 -304. Tokyo, Japan: Universal Academy Press.
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.
Yang, Y. H., Dudoit, S., Lu, P., Lin, D. M., Peng, V., Ngai, J. and Speed, T. P. (2002). Normalization for cDNA microarray data: a robust composite method addressing single and multiple slide systematic variation. Nucleic Acids Res. 30, E15.[Medline]
Zhou, S., Schuetz, J. D., Bunting, K. D., Colapietro, A., Sampath, J., Morris, J. J., Irina Lagutina, I., Grosveld, G. C., Osawa, M., Nakauchi, H. and Sorrentino, B. P. (2001). The ABC transporter Bcrp1/ABCG2 is expressed in a wide variety of stem cells and is a molecular determinant of the side-population phenotype. Nat. Med. 7,1028 -1034.[CrossRef][Medline]