Received 13 June 1997/Returned for modification 28 July
1997/Accepted 27 October 1997
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INTRODUCTION |
One of the key points of cell cycle
regulation is the transition from G1 into S phase (50,
58). Despite the identification of a number of the regulators
that influence this transition, we do not fully understand how it is
controlled. Detailed analyses of the phenotypes of mutations in the
genes that regulate the transition might provide more insight into the
nature of the decisive events that trigger the onset of DNA
replication. Such analyses of Saccharomyces cerevisiae have
demonstrated two types of phenotypes when entry into S phase is
partially compromised (40, 55). Defects in some genes do not
delay the transition into S phase but prolong S phase, apparently by
reducing replication efficiency. Defects in other genes appear to have
an all-or-nothing effect on a decisive event that is presumed to
represent a switch from the G1 to the S-phase states.
Analysis of the behavior of cells in other organisms (26)
and of nuclei replicating in Xenopus laevis in vitro
extracts (39) suggests that an all-or-nothing switch is
generally associated with the G1-to-S transition.
Several findings argue for the importance of the activation of
transcription of specific genes that accompanies the G1-S
transition. Chief among these is the finding that activation of
specific transcription factors appears to occur immediately upstream of
the decisive transition into S phase in yeast (1, 5),
Drosophila melanogaster (15, 17), and mammalian
cells (37, 44). These factors stimulate a program of
expression of genes whose products are required for DNA replication.
However, because at least some of the required products are stable and
are present as a result of earlier expression, the activation of this
gene expression does not necessarily provide direct coupling between
replication factor transcription and entry into S phase. The role of
triggering entry into S phase may instead be mediated by a second type
of gene product induced as part of the S-phase transcription program. In yeast (43, 45), Drosophila (16),
and vertebrates (4, 23, 47), G1 cyclins are part
of this transcription program (e.g., CLN1 and
CLN2 in S. cerevisiae and cyclin E in
metazoans). It is not entirely clear what roles each of these two
classes of induced gene products, replication factors and
G1 cyclins, plays in the G1-to-S transition.
In mammals and Drosophila, the G1-S-phase
transcription program is controlled by the E2F-DP family of
heterodimeric transcription factors (17, 37, 44, 59). Five
related E2F genes and two related DP genes have been identified in
mammals (59). Mice containing a homozygous knockout mutation
of the E2F-1 gene develop normally and are viable,
indicating that E2F-1 is not essential for cell cycle progression
(22, 72). While E2F-1 knockout mice do display
several abnormal phenotypes, most strikingly the eventual acquisition
of a variety of tumors in older individuals, the absence of a more
dramatic phenotype due to loss of E2F-1 function could be explained by
redundancies among the five known E2Fs. In Drosophila, only
a single E2F gene (dE2F) and DP gene (dDP) have
yet been reported (18, 27, 48), although an
E2F-related sequence has recently been identified as an EST
from a Drosophila embryonic cDNA library (28). As
in mice, therefore, there exists a potential for functional redundancy
among Drosophila E2F genes. Nevertheless, mutations of
dE2F cause lethality (6, 17). In embryos,
dE2F mutations block the activation of transcription that
usually occurs at the G1-to-S transition and inhibit DNA replication (17). Thus, dE2F plays an essential role, likely as a requirement for normal cell cycle progression.
Here we explore further the role of the activation of G1-S
phase transcription by identifying mutations of the dDP gene
and comparing embryonic phenotypes caused by genetic reductions in the
function of dDP, dE2F, or cyclin E, a downstream target of dE2F-dDP
that is required for S phase (16, 33, 51). We find that both
dDP and dE2F are required for the activation of
transcription of the same genes at the G1-to-S transition,
consistent with dE2F and dDP acting as a heterodimer in vivo. If cyclin
E and dE2F-dDP are jointly required to trigger a common step needed for
the initiation of DNA replication, then reductions in the functions of
these gene products should lead to similar phenotypes. In contrast to this, we find distinct behaviors. Reducing dE2F-dDP function first diminishes the rate of DNA replication, and only with more severe reductions in functions do we detect delays or defects in the transition into S phase. Reduction of cyclin E function leads to a
stochastic increase in the length of G1 without
compromising replication in those cells that initiate DNA synthesis. We
discuss these results in terms of two distinct roles for dE2F-dDP and cyclin E in the pulse of transcription at the G1-to-S
transition: one role is a direct and dose-dependent participation in
DNA replication and the other is a regulatory role that triggers an
all-or-nothing change required for S phase.
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MATERIALS AND METHODS |
Genetics and fly stocks.
The overlapping
Df(2R)vg56 and
Df(2R)vg33 deficiencies
(breakpoints 49D;49F and 49D;50A, respectively) and the
recessive-lethal EMS-induced mutations from the 49F region were
isolated in a previously described mutagenesis screen (36).
Single mutant stocks representing a subset of the 18 complementation
groups were screened with the DmRNR2 probe (vr-3, -4, -6, -8, -9, -10, -11, -13, -15, -17, -18, and -19). These were graciously
provided to us by Ting Wu of Harvard University. The
dDPvr10 mutation we refer to throughout the text
was originally designated vr10-13 (36) and contained the
recessive markers al, b, c, and sp. We separated three other lethal mutations from the
original dDPvr10 chromosome by two successive
meiotic recombinations with a wild-type Sevelen second chromosome. The
possibility that this new recombinant chromosome (designated
dDPvr10-5D) only contains the
dDPvr10 semilethal allele is supported by the
rare (<2%) appearance of dDPvr10/dDPvr10 homozygous flies in
the stock. The presence of these three additional lethal mutations did
not significantly affect the phenotype in comparisons of
DmRNR2 staining of
dDPvr10/dDPvr10,
dDPvr10/Df(2R)vg56,
dDPvr10-5D/dDPvr10-5D,
and
dDPvr10-5D/Df(2R)vg56
mutant embryos. The cyclin EP28 and cyclin
E05206 alleles are described in references
2 and 32 and
33, respectively. The dE2F164
P-element allele is described in reference 56; all
other dE2F alleles are described in reference
17. All second chromosome mutant stocks were
maintained with a CyO P[w+;
wg-lacZ] balancer, allowing unambiguous identification of
mutant progeny embryos, which fail to express lacZ.
Molecular analysis of the dDP locus and
dDP cDNA.
Polytene chromosome hybridizations were
performed with digoxigenin-labeled dDP cDNA (encoding amino
acids 76 to 322) exactly as described previously (15). To
identify the dDPvr10 mutation, dDP
genomic DNA was PCR amplified in three overlapping regions with primers
designed from the sequence of the full-length cDNA (see below).
Template DNA was prepared from single adult dDPvr10/Df(2R)vg56
or Canton S control flies as described previously (25). For each genotype, products from two independent PCR amplifications with
Thermococcus litoralis DNA polymerase (New England Biolabs) were subcloned into pBluescript and sequenced. The missense mutation was found in both dDPvr10 PCR amplifications but
not in the wild-type amplifications. Several pieces of evidence
suggested that our original dDP cDNA (18) was not
full length. First, immunoblot analysis revealed that heat shock
expression of this cDNA produces a polypeptide that by sodium dodecyl
sulfate (SDS)-polyacrylamide gel electrophoresis migrates with an
apparent molecular mass ~8 kDa smaller than that of the endogenous
dDP (14). Second, Hao et al. (27)
reported the amino acid sequence of a Drosophila dDP cDNA
that was nearly identical to the previously reported dDP
cDNA translation product (18) from residues 8 to 377 but
contained an amino-terminal extension of 74 amino acids. Third, the
Dynlacht et al. (18) sequence does not contain an in-frame
stop codon upstream of the assigned initiator Met. We PCR amplified the
5' region from a probable full-length dDP cDNA with nested
primers and template DNA prepared from a 0- to 4-h embryonic cDNA
library (7). In the first PCR, a vector primer that
hybridizes 5' to the cloning site and contains a HindIII
recognition sequence was paired with a primer (5'
ACAGTTGTTGTCGTACGCAT) that hybridizes at the unique BsiWI site in the dDP cDNA. A 1:1,000 dilution of
the first reaction mixture was used as the template in a second PCR
that paired the same 5' vector primer with a dDP primer
(5' TTGCCCGATGCCGCTAGCAC) that hybridizes at the unique
NheI site just upstream of the initiator Met assigned by
Dynlacht et al. (18). The single product from this reaction
was cut with HindIII/NheI and subcloned in
frame with the Dynlacht et al. (18) cDNA contained in
pBluescript SK.
Embryo in situ hybridization and BrdU labeling.
Digoxigenin-labeled RNA probes from DmRNR2 genomic DNA
(15) and cyclin E type I cDNA (51)
were used for embryo in situ hybridizations as described previously
(15, 38, 65). Embryos were pulse labeled for 15 min in
Schneider's medium containing 1 mg of BrdU/ml as described previously
(19, 54). Incorporated BrdU was detected with anti-BrdU
mouse monoclonal primary antibody (Becton Dickinson) and
rhodamine-conjugated goat anti-mouse secondary antibody (Jackson). For
heat shock experiments, eggs from dDPvr10/CyO
P[w+; wg-lacZ];
hsp70-dDP/hsp70-dDP flies were collected on grape juice agar
plates, aged at 18°C, and heat shocked by floating the collection
plate on the surface of a 37°C water bath for 30 min. After a 70-min
recovery at room temperature, the embryos were fixed and stained for
DmRNR2 expression. The hsp70-dDP P-element construct is described in reference 14.
Immunoblot analysis.
Eggs were collected for 1 h at
room temperature from w;
Df(2R)vg56/CyO
P[w+; wg-lacZ] flies and aged
23 h at 17°C. The eggs were dechorionated in 2.5% sodium
hypochlorite, and the embryos were devitellinized and fixed by shaking
in a 1:1 mixture of methanol and heptane. The embryos were washed twice
with MeOH, transferred to an aqueous buffer containing protease
inhibitors (10 mM Tris [pH 7.8], 150 mM NaCl, 1 mM EDTA, 1 mM
phenylmethylsulfonyl fluoride, 2 mM benzamidine, 0.05% Tween 20), and
stained with the DNA binding dye Hoechst 33258. Ten
Df(2R)vg56/Df(2R)vg56
or 10 Df(2R)vg56/CyO
P[w+; wg-lacZ] methanol-fixed
embryos were hand selected under UV light with a fluorescence
microscope. The three types of progeny from this stock
{Df(2R)vg56/Df(2R)vg56,
Df(2R)vg56/CyO
P[w+; wg-lacZ], and CyO
P[w+; wg-lacZ]/CyO
P[w+; wg-lacZ]} could easily be
distinguished from each other at germ band-retracted stages due to
differences between the aberrant morphology of the deficiency
homozygotes and the CyO/CyO balancer homozygotes, which have a wg
mutant phenotype. Selected embryos were dissociated by boiling them for
3 min in sample buffer (62.5 mM Tris [pH 6.8], 2% SDS, 5% glycerol,
5% 
-mercaptoethanol) and subjected to SDS-polyacrylamide gel
electrophoresis. Separated proteins were electrophoretically
transferred to nitrocellulose, and dDP polypeptides were detected with
a 1:10 dilution of monoclonal antibody supernatant Yun1
(13).
Nucleotide sequence accession number.
The additional
dDP cDNA sequence discussed above has been submitted to
GenBank (accession no. AF031700).
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RESULTS |
Identification of a dDP mutation.
Drosophila
dDP cDNA was cloned previously and shown to encode a functional
binding partner for dE2F (18, 27). We took a reverse genetic
approach to identify mutations of dDP in order to test its
role in transcription and cell cycle control. Hybridizations of a
dDP cDNA probe to salivary gland polytene chromosomes (Fig. 1A) allowed us to map the dDP
locus to position 49F on the right arm of the second chromosome (not
shown). A series of overlapping deletion mutations that uncover region
49F were generated by Lasko and Pardue (36). In situ
analysis of Df(2R)/+ polytene chromosomes revealed that the
Df(2R)vg56 deletion
chromosome failed to hybridize with our dDP cDNA probe (Fig.
1A). We conclude from this result that dDP coding sequences are missing in the
Df(2R)vg56 deletion. Hao
et al. previously reported that dDP mapped outside this
particular deficiency (27). While we cannot readily explain this discrepancy, two other observations confirm our mapping of dDP within the
Df(2R)vg56 deletion.
First, in embryos that had progressed to cellularization or beyond, in
situ hybridization detected zygotic dDP expression in
control but not in
Df(2R)vg56/Df(2R)vg56
embryos, consistent with the elimination of maternal dDP RNA by cell cycle 14 (data not shown). Second, Western blotting (see Materials and Methods) of total protein extracted from germ
band-retracted Df(2R)vg56/Df(2R)vg56
embryos with anti-dDP monoclonal antibodies indicated a severe reduction in the amount of dDP protein compared to that of
Df(2R)vg56/+ sibling
embryos (Fig. 1B). Taken together these data indicate that the
Df(2R)vg56 chromosome
lacks a functional dDP gene.
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FIG. 1.
Identification of dDP mutations. (A)
Hybridization of a
Df(2R)vg56/+ salivary
gland polytene chromosome with a dDP cDNA probe. The
hybridization signal (arrow) is lost in the
Df(2R)vg56 deletion and
consequently appears over half the width of the heterozygous polytene
chromosome. (B) Western blot of extract of germ band-retracted embryos
collected from the
Df(2R)vg56/CyO stock
probed with anti-dDP monoclonal antibody. Since
Df(2R)vg56 deletes the
dDP gene, any dDP protein present in
Df(2R)vg56/Df(2R)vg56
embryos must be of maternal origin. (C) Germ band-extended control
(+/?) and
Df(2R)vg56/Df(2R)vg56
mutant stage 11 embryos subjected to in situ hybridization with a
digoxigenin-labeled probe derived from the DmRNR2 gene.
DmRNR2 expression is failing in the developing ventral
(toward the bottom) nerve cord of the mutant embryo. (D) Intron and
exon structure of the transcribed region of the dDP gene.
Exons are represented by shaded boxes, and introns are represented by a
straight line. ATG and TGA indicate the beginning and end of the
dDP open reading frame, respectively. The Arg-to-His
missense mutation in exon 5 of the dDPvr10
allele is indicated. (E) Alignment of a highly conserved region of the
heterodimerization domain of DPs from Drosophila (this
paper), human (hum) (30, 70), mouse (mus) (30,
49), and Xenopus (xen) (24). The numbers
indicate the position in each primary amino acid sequence. The position
of the Arg-to-His change in the dDPvr10 allele
is indicated at the top.
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Mutations of dE2F reduce the zygotic expression of several
genes involved in DNA replication, including those encoding the small
subunit of ribonucleotide reductase (DmRNR2) and
PCNA (17). This phenotype is first apparent in
the central nervous system (CNS) of germ band-extended embryos during
late stage 11. In germ band-extended
Df(2R)vg56/Df(2R)vg56
embryos the amount of DmRNR2 and PCNA mRNA in the
CNS was significantly reduced relative to that of the wild type (Fig.
1C). An essentially identical gene expression defect was observed in
embryos homozygous for
Df(2R)vg33 (not shown),
another deletion mutation isolated by Lasko and Pardue that removes
polytene region 49F (36). Therefore, loss of the
dDP locus is associated with a phenotype that closely
resembles the phenotype associated with the loss of dE2F
function.
Embryos homozygous for the
Df(2R)vg56 and
Df(2R)vg33 deletions
deviate dramatically from wild-type morphology during and after germ
band retraction, presumably because of the loss of multiple gene
functions. This precluded an accurate analysis of gene expression patterns in the deficiencies at stages when we usually analyze the
G1-S transcription program in wild-type embryos. We
therefore examined existing mutations that mapped to 49F for
perturbations in embryonic DmRNR2 expression. Mutations
causing defective DmRNR2 expression would be candidates for
alleles of dDP. This approach had proven successful in
identifying dE2F mutations (17). A collection of
EMS-induced mutations representing 18 lethal complementation groups
that fail to complement
Df(2R)vg56,
Df(2R)vg33, and other
deletions in the 49F region were generated by Lasko and Pardue
(36). Embryos were collected from extant stocks carrying representative alleles from most of these complementation groups (see
Materials and Methods) and probed for DmRNR2 expression by in situ hybridization. In one lethal complementation group, designated dDPvr10, DmRNR2 expression was
greatly reduced compared to that of wild-type embryos (for germ
band-retracted stages, cf. Fig. 2A and
B). No obvious change in the expression of DmRNR2 was
observed in any of the other mutants tested. At germ band-extended
stages, the dDPvr10 DmRNR2 expression phenotype
was similar to that caused by
Df(2R)vg56 and
Df(2R)vg33 (not shown),
suggesting that the dDPvr10 chromosome contained
a mutation in the relevant gene deleted in these deficiencies.
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FIG. 2.
dDP is required for expression of
DmRNR2. Each panel shows a stage 13 embryo (anterior at the
left and dorsal at the top) subjected to in situ hybridization with a
digoxigenin-labeled probe derived from the DmRNR2 gene. (A)
Wild type (WT). The pattern of DmRNR2 expression is
coincident with the pattern of DNA replication (Fig. 5A). This includes
internal endoreduplicating cells (e.g., midgut cells) and proliferating
cells in the CNS. (B)
dDPvr10/dDPvr10 (dDP ).
DmRNR2 expression is not activated in the dDP
mutant. (C) A dDPvr10/dDPvr10;
hsp70-dDP/hsp70-dDP (dDP hs-dDP) embryo
subjected to a 30-min 37°C heat shock followed by a 70-min recovery
at room temperature. Expression of dDP cDNA in the mutant
restores the normal pattern of DmRNR2 expression. Each of
the images has an internal focal plane in order to observe the anterior
midgut (arrows).
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The original complementation mapping by Lasko and Pardue
(36) placed the dDPvr10 lethal
mutation outside of the
Df(2R)vg56 deletion; i.e.,
dDPvr10 and
Df(2R)vg56 were scored as
complementing mutations in their test crosses. Since this was
inconsistent with our interpretation of the DmRNR2 expression phenotype, we repeated the complementation tests with our
dDPvr10 stock (Table
1). When crosses with
Df(2R)vg56 flies
were reared at 25°C, less than 2% of the adult progeny (n = 683) were of the
dDPvr10/Df(2R)vg56
class (complementing mutations should comprise 33% of adult progeny). Similarly, in crosses with
Df(2R)vg33 flies less than
1% of the adult progeny (n = 531) were of the dDPvr10/Df(2R)vg33
class. These rare "escapers" were not phenotypically normal: many
flies had notched wings, some had rough eyes, and both males and
females were sterile (the females failed to lay eggs). Thus, the
dDPvr10 chromosome is semilethal at 25°C over
both deficiencies. Because this semilethality gives occasional
survivors, it could be scored as complementation when smaller numbers
of progeny are scored: presumably this explains the mapping results of
Lasko and Pardue. When we reared our crosses at 29°C, no
Df(2R)/dDPvr10 flies
eclosed (Table 1). A single copy of a hsp70-dDP construct contained in a P-element vector was sufficient to induce
dDPvr10/Df(2R)vg56
or
dDPvr10/Df(2R)vg33
flies to eclose at 29°C in the absence of a 37°C heat shock
treatment (Table 1). We conclude that the
dDPvr10 mutation fails to complement
deficiencies that delete the dDP locus and that low-level
expression of a dDP cDNA can rescue
dDPvr10/Df(2R) lethality.
Additionally, the incomplete lethality of the dDPvr10/Df(2R) heteroallelic
combination at lower temperatures indicates either that dDP
is not absolutely essential or that dDPvr10 is
not null for the locus.
These data suggested that the dDPvr10 chromosome
contained a mutation in the dDP gene. To confirm this, we
molecularly characterized the dDP locus of wild-type and
dDPvr10 chromosomes. The full-length
dDP transcription unit spans approximately 4 kb of genomic
DNA and contains eight introns (Fig. 1D). Our analysis predicts that
the wild-type dDP translation product consists of 446 residues with a predicted molecular mass of 49,718 Da. This is in
contrast to previously published reports (see Materials and Methods).
Southern blotting indicated that there was no gross rearrangement of
the dDP locus in the dDPvr10
chromosome (data not shown). PCR amplification and sequencing of the
dDP locus from individual
dDPvr10/Df(2R)vg56
or dDPvr10/dDPvr10 adult flies
revealed a single nucleotide change encoding an Arg-to-His substitution
at residue 217 (Fig. 1E). This nucleotide change destroys a unique
AatII restriction site present in our dDP cDNA. This consequently provided a means of readily assessing whether the
change is simply a wild-type polymorphism. PCR amplification products
of dDP genomic DNA isolated from
dDPvr10/CyO flies could be partially digested by
AatII, indicating that the CyO balancer chromosome contains
the AatII site (not shown). In contrast, similar
dDP genomic PCR products derived from a stock (vr18/CyO) containing a mutant chromosome (vr18)
independently isolated in the original Lasko and Pardue (36)
mutagenesis screen could be completely digested by AatII,
strongly suggesting that the vr18 chromosome contains the
AatII site (not shown). Furthermore, vr18
complements dDPvr10 and does not cause defects
in DmRNR2 expression (not shown). We conclude from this data
that the Arg-to-His amino acid change present in
dDPvr10 is not a polymorphism that was present
in the flies originally mutagenized by Lasko and Pardue
(36).
Arg 217 is in a region of the dDP protein that is highly conserved
among Xenopus, mouse, human, and Drosophila DPs
(Fig. 1E). This region of mammalian DP-1 has been shown to be required
for dimerization with E2F-1 (3, 30, 34, 69). Thus, the
Arg-to-His mutation may reduce the ability of the dDPvr10
polypeptide to bind to dE2F. Dimerization of dDP with dE2F is required
for efficient DNA binding activity in vitro (18). Moreover, coexpression of dDP and dE2F is needed for
effective transactivation of promoters containing dE2F-dDP binding
sites in cultured cells (13, 18) and for stimulating
transcription of endogenous target genes in embryos (14). We
conclude that the dDPvr10 chromosome contains an
alteration of the dDP gene and suggest that this change
impairs the ability of dDP protein to act as a transcription factor,
perhaps due to a reduced capacity to dimerize with dE2F. Royzman et al.
(52) have also recently reported the characterization of a
dDP mutant allele (a1) encoding an Arg-to-Cys change at residue 217.
Since dDPvr10 encodes a point mutation of dDP,
we cannot be sure that the phenotypes we describe below represent the
null situation. Moreover, we could still detect a small amount of
maternal dDP protein in germ band-retracted
Df(2R)vg56/Df(2R)vg56
embryos (Fig. 1B). Nevertheless, we did not observe obvious differences in the transcription and DNA replication phenotypes in
dDPvr10/dDPvr10 and
dDPvr10/Df(2R)vg56
embryos. This indicates that a twofold reduction in the zygotic dose of
dDPvr10 does not detectably enhance the
phenotypes we are scoring and suggests that dDPvr10 retains
very little function.
dDP is required for the expression of replication genes
at the G1-S transition.
The first 16 cell cycles of
Drosophila embryogenesis lack a G1 phase.
Following the introduction of a G1 phase in cell cycle 17, subsequent entry into S phase is accompanied by coordinate dE2F-dependent transcriptional activation of replication
factor genes (15). We used probes derived from
DmRNR2 or cyclin E as representatives of this
group of genes (cf. Fig. 2A and 4A). DmRNR2 transcripts are
present continuously and nearly uniformly prior to cycle 17 in
dDPvr10 mutants, as they are in the wild type
(not shown). By cycle 17, DmRNR2 is not expressed in its
usual stereotypic pattern in endoreduplicating tissues in the mutants
(cf. Fig. 2A and B). The dDPvr10 mutation also
dramatically reduces DmRNR2 expression in
proliferating cells of the developing CNS (brain lobes and ventral
nerve cord). Heat shock expression of a dDP cDNA contained
in a P-element transgene restored the wild-type pattern of
DmRNR2 expression (Fig. 2C). We conclude that dDP
function is required for the activation of DmRNR2
transcription in both endoreduplicating and proliferating cells.
The pattern of cyclin E expression resembles that of genes
encoding other replication factors during cycle 17 (16, 33). However, while genes encoding other factors (e.g., DmRNR2)
require dE2F function in all domains of their expression
(16), cyclin E expression relies on
dE2F in all domains except the CNS (16). Similarly, expression of cyclin E relies on dDP
in all domains but the CNS (cf. Fig. 3A
and B). The factors that control cyclin E expression in the
CNS have not yet been identified.
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FIG. 3.
dDP is required for cyclin E
expression at the G1-S transition in endoreduplicating
cells. Each panel shows a stage 13 embryo (anterior at the left and
dorsal at the top) subjected to in situ hybridization with a
digoxigenin-labeled probe derived from cyclin E cDNA. (A)
Wild type (WT). The arrows indicate the midgut. The arrowhead indicates
the ventral nerve cord. (B)
dDPvr10/Df(2R)vg56
(dDP). The cells of the midgut normally express
cyclin E at this stage but fail to do so in the
dDP mutant. CNS expression of cyclin E is normal
in the dDP mutant. (C) cyclin E5206/cyclin
E5206 mutant embryo. This allele contains a P-element
insertion into the upstream regulatory region of the cyclin
E gene (33). cyclin E is not expressed in
the midgut or other cells that normally endoreduplicate but is
expressed in the CNS. (D) cyclin E5206/cyclin
EP28 transheterozygote. Expression of cyclin
E, presumably from the cyclin EP28 homolog,
initiates at the correct developmental time in the midgut. However,
transcription is not down regulated as usual, leading to persistently
high levels of cyclin E mRNA.
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Our results indicate that the dDP mutant phenotype is very
similar to that of dE2F mutants with respect to activation
of transcription of DmRNR2 and cyclin E. This is
consistent with a requirement for dE2F-dDP heterodimers in the
transcription of replication genes in vivo. However, if dE2F
and dDP perform any functions independent of each other,
then we would also expect to observe differences in their respective
phenotypes. This possibility is further suggested by the presence of at
least one other E2F-like gene, and therefore at least one
other potential dDP binding partner, in Drosophila
(28). We detected a phenotypic difference in the embryonic
CNS with a probe derived from the Drosophila dMCM3 gene (22a, 64). dMCM3 is a member of the MCM family of proteins, which act at origins of replication and are required for S phase (8, 61, 67). dMCM genes are expressed in
replicating cells in a pattern similar to that of genes like
DmRNR2 (21, 66). In dE2F mutant
embryos, there is substantially less dMCM3 RNA in the CNS
than in the wild-type CNS, indicating that dE2F is required
for dMCM3 expression in this tissue (cf. Fig.
4A and C). The
dDPvr10 mutation also alters dMCM3
expression, but in very different way. The intense staining of
particular cells seen in wild-type embryos is absent in the mutant,
suggesting that this high-level expression requires dDP (Fig. 4B).
However, other cells of the CNS that normally have extremely low levels
of dMCM3 RNA exhibit increased staining (Fig. 4B). This
derepression of dMCM3 as a result of mutation of
dDP suggests that dDP normally plays a role in the
repression of this gene, at least in some cells of the CNS.
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FIG. 4.
Mutation of dDP and dE2F causes
distinct phenotypes. Each panel shows a stage 14 embryo (anterior at
the left) subjected to in situ hybridization with a digoxigenin-labeled
probe derived from dMCM3 cDNA. The perspective is ventral to
view the nerve cord (VNC) of the CNS (arrowheads). (A) Wild type (WT).
Expression occurs coincident with replication, generating the observed
pattern. (B) dDPvr10/dDPvr10 mutant.
dMCM3 expression continues in the VNC, but occurs in an
expanded domain of cells. This suggests that dDP functions to prevent
dMCM3 from becoming derepressed in this tissue. (C)
dE2F91/dE2F91 null mutant. Very
little if any dMCM3 expression can be detected in the VNC,
indicating that dE2F is required for normal dMCM3
expression. No ectopic dMCM3 expression is observed, perhaps
because dE2F does not perform an essential repression function.
Residual dMCM3 expression in the brain accounts for the
out-of-focus staining at the left.
|
|
Mutation of dDP attenuates DNA replication and
lengthens S phase.
Replicating nuclei can be readily observed in
situ by pulse labeling live embryos with BrdU and subsequently
detecting BrdU incorporation immunologically after fixation. The
wild-type profile of endocycle S phases during embryonic stages 12 to
15 in the midgut has been well described (15, 60). The first
of these S phases begins from G1 of cycle 17 (Fig.
5A).
dDPvr10/dDPvr10 and
dDPvr10/Df(2R)vg56
mutant embryos incorporated much less BrdU in the midgut during a
15-min pulse labeling than did wild-type embryos (for
dDPvr10/dDPvr10, cf. Fig. 5A and B).
Reduced BrdU incorporation is also seen in other endocycling tissues,
such as the hindgut. This decrease in BrdU incorporation was not
restricted to endocycles, as mitotic cells in the CNS also labeled
poorly relative to the wild type (cf. Fig. 5A and B). We conclude that
dDP function is required for wild-type levels of DNA
synthesis in endocycling and dividing cells.
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FIG. 5.
Different cell cycle roles for cyclin E and dDP. Embryos
were collected and aged until stage 13 (A to C) or stage 14 (D to F),
pulse labeled for 15 min with BrdU, and then immediately fixed. BrdU
that incorporated into replicating nuclei during the pulse was
subsequently detected by indirect immunofluorescence. The orientation
of each embryo is the same as in Fig. 2. (A and D) Wild type (WT). (B
and E) dDPvr10/dDPvr10 (dDP). (C and
F) cyclin EP28/cyclin E05206 (cyc
E). The arrows indicate the cells of the anterior midgut (AMG). These
cells begin S phase 17 during stage 12, continue replication into stage
13 (A), and exit S phase by stage 14 (D). In the dDP mutant,
all or most of the AMG cells enter S phase at the correct time but
incorporate much less BrdU (B). By stage 14 these cells are still
replicating (E), indicating that S phase is longer than it is in the
wild type. In the hypomorphic cyclin E mutant, the AMG cells do not
begin S phase 17 on schedule (C) but eventually enter S phase in a
random fashion beginning in stage 14 (F). Thus, G1 of cycle
17 is longer than usual in these cells. The actual length of
G1 depends upon when the cells reach the critical level of
cyclin E activity that triggers S phase. Also note that replication in
the CNS appears normal, consistent with wild-type cyclin E expression
in the CNS of cyclin E05206 mutants.
|
|
Analysis of BrdU incorporation in an embryo allows one to score the
timing of entry into S phase and the duration of S phase in addition to
scoring the level of labeled nucleotide incorporated during the pulse.
As described above, the dDPvr10 mutant embryos
exhibit an obvious reduction in BrdU incorporation, but the pattern of
BrdU incorporation reveals that the programming of the
G1-to-S transition still operates. The tissues that enter S
phase 17 relatively early in the wild type, such as the anterior midgut, posterior midgut, and hindgut, appear to begin their
compromised S phase at a similar embryonic stage in the dDP
mutant (Fig. 5B). This demonstrated that reduction of dDP
function first affects the level of incorporation rather than the
timing of the transition from G1 to S phase. Analysis of
the incorporation pattern in successively older embryos, whose age is
determined both by time from egg deposition and by staging according to
morphological criteria, defines the duration of S phase. Whereas the
tissues that begin S phase 17 early in wild-type embryos discontinue
incorporation by stage 14, in the mutant incorporation continues at
this stage. This changes the pattern of BrdU incorporation such that
cells which no longer label in the wild type are still labeled in the
mutant (e.g., anterior midgut cells [Fig. 5D and E]). The
prolongation of S phase combined with decreased BrdU incorporation
suggests that the replication rate is reduced. Since the
dDPvr10 mutant compromises the expression of
genes such as DmRNR2, we suggest that the mutant exhibits a
reduction in DNA synthesis as a result of decreases in the levels of
replication functions.
Our original analysis of the replication phenotype in dE2F
mutant embryos led us to the conclusion that the mutation blocked S
phase (17), whereas the analysis of the dDP
mutation suggests a more quantitative defect. We thus conducted more
detailed analyses of dE2F mutants. The results suggest that
reductions in the function of dE2F also have a quantitative
effect on replication. dE2F164 is a homozygous,
viable, hypomorphic P-element-induced allele of dE2F that
expresses nearly undetectable levels of dE2F mRNA (56). Nonetheless, dE2F164 fully
complements a deletion
[Df(3R)eD7] of the
dE2F gene: adult flies are viable and fertile. Despite the
absence of an effect on viability, we detected a change in the pattern
of BrdU incorporation in
Df(3R)eD7/dE2F164
mutant embryos. As with the dDPvr10 mutant, this
pattern is consistent with a prolongation of S phase 17 in the midgut
(cf. Fig. 5E and 6A). This suggests that
the requirement for dE2F can be satisfied by relatively low
levels of activity and that a reduction in the efficiency of S phase is
not lethal. We therefore reexamined embryos homozygous for lethal null
alleles of dE2F. We could detect a low level of BrdU incorporation in the midgut, as well as in the CNS (Fig. 6B). This
incorporation can be more readily detected when longer BrdU-labeling periods are used (e.g., 40 vs. 15 min). Again, the pattern of BrdU
incorporation was altered in a way that suggests the prolongation of S
phase 17 in the midgut (cf. Fig. 5E and 6B). We conclude that
reductions in function of dE2F/dDP activity over a large range result in a graded reduction in S phase.
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FIG. 6.
Prolonged S phase in dE2F mutants. Stage 14 dE2F164/Def(3R)eD7
(hypo) (A) and dE2F7172/dE2F7172
(null) (B) embryos pulse labeled for 15 min with BrdU. S phase 17 continues in the anterior midgut (arrows) when in the wild type it has
already finished (Fig. 5D). This is an indication that S phase is
prolonged in the mutants. In the central portion of the midgut
(arrowheads) S phase 18 has not started on schedule for most of the
cells (Fig. 5D). This indicates that dE2F also contributes
to correct timing of the G1-S transition.
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|
Cyclin E triggers S phase in an all-or-nothing manner.
During
G1 phase, eukaryotic cells are thought to respond to
growth-promoting signals by making a commitment to enter the cell cycle
and begin S phase. A typical feature of the process of commitment is
that it is an all-or-nothing step. Reductions in function of a
component contributing to such an all-or-nothing step ought to exhibit
a sharp threshold, with reductions below a critical level blocking the
ability to progress beyond the decisive step. As described above,
mutations that compromise the activity of dE2F-dDP suggest that this
transcription factor makes a graded contribution to the efficiency of S
phase, the antithesis of the expectation for a regulator of a decisive
all-or-nothing transition.
Cyclin E mutations on the other hand show the behavior expected for a
contribution to an all-or-nothing transition. Cyclin E null mutations
complete embryonic cell cycle 16 and cause an arrest in G1
of cell cycle 17 (33). Our immunofluorescent methods cannot
detect any BrdU incorporation in these mutants. Thus, in this instance
the cyclin E threshold is never reached. To test the consequence to
cell cycle progression of reduced but significant levels of cyclin E
function, we used two hypomorphic mutations of cyclin E: cyclin
E05206, a P-element insertion mutation that eliminates
cyclin E expression in endocycling cells but not in the CNS (Fig. 3C),
and cyclin EP28, an EMS-induced mutation. We
examined BrdU incorporation in the anterior midgut cells of
cyclin E05206/cyclin EP28
transheterozygous mutant embryos. At stage 13, no BrdU incorporation was detected (Fig. 5C). These cells are therefore still in
G1 of cycle 17 at a time when wild-type cells have already
begun S phase 17 (Fig. 5A). However, at stage 14 we could detect BrdU incorporation in some of the anterior midgut cells (Fig. 5F). The
number of BrdU-positive cells varied from embryo to embryo, and the
distribution of cells within the tissue was random. In sharp contrast
to the results with the dDP and dE2F mutations, the level of BrdU staining in those nuclei that had made the transition to S phase was just as intense as in the wild type. We interpret this
phenotype as a delay of the G1-S transition, or a reduction of the probability of undergoing this transition, and suggest that it
reflects a partial defect in the execution of an all-or-nothing event
leading to S phase (Fig. 7).
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FIG. 7.
Model of dE2F-dDP and cyclin E function at the
G1-S transition and schematic representation of the
G1-S transition and BrdU incorporation in the embryonic
midgut. (A) Wild type. Components of the prereplicative complex (open
symbols) assemble on chromosomes (parallel lines) at origins of DNA
replication (solid rectangles). Since E2F-DP is required for the
transcription of genes encoding known components of prereplicative
complexes like dMCM3 in Drosophila and HsOrc1 in humans
(46), we propose that dE2F-dDP contributes to prereplicative
complex assembly. Cells remain in G1 until they are induced
to progress through the cell cycle by a developmental signal. DNA
synthesis is then triggered at the beginning of S phase via cyclin E
action, and bidirectional replication forks appear in the chromosome
(ovals). After BrdU pulse labeling and in situ immunofluorescent
detection of incorporated BrdU, the nuclei in a field of cells that
synchronously entered S phase stain brightly and uniformly (nine open
circles on the black background). (B) Reduced cyclin E function.
Prereplicative complexes form normally at a frequency identical to that
of the wild type, because dE2F-dDP is activated independently of cyclin
E (16). If cyclin E function is eliminated, replication is
not triggered and cells remain in G1. If the cyclin E
function is reduced but not eliminated, S phase will still be triggered
provided cyclin E function can eventually achieve the critical
threshold. In the latter situation, G1 is prolonged and DNA
replication is normal once S phase begins. In a field of cells of this
type, the length of G1 might vary stochastically as cells
approach the S phase threshold. The pattern of BrdU-labeled nuclei
would appear random, and each nucleus would have the wild-type
intensity of staining (three open circles instead of nine on the black
background). (C) Reduced dE2F-dDP function. Decreased provision of
components causes limited assembly of prereplicative complexes. If
those that assemble are triggered at the usual time in development, the
length of G1 does not change. However, fewer prereplicative
complexes initiate fewer bidirectional replication forks. Consequently,
S phase is prolonged and BrdU incorporation during pulse labeling is
significantly reduced. A field of cells of this type appears uniformly
and weakly labeled (nine shaded circles on the black background). (Note
that the symbols designating components of the prereplicative complex
are not shown on the DNA after bidirectional DNA synthesis has begun
for the purpose of clarity. Some components of this complex [e.g.,
ORC] are thought to remain bound to the DNA throughout the entire cell
cycle [11].)
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|
Strikingly, a small number of cyclin E05206/cyclin
EP28 progeny eclosed as adult flies (12 out of 317 total progeny [3.7%]). These flies were sterile, had rough eyes, and
had incomplete formation of the fifth wing vein. Therefore, this
combination of cyclin E alleles is semilethal. This result
indicates that dramatic alterations to the timing of the
G1-S transition at embryonic stages is still compatible
with a substantial amount of development.
Drosophila cyclin E inhibits its own expression in
endocycles by one or more negative feedback loops (16, 41,
53), and mammalian cyclin E has been shown to promote its own
destruction (9, 68). As a result, cyclin E is ordinarily
expressed only transiently at the G1-S transition. We
examined the behavior of cyclin E expression in cyclin
E05206/cyclin EP28 embryos and found that
these embryos activate cyclin E transcription (presumably
from the mutant cyclin EP28 chromosome) at the
correct time in cycle 17 in the midgut but fail to terminate
transcription at the normal time (Fig. 3D). This, and similar results
with cyclin E null mutants (16, 53), suggests
that the normal down regulation of cyclin E synthesis involves negative
feedback. There appear to be multiple components to this negative
feedback. Cyclin E acts particularly effectively as a repressor of its
own expression (53), and in a separate step it plays a role
in the down regulation of dE2F-dDP-dependent transcription in general
(16). This mode of regulation buffers the effect of mutation
on cyclin E function in that mutant alleles of cyclin E are
expressed for a longer time and mRNA (and presumably the mutant protein
it encodes) can accumulate to higher levels (Fig. 3D). Therefore, we
propose that part of the delay in progress to S phase in the
cyclin E05206/cyclin EP28 mutant is
due to an extension in the time required for mutant cyclin E protein to
achieve a critical threshold of cyclin E function (e.g., activating
cdk2).
dE2F/dDP contributes to the all-or-nothing transition into S
phase.
Based on the characteristic defects seen in hypomorphic
conditions for cyclin E and dE2F, and in the
dDPvr10 mutant, it appears that cyclin E has
important inputs into an all-or-nothing event required for transition
to S phase while dE2F-dDP makes a quantitative contribution to
functions required for S phase. However, our work has demonstrated that
cyclin E expression requires dE2F and dDP at the
G1-S transition of cycle 17. We consequently expected that
dE2F-dDP should also make a contribution to the G1-S phase
trigger. Such a contribution might be masked if a relatively high level
of dE2F-dDP activity were required to fully satisfy the quantitative
contribution to S phase while a very low level of activity of dE2F-dDP
provided sufficient cyclin E to satisfy the threshold of the
all-or-nothing event. Indeed, further characterization of the
dDP and dE2F mutant phenotypes suggests that this
is the case.
As we described above, cells that enter S phase 17 relatively early
(e.g., anterior midgut cells) in the wild type do so at approximately
the normal time in the dDPvr10 mutant. However,
the ensuing S phase is prolonged in the mutant and displays a reduced
rate of BrdU incorporation. If a low level of maternal dDP persists
until the time that tissues begin to enter S phase 17, one might expect
that, as a result of continued decay, cells which enter S phase early
would benefit from a higher level of this maternal dDP than cells which
enter S phase at a later stage. Consistent with this, S phase 18 in the
central midgut (Fig. 5D), which begins about 2 h after the
beginning of S phase 17 in the anterior midgut, seems to be more
dramatically affected in the dDPvr10 mutant
embryos (Fig. 5E). Importantly, the timing of entry into S phase 18 is
affected as well as the efficiency of replication. In stage 14, only a
few cells in the mutant's central midgut have begun to incorporate
BrdU (Fig. 5E) compared to the number in wild-type embryos (Fig. 5D).
These cells have either been arrested or they initiate S phase at a
much later time than the wild type (i.e., beyond the stages we have
examined). Moreover, the few cells that do incorporate BrdU are stained
weakly, again consistent with a reduced rate of DNA replication once S
phase is initiated. A very similar phenotype is seen in the same cells
of dE2F mutant embryos (Fig. 6). These data suggest that
dE2F-dDP contributes both to the correct timing of the G1-S
transition and to the efficiency of S phase.
| |
DISCUSSION |
In this report we describe the identification and characterization
of mutations of the Drosophila dDP gene, which encodes a
component of the E2F-DP transcription factor. dDP is
required for activation of expression of replication genes at the
G1-S transition in embryonic cells, and mutants have an
attenuated and sometimes delayed S phase. Characterization of BrdU
incorporation in different mutant backgrounds and at different
embryonic stages suggests that reductions in dE2F-dDP over a large
range results in graded attenuation of DNA replication and that only
severe reductions in dE2F-dDP function delay or block entry into S
phase. In contrast, partial loss of function of cyclin E results in a delay of S-phase entry without detectable diminution of BrdU
incorporation. We suggest that dE2F-dDP makes two different kinds of
contributions to DNA replication. It drives the expression of numerous
replication functions, some of which are limiting for the rate of DNA
replication. Additionally, it drives the expression of at least one
product, cyclin E, that is required to trigger an all-or-nothing
transition from G1 into S phase.
dE2F and dDP collaborate to activate G1-S
transcription.
A substantial amount of biochemical evidence
indicates that E2F-DP heterodimers function as a DNA binding
transcription factor that contributes to the activation of
transcription at the G1-S transition (reviewed in
references 37 and 59). Our
previous genetic analysis had demonstrated that the Drosophila
dE2F gene is essential for transcriptional activation of several
replication genes at the G1-S transition in embryos. The
present work demonstrates that mutation of the Drosophila
dDP gene also eliminates this activation of expression for the
DmRNR2 and cyclin E genes. While this article was
undergoing review, Royzman et al. reported similar transcriptional
phenotypes due to mutation of dDP (52). The parallel defects in the transcription of several genes are consistent with the joint action of dE2F and dDP in this aspect of their function.
Several experiments imply that transcriptional activation is likely a
result of a direct interaction between dE2F-dDP and cis-acting elements in the promoters of replication genes.
Expression of the dE2F cDNA stimulates the DNA polymerase
promoter in cultured Drosophila cells (48).
Furthermore, dE2F-dDP binding sites found upstream of the
PCNA transcriptional start site are required for reporter
gene expression in cell culture and -galactosidase activity in
embryo extracts prepared from flies carrying
PCNA-promoter-lacZ fusion transgenes
(71).
Is normal regulation of S phase essential?
The vr10
allele of dDP produced some adult flies when it was
heterozygous to a deficiency for the locus. While it is possible that
this result is in part due to residual function of this mutant allele,
it is clear that this same allele reduces G1-S
transcription of various replication genes to levels below detection
and substantially compromises replication in the embryonic S phases
that we have examined. Similarly, a dE2F hypomorphic allele
that alters embryonic S phases is viable, and a combination of
cyclin E alleles that greatly altered the embryonic S-phase
program (Fig. 5F) nonetheless gave viable flies. Based on these
observations, we suggest that most aspects of patterning, growth, and
differentiation are regulated in ways that are not easily disturbed by
alterations in the G1-S program. Note that adult
dDPvr10 mutant flies were rare, defective, and
infertile. Hence the activity of the gene and presumably the control of
S phase is clearly of some relevance to development, but it is
nonetheless surprising that an intact fly can be produced when defects
in S phase can be scored in midembryogenesis.
Is dE2F-dDP activity essential to S phase?
We previously
reported that the compromised embryonic S phases in a dE2F
mutant indicated that dE2F function was required for the
G1-S transition (17). However, here we have
shown that the available dE2F and dDP mutations
have their primary effect on the efficiency of S phase. If dE2F-dDP is
an integral part of the normal cascade of signaling that triggers S
phase, we might expect that the mutations would result in an absolute
block to S phase rather than a reduction in its efficiency. However, we find that many cells enter a compromised S phase 17 according to a
normal schedule. Nonetheless, we cannot conclude that dE2F-dDP function
is dispensable for the G1-S transition, because it is possible that the threshold of dE2F-dDP activity required to drive the
transition is lower than the levels of dE2F-dDP required for maximal
efficiency of DNA replication.
There are three possible sources of dE2F-dDP activity in the mutant
embryos. First, it appears that a small amount of maternally expressed
protein persists to late embryonic stages (Fig. 1B). Second, the
mutations might not remove all activity of the gene. Third, there may
be homologs that provide a comparable activity. Indeed, sequences of
random cDNAs (ESTs) by the Berkeley Drosophila Genome
Project uncovered the existence of a second dE2F homolog (28). This and perhaps other undetected homologs of dE2F
and/or dDP might contribute to dE2F-dDP activity in the mutant
backgrounds. Hence, while our experiments allow us to say that
mutations that reduce dE2F-dDP activity affect S phase, it is more
difficult to address whether DNA replication seen in these mutant
backgrounds depends on a low level of residual dE2F-dDP activity from
one or more of these sources.
One important observation leads us to suggest that dE2F-dDP contributes
to the transition from G1 to S as well as to the efficiency of DNA replication. Analysis of a later S phase (S phase 18 in the
central midgut), when maternal functions have further decayed, shows
that entry into S phase is delayed and variable. Additionally, an
involvement of dE2F-dDP in the G1-S transition is also
suggested by the involvement of the dE2F-dDP-stimulated gene
cyclin E in triggering the G1-S transition (see
below) and by the extraordinary correlation between the schedule of
activation of dE2F-dDP and the developmental program of S phases. We
consequently favor the interpretation that dE2F-dDP drives expression
of cyclin E at the G1-S transition and that
cyclin E is essential for this transition.
Our favored interpretation might seem strained because it requires that
undetectable levels of cyclin E RNA trigger S phase. However, the genetics indicates that zygotic expression of cyclin E is absolutely required for S phase 17 (33). Hence,
regardless of our failure to detect cyclin E RNA in the
dE2F and dDP mutant embryos, sufficient cyclin E
protein must accumulate to allow S phase. How can undetectable levels
of cyclin E mRNA produce sufficient cyclin E protein to
trigger S phase? We suggest that particular aspects of cyclin E
regulation might allow gradual accumulation of cyclin E protein at very
low levels of mRNA. Cyclin E inhibits its own dE2F-dDP-mediated
transcription in embryos (16, 53). Moreover, mammalian
cyclin E protein appears to drive its own destruction through
ubiquitin-mediated proteolysis (9, 68). Hence, once
expression begins cyclin E will accumulate until it reaches levels that
activate these negative feedback loops. As the level of cyclin
E RNA is reduced, the rate of accumulation of cyclin E protein
will decline, but it will accumulate over a longer period and
ultimately reach its threshold levels. Accordingly, we suggest that
severe reductions in dE2F-dDP activity would be required before one
could detect a substantial defect in the G1-S transition.
In addition, it is also possible that in the complete absence of
dE2F-dDP, low basal expression of cyclin E might allow accumulation of cyclin E protein over a very long time. In this case,
entry into S phase would not be timed properly, but could still occur.
Such dE2F-dDP-independent expression of cyclin E might
sustain the growth of dE2F and dDP mutant larvae,
as reported by Royzman et al. (52).
The dE2F-dDP contribution to the efficiency of DNA
replication.
In S. cerevisiae, limitation of components
that bind replication origins (e.g., ORC and MCM proteins) causes a
reduced rate of replication that is associated with reduced origin
utilization (reviewed in reference 11). In
Drosophila, reduction in the function of dMCM2 results in a
prolongation of S phase consistent with such a reduction in origin
firing (66). We suggest an analogous interpretation of the
phenotype caused by reduction of dE2F-dDP function (Fig. 7). In both
the dDP and dE2F mutant embryos the first defect
in DNA replication that appears is a reduction in the intensity of BrdU
incorporation. An increase in the length of S phase 17 observed in the
midgut argues that this decreased incorporation is associated with a
reduced rate of DNA replication. Although we have not identified a
specific factor whose expression is rate limiting for these embryonic S
phases, we have demonstrated that dE2F and dDP
are required for the expression of several genes encoding replication
factors. We propose that one or more replication factors become
limiting when dE2F-dDP function is compromised and that the consequence
is a reduction in the number of forks that successfully initiate
replication.
Cyclin E provides an S-phase threshold.
Evidence from many
different systems indicates that cyclin E-cdk2 activity is required for
cell cycle progression (reviewed in reference 29).
In what way is cyclin E-cdk2 activity required for S phase? Here we
have shown that Drosophila cyclin E provides a threshold for
entry into S phase. By reducing but not eliminating cyclin E function
by a hypomorphic combination of alleles, we showed that
endoreduplicating embryonic midgut cells fail to execute a
developmentally controlled G1-S transition on schedule in
cell cycle 17. But rather than remaining in G1 arrest
indefinitely, these cells subsequently enter S phase. In this delayed S
phase, BrdU incorporation appears normal, probably because dE2F-dDP is independently activated in these cells and the replication machinery other than cyclin E is not limiting (17). The midgut cells
do not all enter S phase at the same time, however, but rather at random times. We interpret this stochastic entry into S phase in cycle
17 as evidence for the participation of cyclin E in a molecular
threshold. That is, when cyclin E, and by extension cyclin E-cdk2
activity, reaches a critical level S phase is triggered in an
all-or-nothing manner. This will only be true if cyclin E function can
continue to accumulate during the extended G1 period. Consistent with this, the mutant cyclin E gene is activated at the
normal developmental time and continues to be expressed during the
extended G1 period due to a failure to activate its own
down regulation (see above).
The molecular mechanism behind such an all-or-nothing threshold is not
clear. Positive feedback amplification has been a popular proposal for
controlling all-or-nothing cell cycle transitions. The biological
activities of mammalian and Drosophila cyclin E and E2F-DP
can formally be placed into a positive feedback loop: overexpression of
cyclin E activates E2F-DP, probably due to cyclin E-cdk2
phosphorylation of pRB, and E2F-DP activates cyclin E
transcription. This type of positive feedback loop makes an attractive
model for control of the restriction point (10, 23, 57), the
time during G1 that mammalian cells commit to S-phase entry
(50). Similar types of interactions between the S. cerevisiae Swi4/6 transcription factor and G1 cyclin
genes (CLNs) can be demonstrated (43, 45).
However, during START, the yeast equivalent of the restriction point,
careful genetic experiments have ruled out positive feedback
amplification between CLN1 or CLN2 and the Swi4/6 transcription factor
(12, 63). Similarly, in Drosophila endocycling cells the genetic evidence indicates that there is no positive feedback
amplification between cyclin E and dE2F-dDP at the G1-S transition, although each can induce the activity of the other when
overexpressed (14, 16). The lack of positive feedback appears to be the consequence of negative feedback mechanisms involving
cyclin E, as described above (16, 41, 53). These overcome
any contribution of cyclin E to positive feedback. Thus, because cyclin
E does not make a positive contribution to its own expression in our
experimental context, cyclin E provides a threshold for entry into S
phase by a mechanism that does not require positive feedback between
cyclin E and dE2F-dDP.
What then is the function of cyclin E? After formation of
prereplicative complexes during G1, and a commitment to
progress into S phase, a poorly described mechanism exists that
triggers the initiation of DNA replication. In vitro experiments
suggest that cdk activity could comprise part of the trigger mechanism. In frog and human extracts, cyclin E-cdk2 and cyclin A-cdk2 are required for DNA synthesis (20, 31, 35, 62). These kinases presumably have substrates that must be phosphorylated in order for
replication origins to fire. Moreover, cyclin E has an essential S-phase function that is independent of E2F-DP-mediated transcription (14, 42). Whatever the critical substrates are, our data
suggest that cyclin E-cdk2 must be part of a mechanism that in vivo has an all-or-nothing response. Cyclin E could activate an amplifying kinase cascade, trigger changes in protein degradation, or affect DNA
unwinding and initiation directly at origins of replication.
This work was initiated while R.J.D. was supported by a fellowship from
the Cancer Research Fund of the Damon Runyon-Walter Winchell Foundation
(DRG-1161). This work was funded in part by a grant from the National
Institutes of Health to P.H.O. (GM37193).
| 1.
|
Andrews, B. J., and I. Herskowitz.
1990.
Regulation of cell cycle-dependent gene expression in yeast.
J. Biol. Chem.
265:14057-14060[Free Full Text].
|
| 2.
|
Ashburner, M.,
P. Thompson,
J. Roote,
P. F. Lasko,
Y. Grau,
M. El Messal,
S. Roth, and P. Simpson.
1990.
The genetics of a small autosomal region of Drosophila melanogaster containing the structural gene for alcohol dehydrogenase. VII. Characterization of the region around the snail and cactus loci.
Genetics
126:679-694[Abstract].
|
| 3.
|
Bandara, L. R.,
V. M. Buck,
M. Zamanian,
L. H. Johnston, and T. N. La.
1993.
Functional synergy between DP-1 and E2F-1 in the cell cycle-regulating transcription factor DRTF1/E2F.
EMBO J.
12:4317-4324[Medline].
|
| 4.
|
Botz, J.,
K. Zerfass-Thome,
D. Spitkovsky,
H. Delius,
B. Vogt,
M. Eilers,
A. Hatzigeorgiou, and P. Jansen-Durr.
1996.
Cell cycle regulation of the murine cyclin E gene depends on an E2F binding site in the promoter.
Mol. Cell. Biol.
16:3401-3409[Abstract].
|
| 5.
|
Breeden, L.
1996.
Start-specific transcription in yeast.
Curr. Top. Microbiol. Immunol.
208:95-127[Medline].
|
| 6.
|
Brook, A.,
J. E. Xie,
W. Du, and N. Dyson.
1996.
Requirements for dE2F function in proliferating cells and in post-mitotic differentiating cells.
EMBO J.
15:3676-3683[Medline].
|
| 7.
|
Brown, N. H., and F. C. Kafatos.
1988.
Functional cDNA libraries from Drosophila embryos.
J. Mol. Biol.
203:425-437[Medline].
|
| 8.
|
Chong, J. P.,
P. Thommes, and J. J. Blow.
1996.
The role of MCM/P1 proteins in the licensing of DNA replication.
Trends Biochem. Sci.
21:102-106[Medline].
|
| 9.
|
Clurman, B. E.,
R. J. Sheaff,
K. Thress,
M. Groudine, and J. M. Roberts.
1996.
Turnover of cyclin E by the ubiquitin-proteasome pathway is regulated by cdk2 binding and cyclin phosphorylation.
Genes Dev.
10:1979-1990[Abstract/Free Full Text].
|
| 10.
|
DeGregori, J.,
T. Kowalik, and J. R. Nevins.
1995.
Cellular targets for activation by the E2F1 transcription factor include DNA synthesis- and G1/S-regulatory genes.
Mol. Cell. Biol.
15:4215-4224[Abstract].
|
| 11.
|
Diffley, J. F.
1996.
Once and only once upon a time: specifying and regulating origins of DNA replication in eukaryotic cells.
Genes Dev.
10:2819-2830[Free Full Text].
|
| 12.
|
Dirick, L.,
T. Bohm, and K. Nasmyth.
1995.
Roles and regulation of Cln-Cdc28 kinases at the start of the cell cycle of Saccharomyces cerevisiae.
EMBO J.
14:4803-4813[Medline].
|
| 13.
|
Du, W.,
M. Vidal,
J. E. Xie, and N. Dyson.
1996.
RBF, a novel RB-related gene that regulates E2F activity and interacts with cyclin E in Drosophila.
Genes Dev.
10:1206-1218[Abstract/Free Full Text].
|
| 14.
|
Duronio, R. J.,
A. Brook,
N. Dyson, and P. H. O'Farrell.
1996.
E2F-induced S phase requires cyclin E.
Genes Dev.
10:2505-2513[Abstract/Free Full Text].
|
| 15.
|
Duronio, R. J., and P. H. O'Farrell.
1994.
Developmental control of a G1-S transcriptional program in Drosophila.
Development
120:1503-1515[Abstract].
|
| 16.
|
Duronio, R. J., and P. H. O'Farrell.
1995.
Developmental control of the G1 to S transition in Drosophila: cyclin E is a limiting downstream target of E2F.
Genes Dev.
9:1456-1468[Abstract/Free Full Text].
|
| 17.
|
Duronio, R. J.,
P. H. O'Farrell,
J. E. Xie,
A. Brook, and N. Dyson.
1995.
The transcription factor E2F is required for S phase during Drosophila embryogenesis.
Genes Dev.
9:1445-1455[Abstract/Free Full Text].
|
| 18.
|
Dynlacht, B. D.,
A. Brook,
M. Dembski,
L. Yenush, and N. Dyson.
1994.
DNA-binding and trans-activation properties of Drosophila E2F and DP proteins.
Proc. Natl. Acad. Sci. USA
91:6359-6363[Abstract/Free Full Text].
|
| 19.
|
Edgar, B. A., and P. H. O'Farrell.
1990.
The three postblastoderm cell cycles of Drosophila embryogenesis are regulated in G2 by string.
Cell
62:469-480[Medline].
|
| 20.
|
Fang, F., and J. W. Newport.
1991.
Evidence that the G1-S and G2-M transitions are controlled by different cdc2 proteins in higher eukaryotes.
Cell
66:731-742[Medline].
|
| 21.
|
Feger, G.,
H. Vaessin,
T. T. Su,
E. Wolff,
L. Y. Jan, and Y. N. Jan.
1995.
dpa, a member of the MCM family, is required for mitotic DNA replication but not endoreplication in Drosophila.
EMBO J.
14:5387-5398[Medline].
|
| 22.
|
Field, S. J.,
F. Y. Tsai,
F. Kuo,
A. M. Zubiaga,
W. G. Kaelin, Jr.,
D. M. Livingston,
S. H. Orkin, and M. E. Greenberg.
1996.
E2F-1 functions in mice to promote apoptosis and suppress proliferation.
Cell
85:549-561[Medline].
|
| 22a.
| Follette, P., and P. H. O'Farrell. Unpublished
data.
|
| 23.
|
Geng, Y.,
E. N. Eaton,
M. Picon,
J. M. Roberts,
A. S. Lundberg,
A. Gifford,
C. Sardet, and R. A. Weinberg.
1996.
Regulation of cyclin E transcription by E2Fs and retinoblastoma protein.
Oncogene
12:1173-1180[Medline].
|
| 24.
|
Girling, R.,
L. R. Bandara,
E. Ormondroyd,
E. W. Lam,
S. Kotecha,
T. Mohun, and N. B. La Thangue.
1994.
Molecular characterization of Xenopus laevis DP proteins.
Mol. Biol. Cell
5:1081-1092[Abstract].
|
| 25.
|
Gloor, G. B.,
C. R. Preston,
D. M. Johnson-Schlitz,
N. A. Nassif,
R. W. Phillis,
W. K. Benz,
H. M. Robertson, and W. R. Engels.
1993.
Type I repressors of P element mobility.
Genetics
135:81-95[Abstract].
|
| 26.
|
Gong, J.,
F. Traganos, and Z. Darzynkiewicz.
1995.
Threshold expression of cyclin E but not D type cyclins characterizes normal and tumour cells entering S phase.
Cell Prolif.
28:337-346[Medline].
|
| 27.
|
Hao, X. F.,
L. Alphey,
L. R. Bandara,
E. W. Lam,
D. Glover, and N. B. La Thangue.
1995.
Functional conservation of the cell cycle-regulating transcription factor DRTF1/E2F and its pathway of control in Drosophila melanogaster.
J. Cell Sci.
108:2945-2954[Abstract].
|
| 28.
|
Harvey, D.,
L. Hong,
M. Evans-Holm,
J. Pendleton,
C. Su,
P. Brokstein,
S. Lewis, and G. M. Rubin.
1997.
.
Clone #LD02934.
Berkeley Drosophila Genome Project, Berkeley, Calif.
|
| 29.
|
Heichman, K. A., and J. M. Roberts.
1994.
Rules to replicate by.
Cell
79:557-562[Medline].
|
| 30.
|
Helin, K.,
C. L. Wu,
A. R. Fattaey,
J. A. Lees,
B. D. Dynlacht,
C. Ngwu, and E. Harlow.
1993.
Heterodimerization of the transcription factors E2F-1 and DP-1 leads to cooperative trans-activation.
Genes Dev.
7:1850-1861[Abstract/Free Full Text].
|
| 31.
|
Jackson, P. K.,
S. Chevalier,
M. Philippe, and M. W. Kirschner.
1995.
Early events in DNA replication require cyclin E and are blocked by p21CIP1.
J. Cell Biol.
130:755-769[Abstract/Free Full Text].
|
| 32.
|
K |
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Abstract
Introduction
