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Molecular and Cellular Biology, April 2005, p. 3027-3039, Vol. 25, No. 8
0270-7306/05/$08.00+0     doi:10.1128/MCB.25.8.3027-3039.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

dDP Is Needed for Normal Cell Proliferation

Maxim V. Frolov,1,2,{dagger} Nam-Sung Moon,1,{dagger} and Nicholas J. Dyson1*

Massachusetts General Hospital Cancer Center, Charlestown, Massachusetts,1 Department of Biochemistry and Molecular Genetics, University of Illinois at Chicago, Chicago, Illinois2

Received 28 September 2004/ Returned for modification 16 November 2004/ Accepted 24 January 2005


   ABSTRACT
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
To gain insight into the essential functions of E2F, we have examined the phenotypes caused by complete inactivation of E2F and DP family members in Drosophila. Our results show that dDP requires dE2F1 and dE2F2 for DNA-binding activity in vitro and in vivo. In tissue culture cells and in mutant animals, the levels of dE2F and dDP proteins are strongly interdependent. In the absence of dDP, the levels of dE2F1 and dE2F2 decline dramatically, and vice versa. Accordingly, the cell cycle and transcriptional phenotypes caused by targeting dDP mimic the effects of targeting both dE2F1 and dE2F2 and are indistinguishable from the effects of inactivating all three proteins. Although trans-heterozygous dDP mutant animals develop to late pupal stages, the analysis of somatic mutant clones shows that dDP mutant cells are at a severe proliferative disadvantage when compared directly with wild-type neighbors. Strikingly, the timing of S-phase entry or exit is not delayed in dDP mutant clones, nor is the accumulation of cyclin A or cyclin B. However, the maximal level of bromodeoxyuridine incorporation is reduced in dDP mutant clones, and RNA interference experiments show that dDP-depleted cells are prone to stall in S phase. In addition, dDP mutant clones contain reduced numbers of mitotic cells, indicating that dDP mutant cells have a defect in G2/M-phase progression. Thus, dDP is not essential for developmental control of the G1-to-S transition, but it is required for normal cell proliferation, for optimal DNA synthesis, and for efficient G2/M progression.


   INTRODUCTION
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
The E2F transcription factor plays an important role in the regulation of cell cycle progression. Changes that activate E2F-dependent transcription, such as the overexpression of E2F genes or the inactivation of pRB family members, promote the progression from G1 to S phase. Conversely, changes that augment the formation of E2F repressor complexes, such as the overexpression of pRB family members or the inhibition of G1 cyclin-dependent kinases, arrest cells in G1.

E2F has been studied primarily in mammalian cells. Mammalian E2F refers to the net activity provided by a large number of proteins. The basic component of E2F is typically a heterodimer of DP and E2F subunits. Mammalian cells contain at least seven E2F genes and two DP genes, and the products of these can be combined in many different permutations (2, 16, 20, 28, 36, 39). Functional overlap between different forms of E2F has made it difficult to identify the precise roles played by individual components. Moreover, the large number of E2F and DP genes has undermined attempts to assess the overall role of E2F in either cell cycle control or animal development.

The Drosophila genome contains two genes with clear homology to the mammalian E2F genes (de2f1 and de2f2) and one gene that is highly homologous to the mammalian DP genes (dDP) (5, 15, 17, 29). dE2F1 and dE2F2 both heterodimerize with dDP and bind to the promoters of E2F target genes. E2F regulation allows responsive genes to be coordinately regulated at the G1-to-S transition. In de2f1 mutant embryos, the expression of E2F target genes, PCNA and RNR2, as revealed by in situ hybridization, is lost (14, 33). Mutation of de2f1 severely reduces cell proliferation and DNA synthesis (14), and de2f1 mutant embryos hatch to produce extremely slow-growing larvae that fail to develop, and die (33). de2f2 mutants have reduced viability and fertility, but de2f2 is not an essential gene (5, 17). However, mutation of de2f2 rescues the strong larval phenotype of de2f1, suggesting that de2f1 and de2f2 act antagonistically during larval development (17).

One of the curious features of de2f2 de2f1 double-mutant animals is that they develop normally until late pupal stages. Imaginal disks taken from these animals display relatively normal patterns of DNA synthesis. This is remarkable, since the two de2fs are the only E2F genes found in a finished sequence of the Drosophila genome and cell proliferation occurs in de2f2 de2f1 double-mutant cells without E2F control. In a similar way, dDP mutant embryos have defects in the spatiotemporal pattern of E2F-dependent transcription, but these animals develop to late pupal stages with no clear larval defects. dDP mutant embryos appear to have fairly normal timing and levels of DNA synthesis at stage 13 (33), but cells in the central midgut initiate S phase later than in wild-type embryos, during stage 14, raising the possibility that dDP may contribute to the correct timing of S phase (13). It is uncertain precisely when maternal supplies of dDP are fully depleted, and this complicates the interpretation of the homozygous mutant phenotypes (11).

To date, studies of dDP or de2f2 de2f1 mutants have been restricted to comparisons between wild-type and mutant animals. The properties of these mutant cells have not yet been compared side by side with wild-type cells in vivo. In addition, although dDP and the dE2Fs are heterodimeric partners, the issue of whether mutating dDP is equivalent to mutating de2f1 and de2f2 has not been carefully examined. Previous studies have stressed that the E2F-regulated patterns of RNR2 and PCNA expression are lost in dDP mutants (13, 33), whereas analysis of de2f1 de2f2 double mutants has shown that the overall levels of expression of these and other E2F-regulated genes are similar to those of wild-type animals (17). To reconcile these discrepancies, and to better understand the roles of E2F and DP proteins in the regulation of cell proliferation, we have compared the consequences of inactivating dDP, dE2F1, and dE2F2. We have generated somatic clones of dDP mutant cells and used these to study the effects of eliminating E2F regulation. Here, we show that the somatic mutation of dDP causes a strong reduction in cell proliferation and that dDP mutant clones display defects both in 5-bromo-2-deoxyuridine (BrdU) incorporation and in the number of mitotic cells. We conclude that dDP is needed for efficient progression through both S phase and G2/M and that the development of trans-heterozygous dDP mutant animals to late pupal stages disguises the fact that dDP mutant cells are at a severe disadvantage when compared directly with wild-type neighbors.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Fly stocks. The following null alleles were used in this work: de2f1rm729 and de2f191 (12); dDPa2, dDPa3, dDPa4, and Df(2R)vg-B, which deletes the dDP gene (27); and de2f276Q1 and de2f2G5.1 (17). Extra lethal mutations on the dDPa3 and dDPa4chromosome were recombined out by homologous recombination.

To make an hs-de2f2 transgene, full-length de2f2 cDNA was cloned under the control of a heat shock-inducible promoter in an hs-Casper vector and transformed into y w1118 flies, together with a helper plasmid, {Delta}2-3. Several independent transgenic lines were established, and induction of dE2F2 protein after heat shock was verified by Western blot analysis.

Mitotic clones were induced using the FLP/FRT technique (40). For determination of clone areas hs-FLP, FRT42D dDPa4/FRT42D Ubi-GFP larvae were heat shocked for 15 min at 37°C 48 h after egg deposition, and the imaginal disks were dissected and fixed. Similar results were observed with the dDPa3 allele. Clone areas were measured with the histogram function of Adobe Photoshop. ey-FLP was used to induce the clones in the eye.

Northern and Western blot analyses and immunofluorescence. RNA isolation, Northern blot analysis, in situ hybridization, staining with anti-phos-H3 antibody (Upstate), and BrdU labeling of eye imaginal disks were performed as previously described (17). For immunostaining of eye imaginal disks, a rabbit polyclonal anti-dE2F1 antibody was used (34). Polytene chromosomes were prepared as described previously (30) and stained using rabbit polyclonal anti-dE2F2 antibody (17) and rabbit polyclonal anti-dDP antibody (8). Since both the anti-dE2F2 and anti-dDP antibodies were raised in rabbits, costaining was performed using an anti-dE2F2 antibody directly conjugated to rhodamine. The sequence of staining was as follows: rabbit anti-dDP, Cy3-conjugated anti-rabbit immunoglobulin G (Vector), and rhodamine-conjugated anti-dE2F2. DNA was visualized with YOYO (Molecular Probes). To make rhodamine-conjugated anti-dE2F2 antibody, an antibody-labeling kit (Amersham) was used.

For Western blot analysis, 20 larvae were homogenized in 200 µl of high-salt ELB extraction buffer (19), frozen in liquid nitrogen, and incubated on ice for 30 min. Samples were spun for 10 min at 4°C, and the supernatant was collected. Proteins were resolved by electrophoresis on a sodium dodecyl sulfate-10% polyacrylamide gel electrophoresis gel. A similar procedure was used for SL2 cells. The following antibodies were used for a subsequent Western blot analysis: mouse monoclonal anti-dDP, Yun-3 (11), rabbit polyclonal anti-dE2F2 (17), and guinea pig polyclonal anti-dE2F1 (3). To quantify the mitotic defect in dDP mutant clones, we outlined the clone using a lasso tool in Photoshop CS, recorded the number of phos-H3-positive cells, and then counted the number of stained cells in an identically sized area of neighboring wild-type tissue.

Manipulations with SL2 cells. RNA interference (RNAi) in SL2 cells was carried out using double-stranded RNA (dsRNA) as described previously (35). For fluorescence-activated cell sorting (FACS), [3H]thymidine incorporation, and cell count analyses, cells were incubated with dsRNA for 4 days and then replated at a density of 0.5 million cells per ml and analyzed 3 days later. No differences were found when cells were re-treated with dsRNA at the time of replating.

To incorporate BrdU into SL2 cells prior to FACS analysis, 15 µl of BrdU-5-fluoro-2'-deoxyuridine (FdU) cell proliferation labeling reagent (Amersham) was added to the wells. After being harvested, the cells were washed once with cold phosphate-buffered saline and fixed in 95% ethanol. To detect incorporated BrdU, a standard protocol was used (35). Flow cytometry analysis was performed on a Becton-Dickinson FACScan, collecting data from 20,000 cells per sample.

For [3H]thymidine labeling, on the last day of treatment, cells were incubated with 5 to 15 µCi of [3H]thymidine for 1 h at 25°C, washed with phosphate-buffered saline, harvested, and frozen. The cells were lysed with 0.3 N NaOH, and DNA was precipitated by the addition of an equal volume of 20% trichloroacetic acid. Lysates were spotted on GF-C filters (Whatman) and washed with ice-cold 10% trichloroacetic acid and ethanol. Incorporated radioactivity was measured in a scintillation counter.

EMSA. Electrophoretic mobility shift assays (EMSAs) were performed with 5 µg of total extract from SL2 cells. The samples were incubated at 4°C for 30 min in a final volume of 20 µl of EMSA reaction buffer (40 mM KCl, 20 mM HEPES, pH 7.6, 2.5 mM MgCl2, 0.1 mM EGTA, 4% Ficoll, 2 mM spermine, 0.5 mM dithiothreitol, 10 ng of sonicated salmon sperm DNA/µl, 25 ng of random oligonucleotide/µl, and 1 µg of bovine serum albumin/µl). End-labeled double-stranded oligonucleotides containing E2F sites (~10 pg) were added, and the samples were further incubated for 30 min at 4°C. The samples were loaded on a 4% polyacrylamide gel and separated by electrophoresis for 4 h at 120 V in 0.25x Tris-borate-EDTA. The gels were dried and visualized by autoradiography.


   RESULTS
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Targeting dDP as a means to inactivate dE2F1 and dE2F2. The analysis of de2f2 de2f1 double-mutant animals has revealed that fairly normal patterns of cell proliferation can occur without E2F regulation; however, the mutant animals are retarded in their larval development by ~1 day (17), and this developmental delay might mask, or be caused by, altered rates of cell proliferation. Most commonly, proliferation rates are compared in Drosophila using clonal analysis because it allows a side-by-side comparison of wild-type and mutant cells within a single tissue. The generation of de2f2 de2f1 mutant clones was complicated by two technical hurdles. First, the de2f2 and de2f1 genes reside on different chromosomes. Therefore, the generation of the cell mutant for both E2F genes requires two independent recombination events to occur at the same time. Second, in order to use an FLP/FRT technique (40), the mutation needs to be recombined onto an FRT-containing chromosome. Unfortunately, de2f2 is located at 39B on the second chromosome, close to the centromere, and cannot be easily recombined. This essentially precluded any direct comparison of wild-type and de2f2 de2f1 mutant clones.

To circumvent these problems, we decided to generate clones of cells carrying a mutation in dDP. dDP is the heterodimeric partner of both dE2F1 and dE2F2, and it is generally assumed that the loss of DP proteins should impair an E2F function as effectively and specifically as removing E2F proteins. However, this assumption has not been rigorously examined, and the discovery of a new class of E2F proteins in plant and animal cells that can act independently of a DP partner (6, 9, 24) highlights the need to verify this issue for each experimental system. Hence, prior to analysis of the phenotype of dDP mutant clones, we asked whether removing dDP could be safely said to mimic the loss of de2f1 and de2f2.

E2F-binding activity was assessed by EMSA in extracts prepared from Drosophila SL2 tissue culture cells. Four specific complexes were detected using a probe containing an E2F-binding site (Fig. 1A). The complexes bind to the probe in a sequence-specific manner because their DNA-binding activity can be efficiently competed by a wild-type E2F-binding site, but not when the site has been mutated. E2F-specific DNA-binding activity was further characterized in cells depleted of dE2F1, dE2F2, or dDP by RNAi. The efficiency of depletion was monitored by Western blot analysis (data not shown). Removal of dE2F1 eliminates complexes a and b, whereas depletion of dE2F2 results in the disappearance of the faster-migrating complexes c and d (Fig. 1B). Cells treated with dDP dsRNA show a complete loss of E2F-specific DNA-binding activity and lack all four complexes. Thus, the four E2F-binding complexes detected by using this method contained dDP and either dE2F1 or dE2F2.



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  		FIG. 1. E2F-specific DNA-binding activity is dependent upon dDP in vitro. (A) Total extracts of SL2 cells were prepared and analyzed in EMSA with oligonucleotides containing an E2F-binding site (underlined) (ATTTAAGTTTCGCGCCCTTTCTCAAATTT) (left lane). The specificity of the retarded complex was verified by preincubating the extracts with a 100-fold molar excess of competing unlabeled oligonucleotides containing either a wild-type (WT) or a mutant (MT) E2F-binding site. End-labeled oligonucleotide alone was also migrated in the gel (right lane). The nonspecific band is designated by an asterisk. (B) Cells were treated with nonspecific dsRNA used as a control (NS) or with dE2F1 (E1), dE2F2 (E2), and dDP (dDP) dsRNAs. Note the lack of E2F-specific DNA-binding activity in SL2 cells following depletion of dDP.

 
Although EMSAs are often used to provide a profile of E2F complexes, the method has limitations. EMSAs are in vitro-based assays that employ nonphysiological conditions. The patterns of complexes detected vary in different buffers, and not all E2F-responsive elements bind complexes under standard reaction conditions. We therefore examined the binding of dE2F and dDP proteins in vivo by staining polytene chromosomes. The advantage of this technique is that the immunostaining patterns reveal the genomewide distribution of native chromatin-associated proteins at their physiological targets. Drosophila third-instar larval salivary gland polytene chromosomes were stained with the antibodies raised against dE2F2 and dDP. The chromosomes were counterstained with YOYO to visualize the DNA. YOYO staining is strongest in the condensed regions of euchromatin and in the heterochromatin and is weakest in interbands.

In wild-type animals, anti-dE2F2 antibody recognized ~100 euchromatic sites (Fig. 2A to C). No dE2F2 was detected in chromocenter or telomeric regions. dE2F2 associated mainly with interband regions and appeared to be excluded from brightly stained bands. These sites represent specific locations of dE2F2, because the staining was completely absent in the preparations from the de2f2 mutant larvae (data not shown). When polytene chromosomes were stained with antibody against dDP protein, a similar distribution was observed (Fig. 2D to F). dDP, like dE2F2, was present mostly in interbands and was absent from the chromocenter and from bands. To test whether dE2F2 colocalizes with dDP, chromosomes were costained with anti-dDP and anti-dE2F2 antibodies. Since both antibodies were raised in rabbits, the anti-dE2F2 antibody was directly conjugated to rhodamine, and Cy5-conjugated anti-rabbit secondary antibody was used to detect dDP. Appropriate controls were performed to assure that the dE2F2 signal did not come from nonspecific binding of anti-dE2F2 antibody to Cy5-conjugated anti-rabbit secondary antibody. As seen in Fig. 2G to I, dDP showed almost complete overlap. We noted that there were a couple of sites that immunostained with anti-dDP antibodies but not with antibodies to dE2F2 (see, for example, Fig. 2I).



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  		FIG.2. dE2F2 colocalizes with dDP on polytene chromosomes. (A to F) Wild-type polytene chromosomes were stained with anti-dE2F2 and anti-dDP antibodies and counterstained with YOYO to visualize DNA. (G to I) Higher magnification of a fragment of wild-type polytene chromosome that was costained with anti-dE2F2 and anti-dDP antibodies. The arrows indicate one of the sites that stains for dDP but not for dE2F2. (J to L) dDP mutant chromosomes stained with anti-dE2F2 antibody. (M to O) de2f2 mutant chromosomes stained with anti-dDP antibody. Note the reduction in the number of dDP binding sites in the de2f2 mutant compared to the wild type. (P to R) de2f2 de2f1 double-mutant chromosomes stained with anti-dDP antibody.

 
The dE2F2 staining pattern completely disappeared when polytene chromosomes from dDP mutant animals were examined (Fig. 2J to L), indicating that dE2F2 binds to chromosomes in a dDP-dependent manner. However, the dDP signal was not completely eliminated in de2f2 mutants (Fig. 2M to O). In these mutants, a small number of dDP bands persist that presumably represent the similarly small number of loci that were bound by dDP but not by dE2F2 in wild-type animals. Since dDP heterodimerizes with both dE2F1 and dE2F2, the most likely explanation is that these represent sites of specific binding by dE2F1-dDP heterodimers. We were unable to confirm this by immunochemistry, because none of the anti-dE2F1 antibodies that we have raised or obtained from others stain polytene chromosomes specifically. However, we addressed this question genetically by comparing the distribution of dDP on polytene chromosomes of de2f2 single-mutant and de2f2 de2f1 double-mutant animals. The residual dDP staining pattern evident in de2f2 mutant animals (Fig. 2M to O) is lost in de2f2 de2f1 double-mutant animals (Fig. 2P to R). Thus, we conclude that that dE2F2 binds to its natural chromosomal targets in a dDP-dependent manner and, conversely, that dDP requires both dE2F1 and dE2F2 to associate with chromatin.

To test whether the loss of E2F and DP proteins has a similar effect on E2F activity, we quantitatively compared the phenotype of dDP-deficient SL2 cells with the phenotype of cells depleted of dE2F1 and dE2F2 by RNAi. S-phase entry and E2F-dependent transcription, two functional readouts of E2F activity, were monitored in cells treated with dsRNA and depleted of dDP, dE2F1/dDP, dE2F1/dE2F2, or dE2F1/dE2F2/dDP. The expression of E2F target genes was assessed by Northern blot analysis. To measure changes in cell cycle parameters, cells were labeled with BrdU and analyzed by FACS. These experiments were performed in triplicate. In accordance with our previous data, BrdU incorporation was decreased in the dE2F1-deficient cells by ~1 order of magnitude, and the expression of E2F targets was also strongly reduced (Fig. 3A and B). Codepletion of dE2F2 partially elevated the expression of E2F targets in dE2F1-deficient cells and restored BrdU incorporation to 45% of that in control treated cells. In a similar way, codepletion of dDP removed the S-phase block of dE2F1-deficient cells. Importantly, the level of BrdU incorporation and the level of expression of E2F targets were indistinguishable in cell cultures depleted of dDP, of both dE2Fs, or of all three proteins, dE2F1/dE2F2/dDP (Fig. 3A and B). As a further test, we performed Northern blot analysis to compare the expression levels of E2F targets in vivo in eye imaginal disks dissected from dDP and de2f2 de2f1 mutant third-instar larvae. The levels of DNA pol{alpha}, cyclin E, MCM3, RNR2, and PCNA transcripts isolated from the dDP mutant eye disks were identical to those seen in the de2f2 de2f1 double mutant (Fig. 3C) (8). We noted that cyclin E expression is more severely affected by the loss of E2F/DP proteins in SL2 cells than in eye disks. This difference presumably reflects the fact that in vivo, cyclin E expression is not simply driven by cell cycle progression but also responds to developmentally regulated signaling pathways (12), and these signals are not present in the tissue culture cells. Nevertheless, these results show that targeting dDP has effects on E2F-dependent transcription that are similar to those caused by targeting dE2F1/dE2F2. No further changes were evident when all three proteins were targeted in SL2 cells.



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  		FIG. 3. Proliferative properties of SL2 cells depleted of dE2F1, dE2F2, or dDP. (A) Cells depleted of dE2F1, dE2F2, and dDP or their combinations were labeled with BrdU, and the percentage of BrdU-positive cells were assayed by FACS. The number of BrdU-positive cells is dramatically reduced in the absence of dE2F1. Removal of dE2F2 does not significantly affect the number of BrdU-positive cells but restores BrdU incorporation lost in cells depleted of dE2F1. Note that dE2F1/dE2F2-, dDP-, dE2F1/dDP-, and dE2F1/dE2F2/dDP-depleted cells show similar values of BrdU incorporation. The error bars indicate standard deviations. NS, nonspecific dsRNA. (B) Total RNA was isolated from dsRNA-treated cells, and the levels of PCNA, cyclin E, RNR2, and MCM3 transcripts were followed by Northern blot analysis. (C) Northern analysis of total RNAs isolated from eye disks of different genotypes shows that E2F-regulated genes are expressed at similar levels in dDP mutant and de2f1 de2f2 double-mutant animals.

 
To further characterize cells depleted of dE2Fs and dDP family members, we examined S-phase progression. Following treatment with dsRNAs, one-half of the cell cultures were labeled with [3H]thymidine to determine the level of DNA synthesis while the other half were labeled with BrdU and processed for two-dimensional FACS analysis. As expected, dE2F1-depleted cells showed a dramatic reduction of [3H]thymidine incorporation compared to control treated cells (Fig. 4A). Although the block was relieved by the codepletion of dE2F2, this was insufficient to restore a normal rate of DNA synthesis (Fig. 4A). Incorporation of [3H]thymidine was severely reduced in dE2F1/dE2F2-deficient cells, as well as in cells depleted of dDP (Fig. 4A).



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  		FIG. 4. S-phase defects of SL2 cells depleted of dE2F1, dE2F2, or dDP. (A and B) Cells depleted of dE2F1, dE2F2, and dDP were labeled with [3H]thymidine to measure DNA synthesis or with BrdU for 16 h and subjected to two-dimensional FACS analysis. The error bars indicate standard deviations. (C) Proliferation indices of cells following RNAi treatment. Loss of dE2F1 results in a significant reduction of S-phase cells and an increased G1 population. Removal of dE2F2 restores S-phase entry defects of dE2F1-depleted cells but does not rescue S-phase progression defects. Despite containing BrdU-positive cells, populations of dDP, dE2F1/dE2F2, dE2F1/dDP, and dE2F1/dE2F2/dDP RNAi-treated cells have low rates of DNA synthesis and proliferate very slowly. In populations of dDP- or dE2F1/dE2F2-depleted cells, a substantial number of BrdU-negative cells have intermediate DNA contents, indicating that these cells are stalled within S phase.

 
As further evidence of slowed S-phase progression, the FACS profile of dDP- or dE2F1/dE2F2-depleted cells shows an increase in the population of BrdU-negative cells with a DNA content that is between the major G1 and G2 peaks, suggesting that these cells have stalled within S phase (Fig. 4B). Using a long pulse of BrdU incorporation (16 h), the majority of BrdU-positive cells in the control population accumulate with a G2 DNA content, indicating that most cells that enter S phase during this period have sufficient time to complete S phase. In contrast, in dDP-depleted or dE2F1/dE2F2-depleted populations, the DNA content of BrdU-positive cells was spread between the G1 and G2 peaks, indicating that these cells replicate DNA more slowly than wild-type cells. To measure whether proliferation potential is impaired in dDP- and dE2F1/dE2F2-depleted cells, we counted the cells in the culture following protein depletion (at which time most of the protein product had been lost) and again 3 days later (Fig. 4C). As expected, depletion of dE2F1 blocked cell proliferation, and we found no increase in cell numbers within this 3-day period. While depletion of dE2F2 relieved the block to S-phase entry caused by removing dE2F1, this resulted in only a marginal increase in cell numbers that was indistinguishable from that of dDP-depleted cells. Thus, the direct targeting of depletion of dDP has an impact on DNA synthesis and cell proliferation that is similar to targeting dE2F1 and dE2F2.

E2F expression is altered in dDP mutant clones. Having established that removing dDP impairs E2F function as efficiently as targeting dE2F1 and dE2F2, we set out to use dDP mutant alleles to generate clones of cells lacking E2F activity in vivo. Since somatic recombination generates clones that are homozygous for the mutant chromosome, it is important to be confident that the mutant chromosomes do not carry extra lethal mutations. We examined three dDP mutant alleles, dDPa2, dDPa3, and dDPa4 (33). dDPa2 is a point mutation converting Trp241 within a dimerization domain into a stop codon (33). Genomic fragments containing the dDP gene were sequenced from the two other mutant chromosomes. dDPa3 was found to contain a G-for-A substitution at the 5' end of the first intron that is predicted to prevent correct splicing of the dDP pre-mRNA. A failure to splice the first intron of dDP results in an open reading frame that terminates prematurely at a naturally occurring stop codon in the first intron. Sequencing of the dDPa4 allele revealed that it contains a point mutation converting Gln80 into a stop codon. The short truncated proteins produced from dDPa3 and dDPa4 alleles lack DNA-binding, dimerization, and marked-box domains and therefore are expected to be functional nulls. Western blot analysis confirmed that no wild-type dDP protein was produced from each of three alleles (data not shown).

All three mutant chromosomes cause pupal lethality when placed over Df(2R)vg-B, a deficiency uncovering dDP. However, each of the dDP mutant chromosomes was found to carry multiple additional lethal mutations, and we attempted to remove these by recombination. For our future analysis, we picked recombinants that were pupal lethal as homozygotes and that could be rescued to viable adults by a dDP genomic construct (27). Successful recombinants were recovered for dDPa3 and dDPa4 alleles, but not for dDPa2, indicating that an extra lethal mutation is likely to be closely linked to the dDP locus. Each of the phenotypes described below was observed in clones generated with both dDPa3 and dDPa4, and these effects were rescued by a dDP genomic construct.

We first assayed the expression of dE2F1 and dE2F2 in dDP mutant clones by immunofluorescence. Several reports have described dE2F1 expression patterns, but none of these studies has verified the specificity of the staining patterns with de2f1 mutant alleles (1, 4, 21, 31). We examined the expression pattern of dE2F1 using a rabbit polyclonal antibody that gives a robust staining pattern on eye imaginal disks (34). Eye disks dissected from de2f2 de2f1 double-mutant animals were used as controls, since they are similar in size and development to the wild-type disks, unlike the disks of the de2f1 single mutant (33).

In wild-type eye imaginal disks, dE2F1 was expressed at high levels in cells both anterior and posterior to the morphogenetic furrow and at slightly reduced levels in cells synchronized in G1 in the morphogenetic furrow. dE2F1 expression posterior to the morphogenetic furrow coincides with cells synchronously entering S phase in the second mitotic wave (Fig. 5A), and as previously reported, the disappearance of this signal seems to be tightly coupled to S-phase entry (21, 31). This staining pattern is specific, because it is absent from disks trans-heterozygous mutant for de2f1 (Fig. 5B). One of the interesting features of this staining pattern is that it is broader than the pattern of E2F-stimulated transcription. PCNA is a well-known E2F transcriptional target and is induced by a pulse of dE2F1-dependent transcription preceding the G1/S transition. PCNA is highly expressed in cells immediately posterior to the morphogenetic furrow (Fig. 5C) (10, 17). This stripe of PCNA expression corresponds to the area with a high level of dE2F1 and partially overlaps the second mitotic wave. However, it is striking that no PCNA expression was evident in the region just anterior to the morphogenetic furrow, where dE2F1 is present at similarly high levels (Fig. 5B and C). A transgene expressing a PCNA-green fluorescent protein (GFP) fusion protein (37) was used to simultaneously monitor PCNA and dE2F1 expression in the same disk. As seen in Fig. 5D, PCNA-GFP expression only partially overlaps with the expression of dE2F1 protein in the eye imaginal disk, and no PCNA-GFP is found anterior to and within the morphogenetic furrow. These results are consistent with previous studies that reported that cells entering the second mitotic wave have a transiently high level of E2F1. However, these results show the discordance between the levels of dE2F1 detected by immunostaining and the expression of E2F-regulated targets. The transcriptional activity of dE2F1 is known to be inhibited by RBF1, and cells arrested in G1 with low cdk activity presumably contain dE2F1 complexes that are inactivated by association with RBF1. These results illustrate that, since E2F-dependent transcription can be high or low in cells that have high levels of dE2F1 protein, dE2F1 immunostaining is an inaccurate marker of E2F activity.



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  		FIG. 5. dE2F1 is not sufficient to induce E2F-dependent transcription. (A and B) Eye disks were stained with anti-dE2F1 antibody. (A) In a wild-type disk, dE2F1 is expressed in the morphogenetic furrow and in the cells anterior and posterior to the furrow. (B) No staining with anti-dE2F1 antibody is observed in the eye disks trans-heterozygous for de2f1 mutant alleles. (C) In situ hybridization of eye disks from wild-type larvae with PCNA probe. PCNA transcripts are detected in cells posterior to the furrow. (D) Direct comparison of dE2F1 (red) and PCNA (green) expression patterns using a transgene producing a GFP-PCNA fusion protein. Arrow, position of the morphogenetic furrow.

 
When the levels of dE2F1 and dE2F2 were examined in dDP mutant clones, we found that the expression of both proteins was altered by the absence of dDP. dDP mutant clones were induced in eye imaginal disks using an FLP transgene under the control of an eye-specific promoter, ey-FLP, and identified by the absence of GFP signal. dE2F2 is expressed ubiquitously in the developing eye and is present both in proliferating and in differentiating cells (17). However, dE2F2 was almost undetectable in dDP mutant clones (Fig. 6A and B). The loss of the dE2F2 protein in dDP mutant clones could be rescued by a dDP genomic rescue construct (Fig. 6C and D). A less dramatic change was seen in dE2F1 expression: the level of dE2F1 in dDP mutant clones was reduced, and staining was more diffuse than in control cells (Fig. 6E to G). Similar changes in levels were seen in SL2 cells treated by RNAi: the levels of dE2F2 and dE2F1 were reduced in cells depleted of dDP (Fig. 7A), and reciprocally, cells soaked with dE2F1 and dE2F2 dsRNAs had a reduced level of dDP protein (Fig. 7A). In a similar way, the level of dE2F2 protein was strongly reduced in dDP mutant larvae compared with wild-type controls, and the level of dDP was similarly reduced in de2f2 mutant larvae (Fig. 7B). Moreover, the reexpression of dE2F2 from a heat shock-inducible transgene in de2f2 mutant larvae restored dDP to normal levels within 2 h (Fig. 7C). No decrease in the levels of dDP mRNA was observed in either de2f2 or de2f1 de2f2 mutant larvae, and conversely, no decrease in the levels of de2f1 or de2f2 mRNAs was observed in dDP mutant animals (Fig. 7D). Thus, the changes in protein levels are most likely due to a change in protein synthesis or stability rather than a change in transcription. We conclude that dE2F1 and dE2F2 are required to maintain normal levels of dDP, and vice versa. As a consequence, mutation of dDP reduces the levels of dDP, dE2F1, and dE2F2. Similarly, the mutation of de2f2 and de2f1 also affects the levels of dDP. Although protein levels do not always reflect activity, these changes provide a very simple reason why the mutation of dDP has biological effects that are similar to mutation of de2f1 and de2f2.



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  		FIG. 6. Levels of dE2F1 and dE2F2 are reduced in dDP mutant clones. Clones were induced with ey-FLP, and eye disks were stained with anti-dE2F2 (A to D) or with anti-dE2F1 (E to G) antibody. Mutant clones were identified by the absence of GFP marker (green). (A and B) dE2F2 protein is at a low level in dDP mutant clones. (C and D) The loss of dE2F2 staining is rescued by a dDP transgene. (E to G) dE2F1 staining is reduced and diffuse in a dDP mutant clone. Arrow, position of the morphogenetic furrow.

 


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  		FIG. 7. dDP and dE2F2 levels are interdependent in mutant animals and in cells. (A) Cells were treated with nonspecific (NS), dE2F1 (E1), dDP, and dE2F2 (E2) dsRNAs, and cell extracts were analyzed on Western blots using antibodies specific for dE2F1, dE2F2, and dDP. Note that depletion of dDP affects the levels of both dE2F1 and dE2F2, and conversely, dDP is reduced following depletion of dE2F1 and dE2F2. Anti-tubulin mouse monoclonal antibody was used as a loading control. (B) Western blot analysis of larval extracts from Canton S (wild-type), de2f276Q1/de2f2G5.1 (de2f2–), de2f276Q1/de2f2G5.1, de2f191/de2f1rm729 (de2f2–; de2f1–), and dDPa2/Df(2R)vg-B (dDP–) animals. (C) A wild-type level of dDP can be restored in de2f2 mutants by the reexpression of dE2F2. A heat shock-inducible de2f2 transgene, hs-de2f2, was introduced into de2f2 mutant animals. The level of dDP in larvae was monitored by Western blot analysis before heat shock (no HS) and at 30 min (30' AHS) and 2 h (2 h AHS) following a 1-h heat shock (de2f2– hs-de2f2). The rescue was not due to the heat shock treatment, because the level of dDP was unchanged in de2f2 mutants that lack an hs-de2f2 transgene (de2f2–). (D) Northern blot analysis of larvae of the same genotypes used in panel B. 32P-labeled RNA probes for de2f1, de2f2, and dDP reveal that there is no decrease in the steady-state level of dDP mRNA in the de2f1 de2f2 double mutant or of de2f1 and de2f2 mRNAs in the dDP mutant.

 
Abnormal S-phase progression and G2/M defects in dDP mutant cells. To determine how the loss of dDP affects cell proliferation, homozygous mutant dDP clones were generated following the induction of FLP recombinase from a heat shock-inducible promoter in the first larval stage and were analyzed 72 h later. In eye imaginal disks, dDP mutant clones were significantly smaller than wild-type twin spots marked with GFP (Fig. 8A). The small size of dDP mutant clones was rescued by a dDP transgene, confirming that this defect is attributable to the loss of dDP (Fig. 6A and C and data not shown). The average area covered by dDP mutant cells in wing imaginal disks at the time when most of the cells are asynchronously proliferating was approximately one-ninth of the area covered by twin spots (Fig. 8B and C). The mutant clones do not show an increase in apoptosis, as judged by staining with acridine orange, a dye that stains apoptotic cells, or by the absence of the cleaved form of caspase 3 (data not shown) or terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling staining. Thus, cells lacking E2F function are at a severe proliferative disadvantage when directly compared with wild-type cells.



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  		FIG. 8. dDP mutant cells have a severe proliferative disadvantage. Clones of dDP mutant cells were induced at the first-instar larval stage by mitotic recombination. Examples of the clones observed in the eye (A) and wing (B) imaginal disks are presented. Homozygous mutant clones were visualized by the absence of GFP, while homozygous wild-type twin spots (+/+ in panel B) show a doubled GFP signal on a heterozygous background. (C) Clones were induced as described for panel B, wing imaginal disks were dissected and fixed at the early third-instar larval stage, and the sizes of the mutant clones and the corresponding twin spots were determined. The bars are ordered according to the size of the wild-type twin spots. Values for average clone areas of dDP mutant cells are indicated.

 
The small size of dDP mutant clones made it technically difficult to determine a cell cycle profile by flow cytometry. As an alternative approach, we stained eye imaginal disks carrying dDP mutant clones for various cell cycle markers. Clones were induced using an ey-FLP transgene, which is expressed very early in development, and as a result, almost no heterozygous tissue is present in the third larval instar. Eye imaginal disks were labeled with BrdU to follow the cells in S phase, and the mutant clones were distinguished by the absence of GFP. In wild-type disks, cell proliferation is localized to two distinct areas, the first and second mitotic waves, which are separated by the morphogenetic furrow. In the morphogenetic furrow, cells pause in G1 and enter S phase in a highly synchronized manner, generating a sharp band of BrdU-positive cells (the second mitotic wave). A previous study of homozygous dDP mutant embryos suggested that cells display a delay in S-phase entry as the maternally supplied dDP products are depleted (13). Since the mutant clones generated by somatic recombination are adjacent to normal cells, this method allowed us to precisely compare the timing of S-phase entry in cells lacking E2F function in our analysis. Strikingly, dDP mutant cells enter S phase at exactly the same time as wild-type cells (Fig. 9A and B), and since the stripe of BrdU labeling was not extended within the mutant clones, there was no recognizable delay in the completion of S phase. Hence, at least in the eye disk, wild-type and mutant cells enter and exit S-phase at the appropriate times. We noted that the maximal intensity of BrdU incorporation that was detected in the second mitotic wave was substantially decreased in dDP mutant clones (Fig. 9A and B). This phenotype is attributed to the loss of dDP, since reintroduction of a dDP transgene restored BrdU incorporation to normal levels (Fig. 9C and D). The intensity of BrdU labeling serves as a rough indication of the amount of nucleotide incorporated during the labeling period, and this reduction suggests that the rate of DNA replication is lower in cells lacking E2F function. This is consistent with the low level of DNA synthesis observed in dDP- and dE2F1/dE2F2-depleted tissue culture cells (Fig. 4A).



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  		FIG. 9. Cell cycle characteristics of dDP mutant cells. Clones of dDP mutant cells were induced with ey-FLP, and eye imaginal disks were labeled with BrdU to visualize cells in S phase (A to D, J, and K) or stained with anti-cyclin A antibody (E and F), anti-cyclin B antibody (G and H), and anti-phos-H3 antibody (I to L). Mutant cells lack GFP signal. (A and B) Cells in dDP mutant clones incorporate BrdU at the same time as wild-type cells but with less efficiency. (C and D) Reduced efficiency of BrdU labeling in dDP mutant clones is fully rescued by a dDP genomic rescue construct. The timing of induction of cyclin A (E and F) and cyclin B (G and H) is normal in a dDP mutant clone. The number of cells in mitosis is reduced in dDP mutant cells, as visualized by anti-phos-H3 staining (I). Ectopic expression of cyclin E restores the efficiency of BrdU incorporation (J and K) but not the reduced number of cells entering mitosis (L) in dDP mutant clones. Arrow, position of the morphogenetic furrow.

 
Next, we examined progression through G2 into M phase in dDP mutant clones. Cyclin A is expressed at the end of S phase and degraded during mitosis and therefore marks cells in G2. Cyclin A is properly induced in dDP mutant clones, suggesting that the timing of G2 is normal (Fig. 9E and F). Cyclin B is expressed following cyclin A, and no delay in cyclin B expression was evident in dDP mutant clones (Fig. 9G and H).

Despite the normal appearance of cyclins, dDP mutant cells exhibit defects in G2/M progression. Eye disks were stained with anti-phos-H3 to reveal mitotic cells. In examining large numbers of dDP mutant clones, we noticed that the number of mitotic cells was consistently reduced in dDP mutant clones (Fig. 9I). On average, dDP mutant clones contained only one-third the number of phos-H3-stained cells found in the adjacent wild-type tissue. Such a substantial decrease in the number of mitotic cells could easily explain the small size of the dDP mutant clones.

There are at least two different potential explanations for this defect. One possibility is that dDP is needed for the expression of genes that are required during mitosis. Indeed, previous microarray studies have shown that the depletion of dDP reduces transcription from several genes encoding proteins that act at mitosis, such as string, cdc2, and Bub1 (8). An alternative explanation suggested by the observation that dDP mutant cells enter and exit S phase at the normal time but have a lower maximal level of BrdU incorporation is that dDP mutant cells may be prone to incomplete DNA replication, a phenomenon that occurs in Drosophila during endoreduplication cycles. However, we have failed to find evidence of incomplete DNA replication in the dDP mutant clones. The dDP mutant clones showed no elevated staining in terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling assays, which label free DNA ends, or using an antibody that recognizes the phosphorylated form of histone H2Av, a marker of double-strand breaks (25). Thus, these cells do not appear to contain an excessive level of DNA ends.

While examining this issue, we used an hs-cyclin E transgene to synchronously drive cells into S phase and monitored the progression of these cells through the cell cycle. Wild-type and dDP mutant cells were driven into S phase in the anterior part of the morphogenetic furrow (Fig. 9J and K). BrdU labeling 2 h after the heat shock pulse showed that cyclin E drives wild-type and dDP mutant cells into S phase with equal efficiency. Strikingly, the transient expression of cyclin E was sufficient to restore the normal level of BrdU incorporation in clones of dDP mutant cells. The finding that the S-phase defects of dDP mutant cells can be rescued by elevated levels of cyclin E is consistent with an earlier observation that the inactivation of DACAPO, a negative regulator of cyclin E kinase activity, rescues DNA synthesis defects of dE2F1/dE2F2-deficient SL2 cells (18). Eye disks were stained with anti-phos-H3 antibody at 0, 1.5, 3, 6, and 9 h after induction of cyclin E. The number of ectopic phos-H3-positive cells was highest 6 h after cyclin E induction (Fig. 9L); however, the number of phos-H3-positive cells in the dDP mutant clones remained extremely low—approximately one-third of the number in the adjacent wild-type tissue. Thus, the transient expression of cyclin E is sufficient to rescue the BrdU incorporation defects of dDP mutant cells, but this is not sufficient to correct the defects in M-phase entry. This suggests that the reduced number of mitotic cells in the dDP mutant clones is unlikely to be simply a consequence of inefficient DNA synthesis.

Taken together, these experiments show that the mutation of dDP, and the loss of E2F regulation that occurs as a result, impairs cell proliferation significantly. These changes do not seem to affect the timing of S-phase entry in vivo, but they reduce the efficiency of S-phase progression and cause a substantial drop in the number of cells that progress through M phase.


   DISCUSSION
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 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
In this study, we have exploited the relative simplicity of Drosophila E2F family members to examine the effects of removing E2F or DP protein. The surprising finding that has emerged from the study of Drosophila de2f1 de2f2 double mutants is that these animals develop until late pupal stages with relatively normal patterns of cell proliferation (17). This begs the question, which cellular functions require E2F regulation? Or, put another way, how "normal" are E2F-deficient cells?

We draw two main conclusions from the experiments described here. The first conclusion is that in Drosophila the effects of inactivating dDP appear indistinguishable from the effects of inactivating both dE2F1 and dE2F2. This finding is of practical value because it validates the use of dDP mutant alleles to eliminate E2F activity. It is also an important conceptual point. E2F is generally considered to be a heterodimeric factor, but the idea that E2F and DP proteins function only in partnership with one another has not been rigorously examined in any experimental system. Indeed, there are several indications that this assumption is unlikely to be true in all species. Mammalian cells contain many different E2F and DP proteins, and several of them have been reported to interact with additional proteins (7, 22, 26, 38). More significantly, the recent characterization of E2F7, an E2F family member with a duplicated DNA-binding domain that appears to bind to E2F-regulated promoters without a DP partner, has added a new level of complexity. Similar genes are found in plants, suggesting that this DP-independent mode of action is conserved. Although the functions of most of these novel E2Fs are not yet known, they can, at least in the case of E2F7, bind to classic cell cycle E2F targets. Thus, in these species, it is expected that eliminating all E2F proteins will most likely have consequences different from those of removing all DP proteins.

The Drosophila genome lacks any clear ortholog of E2F7, and the results described here strongly suggest that both of the known Drosophila E2F proteins require dDP to function, and vice versa. This conclusion is based on both biochemical and genetic data. In EMSAs, all of the complexes that bound to an E2F probe in a sequence-specific manner contained both dE2F and dDP proteins and were eliminated when dDP or dE2F1 and dE2F2 were removed by RNAi. Immunostaining experiments on polytene chromosomes show that E2F and DP proteins colocalize in vivo and that the presence of dDP at its natural targets requires dE2F1 and dE2F2; conversely, dE2F2 requires dDP. In addition, the removal of dDP (either by RNAi or in trans-heterozygous mutant animals) has effects on the expression of E2F target genes and on S-phase entry that are indistinguishable from the changes seen when dE2F1 and dE2F2 are removed. The direct targeting of dDP, dE2F1, and dE2F2 gave no further changes over cells in which either dDP or dE2F1 and dE2F2 were targeted.

These results strongly suggest that dDP and the two dE2F proteins are exclusive and interdependent partners. However, we note that there is a caveat to this interpretation. The levels of dDP and dE2F1/dE2F2 are strongly reduced in the absence of their binding partners. This may help to explain why dDP and de2f1 de2f2 mutant cells have similar properties: these cells may, in essence, lack the same set of proteins. However, this change in levels also means that there may not be sufficient dDP protein in de2f1 de2f2 mutants to perform any functions that are normally independent of dE2F1 or dE2F2. The converse could also be true for dDP-independent functions of dE2F1 and dE2F2. The idea that the levels of partner proteins can change dramatically in mutant animals adds an interesting complication to the interpretation of E2F/DP knockout phenotypes. Potentially, an indirect function of repressor E2F/DP complexes may be to maintain a reservoir of DP components that can be used to partner newly synthesized activator E2Fs. According to the Fly GRID database of two-hybrid interactions (http://biodata.mshri.on.ca/fly_grid/servlet/SearchPage), dE2F1, dE2F2, and dDP have the potential to associate individually with >30 other proteins. At present, there is no evidence that most of these interactions occur in vivo, but we cannot formally exclude the possibility. Nevertheless, the results described here provide compelling evidence that E2F function can be eliminated in Drosophila by removing either dDP or dE2F1 and dE2F2, and hence, we can safely assume that dDP mutants are deficient for E2F regulation, once inherited products are exhausted.

The second major conclusion that we draw from these experiments is that dDP mutant cells are at a strong proliferative disadvantage compared to wild-type cells. A requirement for E2F was seen in the slow proliferation of dDP mutant clones in vivo and the slow proliferation of dE2F1/dE2F2- or dDP-depleted cells in tissue culture. We found that cells in dDP mutant clones progress slowly through S phase and exhibit G2/M defects. Since transient overexpression of cyclin E restores the normal rate of DNA synthesis but does not restore the normal number of M-phase cells, it is unlikely that abnormal G2/M progression is a consequence of slow S phase. In agreement with this, we do not find any evidence of a significant extension of S phase in dDP mutant cells or that the cells enter mitosis with partially replicated DNA or suffer DNA damage and activate a checkpoint-mediated cell cycle arrest. This finding is consistent with the observation that many E2F target genes are expressed at abnormal levels in dDP-depleted cells. The reduced efficiency of DNA synthesis and G2/M phase progression fits with the idea that proteins needed for the control of DNA synthesis and G2/M progression are present at suboptimal levels when E2F regulation is absent (37). Indeed, E2F has been implicated in the regulation of expression of a set of G2/M-specific genes both in Drosophila (8) and in mammalian cells (23, 32). The fact that dDP and de2f1 de2f2 mutant animals progress to such a late stage of development is a testament to the remarkable resilience of animal development, and in many respects, this development disguises the fact that cell proliferation is severely compromised in the absence of E2F regulation. This reduced rate of cell proliferation presumably contributes to the developmental delay that has been noted for dDP or de2f1 de2f2 mutants.

It has been proposed from studies of embryos carrying a hypomorphic dDP allele that as maternally supplied dDP products run out, reduced E2F activity would result in delayed S-phase entry (13). Such an effect may be transient, or cell type specific, because it is clear from the somatic clones described here that dDP mutant cells enter S phase of the second mitotic wave in the eye imaginal disk at precisely the same time as their wild-type neighbors. Although dDP mutant cells have been shown to be unable to control cell cycle phasing when challenged by the ectopic expression of cell cycle regulators (31), the relatively normal timing of S-phase entry in dDP mutants clones and in imaginal disks of trans-heterozygous dDP mutants shows that dDP mutant cells do respond fairly normally to the physiological signals that pattern S-phase entry during development. Clearly, further studies are needed to determine the precise settings in which normal cell cycle control requires E2F, and the FRT dDPa3 and FRT dDPa4 chromosomes described here will be invaluable tools for this work.


   ACKNOWLEDGMENTS
 
We thank our colleagues in the MGH Cancer Center and Molecular Oncology Laboratory for valuable discussions. We are especially thankful to Fred Dick and Marie Classon for help with FACS analysis. We thank Terry Or-Weaver for generously providing dDP mutant alleles and dE2F1 antibody, Carol Seum for the rabbit polyclonal dE2F1 antibody, and Bob Duronio for dDP transgenic flies, for PCNA-GFP alleles, and for the genomic dDP rescue constructs.

This work was supported in part by a Tosteson Postdoctoral Fellowship from MBRC to M.V.F. and CIHR fellowship 210853 to N.-S.M. and by NIH grants GM53203 and PO1 CA95281. M.V.F. is a Leukemia and Lymphoma Society Special Fellow.


   FOOTNOTES
 
* Corresponding author. Mailing address: Massachusetts General Hospital Cancer Center, Bldg. 149, 13th St., Charlestown, MA 02129. Phone: (617) 726-7800. Fax: (617) 726-7808. E-mail: dyson{at}helix.mgh.harvard.edu. Back

{dagger} M. V. Frolov and N.-S. Moon contributed equally to this work. Back


   REFERENCES
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 Introduction
 Materials and Methods
 Results
 Discussion
 References
 
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Molecular and Cellular Biology, April 2005, p. 3027-3039, Vol. 25, No. 8
0270-7306/05/$08.00+0     doi:10.1128/MCB.25.8.3027-3039.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.



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