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First published online 16 June 2009
doi: 10.1242/jcs.042861

Journal of Cell Science 122, 2393-2401 (2009)
Published by The Company of Biologists 2009
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Research Article

Cul4 and DDB1 regulate Orc2 localization, BrdU incorporation and Dup stability during gene amplification in Drosophila follicle cells

Hsiu-Chen Lin1,2, June-Tai Wu3,4, Bertrand Chin-Ming Tan5 and Cheng-Ting Chien1,2,*

1 Graduate Institute of Life Sciences, National Defense Medical Center, Taipei, Taiwan
2 Institute of Molecular Biology, Academia Sinica, 128, Sec No. 2, Academia Road, Taipei 115, Taiwan
3 Department of Medical Research, National Taiwan University Hospital, No. 7, Chung San South Road, Taipei 115, Taiwan
4 Gratuate Institute of Molecular Medicine, Medical College, National Taiwan University. No. 1, Ren-Ai Road Section 1., Taipei 100, Taiwan
5 Department of Life Science, College of Medicine, Chang Gung University, 259 Wen-Hwa 1st Road, Kwei-San, Tao-Yuan 333, Taiwan

* Author for correspondence (e-mail: ctchien{at}gate.sinica.edu.tw)

Accepted 1 April 2009


   Summary
Top
 Summary
Introduction
 Results
 Discussion
 Materials and Methods
 References
 
In higher eukaryotes, the pre-replication complex (pre-RC) component Cdt1 is the major regulator in licensing control for DNA replication. The Cul4-DDB1-based ubiquitin ligase mediates Cdt1 ubiquitylation for subsequent proteolysis. During the initiation of chorion gene amplification, Double-parked (Dup), the Drosophila ortholog of Cdt1, is restricted to chorion gene foci. We found that Dup accumulated in nuclei in Cul4 mutant follicle cells, and the accumulation was less prominent in DDB1 mutant cells. Loss of Cul4 or DDB1 activity in follicle cells also compromised chorion gene amplification and induced ectopic genomic DNA replication. The focal localization of Orc2, a subunit of the origin recognition complex, is frequently absent in Cul4 mutant follicle cells. Therefore, Cul4 and DDB1 have differential functions during chorion gene amplification.

Key words: Cul4, DDB1, Proliferation, DNA replication, pre-RC, Dup, Cdt1, Orc2, Chorion gene amplification


   Introduction
Top
 Summary
 Introduction
Results
 Discussion
 Materials and Methods
 References
 
To ensure one round of DNA replication per cell cycle, several pathways function redundantly to control the `licensing' event, a process that includes formation of the pre-replication complex (pre-RC) in the G1 phase (Arias and Walter, 2007Go; Blow and Dutta, 2005Go; DePamphilis, 2003Go; Machida et al., 2005Go). During cell cycle progression from mitosis to the G1 phase in higher eukaryotic cells, the origin recognition complex (Orc) localized at the replication origins recruits Cdc6, Cdt1 and the minichromosome maintenance (MCM) complex sequentially, thus forming the pre-RC. With the initiation of DNA replication in the S phase of eukaryotic cells, the MCM complex functions as the DNA helicase to unwind the double helix, thus allowing DNA replication to move forward (Labib and Gambus, 2007Go). By completion of the S phase, the pre-RC is disassembled, with the detachment of some Orc proteins from the chromatin and the degradation of Cdt1 in Xenopus and mammalian cells (Arias and Walter, 2007Go; Blow and Dutta, 2005Go; DePamphilis, 2003Go; Machida et al., 2005Go). In addition, excess CDC6 is exported to the cytosol by a p21-dependent mechanism during S phase in human cells (Kim et al., 2008Go). In Xenopus and mammalian cells, Cdt1 is also inactivated in the G2 and M phases through binding to geminin, another licensing factor in DNA replication control (Saxena and Dutta, 2005Go; Xouri et al., 2007Go). Together, these regulatory mechanisms ensure that the genome replicates only once per cell cycle.

The Cul4-DDB1 ubiquitin ligase mediates polyubiquitylation of many crucial regulators in the eukaryotic cell cycle (Abbas et al., 2008Go; Dai and Wang, 2006Go; He et al., 2006Go; Kim et al., 2008Go; Lee and Zhou, 2007Go; Nishitani et al., 2008Go; Shibutani et al., 2008Go). Previous studies showed that polyubiquitylation by this ligase is required for Cdt1 degradation during the S phase in C. elegans and human cells (Kim and Kipreos, 2007Go; Nishitani et al., 2006Go). Cdt1 docking at the chromatin recruits Cdt2, the substrate recognition component of the Cul4-DDB1 ligase complex in Xenopus egg extracts and human cells (Arias and Walter, 2006Go; Senga et al., 2006Go). Although Cdt1 degradation is regulated by the Cul4-DDB1 complex during the S phase to ensure DNA replication precisely once per cell cycle (Wohlschlegel et al., 2000Go), it is not clear how the Cul4-DDB1 ligase is involved in the DNA replication process.

Chorion gene amplification in Drosophila follicle cells is a model system to explore the regulatory mechanisms in DNA replication. Amplification of chorion genes during oogenesis is a strategy to meet the mass demand of gene products such as eggshell proteins (Calvi et al., 1998Go; Calvi and Spradling, 1999Go; Cavaliere et al., 2008Go; Claycomb and Orr-Weaver, 2005Go; Tower, 2004Go). During oogenesis, follicle cell precursors derived from stem cells in the germarium proceed with six rounds of mitosis, followed by four rounds of endocycle to reach a polyploidy of 16C DNA content. In these polyploidy follicle cells, chorion genes are specifically amplified further by repetitive firing at DNA replication origins. Two major clusters of chorion genes that are amplified, named Drosophila amplicons in follicle cells (DAFC) located at 66D (DAFC-66D) and 7F (DAFC-7F), and two newly defined DAFCs, DAFC-62D and DAFC-30B, are amplified 60- to 80-, 20-, 6.5- and 4-fold, respectively (Claycomb et al., 2004Go; Delidakis and Kafatos, 1989Go; Spradling, 1981Go). The chorion gene amplification system takes the advantage of easy visualization of replication-related proteins at the amplification foci by immuno-detection. In the initiation of amplification, the Orc complex and the Drosophila Cdt1 ortholog Dup localize at gene amplification foci in stage 10B follicle cells (Royzman et al., 1999Go; Whittaker et al., 2000Go). Moreover, the elongation step in DNA replication is readily detectable when replication proteins Dup and PCNA travel away from the chorion origins in stage 13 (Calvi et al., 1998Go; Claycomb et al., 2002Go).

Equally important for chorion gene amplification are the cis- and trans-regulatory factors (Claycomb and Orr-Weaver, 2005Go). Several cis elements, named amplification enhancing regions (AERs) and amplification control element enhancers (ACEs), are also required for amplification (Lu et al., 2001Go; Zhang and Tower, 2004Go). Some of these ACEs and AERs contain identifiable origins, such as ACE3 and Ori-β that are bound by Orc2 in vivo (Austin et al., 1999Go). Moreover, the trans regulators Myb and E2F transcriptional complexes bind ACE3, and the Myb complex also binds Ori-β. Defects in those transcription factors result in impaired gene amplification (Beall et al., 2004Go; Beall et al., 2002Go; Bosco et al., 2001Go; Cayirlioglu et al., 2001Go; Cayirlioglu et al., 2003Go; Royzman et al., 1999Go).

In this study, we have generated Cul4 mutant alleles and found that Cul4 mutations are associated with defects in cell proliferation in larval stages and DNA replication in the follicular tissue during the gene amplification stage. We also generated DDB1 mutants to examine the involvement of Cul4 and DDB1 in regulating the protein stability of Dup. In addition, gene amplification shown by BrdU incorporation assays was found to have complex defects in Cul4 and DDB1 mutants. Localization of the pre-RC component Orc2 at replication foci was also disrupted in Cul4 mutants. The similarities and differences between Cul4 and DDB1 mutant phenotypes reveal their differential functions during gene amplification.


   Results
Top
 Summary
 Introduction
 Results
Discussion
 Materials and Methods
 References
 
Cul4 mutants are larval lethal
The Drosophila genome includes one Cul4 homolog (CG8711) with an ORF of 821 amino acids that share 54-60% identity with mammalian Cul4A and Cul4B. To address how Cul4 functions in vivo, we isolated four Cul4-null alleles: Cul4G1-3, Cul4L2-1, Cul4G3-5 and Cul4JJ11 (see Fig. 1A). The first three alleles are deletion mutants in which various lengths of the Cul4 N-terminus are truncated and Cul4JJ11 carries a nonsense mutation at Trp199. These four alleles failed to complement each other and the deficiency chromosome Df(2R)Exel6056 (44A4;44C2) that uncovers CG8711. Homozygous mutants for these Cul4 alleles were arrested in the second-instar larval stage (Fig. 1C). To characterize Cul4 expression in these mutants, we generated polyclonal antibodies against Cul4 and tested their specificity for Cul4. Western blot analysis of second-instar larval extracts by immuno-purified Cul4 antibodies detected two signals near the molecular mass of 100 kDa, slightly more than the predicted 94 kDa for Cul4 (Fig. 1B). Whereas the lower size signal remained constant in Cul4 mutants and is therefore a non-specific signal, the greater size signal that was almost undetectable in these Cul4 mutants represents Cul4 expression (Fig. 1B, left panel). This result also indicates that these Cul4 alleles are null. In addition, we also utilized the P-element allele KG02900 that localizes at 5'UTR as a hypomorphic mutant for subsequent Cul4 mutant analysis (Fig. 1A). Homozygous KG02900 larvae expressed lower but significant levels of Cul4 proteins (Fig. 1B, right panel) and survived to later stages than homozygous null mutants (not shown).


Figure 1
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  		Fig. 1. Characterization of Drosophila Cul4 mutants. (A) Scheme of the Cul4 genomic region. Arrows represent three genetic loci, Cul4 (CG8711) in orange, and flanking CG18316 and CG11210 in green. P-elements GE17790 and KG02900 inserted in CG8711 are marked in blue and black. Deleted chromosomal regions in three Cul4 alleles isolated in this study are shown as grey boxes. Black asterisk indicates the position of the single nucleotide substitution in the EMS mutant allele JJ11. (B) Western blots of extracts prepared from late second-instar larvae. Upper panel shows Cul4 protein expression, detected by anti-Cul4 antibodies, in wild-type (lane 1) and in four Cul4 mutants (lanes 2-5). Right panel compares Cul4 expression in WT, KG02900/+ and KG02900/KG02900. Upper arrows indicate specific Cul4 signals of 100 kDa, asterisks indicate nonspecific bands below the Cul4 signal. Lower panel shows the loading control by detecting the anti-{alpha}-tubulin antibody. (C) Size comparison of the homozygous mutant for the L2-1 allele and wild type at 5 days AEL. The wild-type larva (right) is in the third-instar stage whereas the Cul4 mutant (left) remains at the second-instar stage, as judged from the pattern of their mouth hocks. (D-E'') Wing imaginal discs of mosaic clones for Cul4G1-3 (D-D'') or KG02900 (E-E''). Cul4 levels (magenta) are almost undetectable in Cul4G1-3 or greatly reduced in KG02900 clones. Clone cells are negative for the GFP signal (green). Boxed areas are enlarged in the right lower corner. Cul4 is abundant in nuclei, as shown by the colocalization with nuclear GFP signals indicated by arrow in D'. Scale bars: 20 µm.

 
In mutant clones homozygous for Cul4G1-3 generated in wing discs, Cul4 proteins were depleted to a background level (Fig. 1D-D''). Expression of Cul4 proteins in homozygous KG02900 wing-disc clones was reduced, but still remained higher than that in the Cul4G1-3 clone (Fig. 1E-E''). In addition, the clone size was larger for KG02900 compared with Cul4G1-3 clones (see Fig. 1E,D). These results suggest that KG02900 behaves as a partial loss-of-function allele for Cul4.

Cul4 mutant cells have severe proliferation defects
When induced between 48-66 hours after egg laying (AEL), homozygous mutant clones for Cul4G1-3 were hardly recovered in wing discs of late third-instar larvae. This result indicates a requirement for Cul4 in cell proliferation or survival. Small Cul4G1-3 clones, however, were successfully recovered when induced between 66-92 hours AEL (Fig. 2A). These Cul4G1-3 clones were significantly reduced in size compared with the twin clones of Cul4+ (Fig. 2B).


Figure 2
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  		Fig. 2. Cul4 mutants are defective in cell proliferation. (A) A wing disc carrying mosaic clones for the Cul4G1-3 allele. Note that the bright GFP signals are twin-spot Cul4+/+cells and the dimmer GFP signals are heterozygous Cul4+/G1-3 cells. Homozygous Cul4G1-3/G1-3 mutant cells lack any GFP signals. (B) Total areas for homozygous Cul4 clones and twin-spots in individual pairs were quantified for seven discs. Clone induction was performed at 3 days AEL, and analyzed at third-instar larva stage. (C-E') Wing discs of MARCM clones labeled with mCD8-GFP (C-E) and cleaved caspase 3 signals in (C'-E') for Cul4+(C,C'), Cul4L2-1 (D,D'), and Cul4L2-1 with p35 expression (E,E'). Boxed areas are enlarged in upper right corners. (F,G) Wing-discs carrying MARCM clones for Cul4+(F) and Cul4G1-3 (G) show BrdU incorporation (magenta). Scale bars: 20 µm. Arrows indicate cells labelled with both mCD8-GFP and BrdU signals. (H) Statistics for the ratio of the number of BrdU-incorporated and GFP-positive cells to the total number of GFP-labeled cells in a wing disc. A total of 1561 GFP-labeled cells from 17 Cul4+/+ wing discs and 922 GFP-labeled cells from 7 Cul4G1-3/+ wing discs were scored. Results are shown as mean ± s.e.m. Statistically significance was calculated using the Student's t-test, *P<0.05.

 
To test whether the growth defect in Cul4 mutant clones was caused by defects in cell survival, cell proliferation or both, MARCM clones of Cul4+ and Cul4L2-1 were generated with cell morphology marked by membrane-tethered GFP, mCD8-GFP. Similarly to Cul4G1-3 clones, Cul4L2-1 MARCM clones were much smaller in size than Cul4+ clones generated via the same procedure (Fig. 2C,D). Interestingly, cells within mutant clones displayed a rounded and shrunken morphology with irregularly condensed membrane labeled with mCD8-GFP (Fig. 2D). In addition, active caspase 3 could be detected in Cul4L2-1 mutant cells (Fig. 2D'). These typical features of apoptotic cells were not seen in the large Cul4+ MARCM clones (Fig. 2C,C'). To test whether apoptosis contributes to the reduction in Cul4L2-1 clone size, the baculoviral apoptotic inhibitor p35 (Callus and Vaux, 2007Go; Zhou et al., 1998Go) was expressed by the tub-GAL4 driver while Cul4 mutant clones were generated in wing discs. Cul4L2-1 cell morphology was restored by the expression of p35 while retaining low levels of caspase 3 expression (Fig. 2E,E'). However, Cul4L2-1 MARCM clones remained small, with fewer cells than in Cul4+ clones, indicating that cell proliferation is affected by the lack of Cul4 activity.

The cell proliferation defect in Cul4 mutant wing-disc cells could be caused by cell-cycle arrest in the G1 phase, which was evident in S2 cells with Cul4 dsRNA knockdown (data not shown) (Higa et al., 2006bGo). To examine whether S phase progression is also affected in Cul4 mutants, the BrdU incorporation assay was performed for wing imaginal discs that included either Cul4+ or Cul4G1-3 MARCM clones (Fig. 2F,G). The wing disc is formed by a monolayer of epithelial cells, which take 4 days to develop from 50 original primordial cells to the final 50,000 cells with a cell-doubling time of 8-12 hours (Prober and Edgar, 2000Go). Third-instar larval wing discs were incubated with BrdU for 1 hour after MARCM-clones had been induced for 28 hours, which accounts for 2-4 cell-doubling times to generate 4-16 cells in individual clones (Fig. 2F). In Cul4+ clones, 13.3% of GFP-positive cells were also positive for BrdU, whereas in Cul4G1-3 clones this ratio decreased significantly to 8.8% (Fig. 2F-H). This result indicates that Cul4 might be required for entry into S phase or for progression of S phase during wing-disc development.

Dup accumulates in Cul4 mutant cells during chorion gene amplification
To specifically examine the Dup levels during S phase progression, we then used the chorion gene amplification system in ovarian follicle cells. Dup nuclear expression in follicle cells is asynchronous in the endocycle stage but is synchronously suppressed in stage 10A (Whittaker et al., 2000Go). When gene amplification is initiated by stage 10B, Dup localization at two major foci DAFC-66D and DAFC-7F is readily detectable and subsequently Dup is retained at DAFC-66D in late stage 10B (Fig. 3A,B) (Whittaker et al., 2000Go). However, in mutant follicle cells homozygous for Cul4G1-3 in early stage 10B, Dup signals at chorion gene amplification foci were easily discernable but with ectopic Dup punctate spots decorating the nuclear periphery (open arrows in Fig. 3A'). In some cases, such ectopic Dup signals were accompanied with reduced or undetectable Dup chorion gene focal signals (Fig. 3A', right open arrow). By late stage 10B, nuclear Dup accumulation in Cul4G1-3 mutant follicle cells reached peak levels and masked the focal signals (open arrows in Fig. 3B'). Interestingly, cytosolic accumulation of Dup was occasionally found (arrow in Fig. 3C'). Accumulation of Dup was observed in almost all Cul4G1-3 cells (Fig. 3D) except those undergoing apoptosis (asterisk in Fig. 3A''). These analyses suggest that Cul4 regulates the Dup protein level in follicle cells during gene amplification stages.


Figure 3
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  		Fig. 3. Dup accumulation in Cul4 mutant cells during chorion gene amplification. (A-C') Dup expression in follicle tissues including some Cul4G1-3 mutant cells (GFP-negative) at the stage of early 10B (A-A') and late 10B (B-C'). Dup (red) accumulates weakly (open arrows in A') in the nuclei (Hoechst 33342 in blue) compared with two major replication foci in neighboring heterozygous cells (arrows) in early 10B stage, and accumulates strongly in mutant nuclei in the late 10B stage (open arrows in B') or even in cytosol (arrow in C') when neighboring wild-type nuclei include only one Dup focal signal (arrows in B'). Asterisk in A'' indicates an apoptotic cell. See key in D for summary. Scale bar: 10 µm. (D) Statistics showing the percentages of Cul4G1-3/G1-3 mutant and twin-spot Cul4+/+ cells in expressing different Dup signals (1-5, see key on right) at stage 10B-11. Results are shown as mean ± s.e.m. of 76 cells for Cul4G1-3 and 156 cells of twin-spot Cul4+ cells from two experiments.

 

Some DDB1 mutant cells accumulate Dup protein during gene amplification
We also determined whether the Dup protein levels are regulated by DDB1. The EY01408 P-element is inserted at the 5' UTR of the DDB1 gene and was used to generate the deletion allele 5-1 that is predicted to truncate the 208 amino acids at the N-terminus (Fig. 4A). Expression of DDB1 mRNA was abolished in DDB15-1 homozygous larvae, indicating that 5-1 is a null allele (Fig. 4B). The EY01408 insertion also abolished DDB1 mRNA expression (Fig. 4B) and homozygous larvae for either EY01408 or 5-1 survived to second instar stage, suggesting that EY01408 behaves similarly to a null allele (also see below).


Figure 4
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  		Fig. 4. Dup accumulation in DDB1 mutant cells during chorion gene amplification. (A) Schematic for the DDB1 genomic region with red arrow representing DDB1 (CG7769) and green arrows for CG7966 and sim. The EY01408 insertion is indicated in blue and the deleted region in the DDB15-1 allele by the grey bar. (B) RT-PCR shows that DDB1 mRNA expression is reduced in homozygous mutants for EY01408 and DDB15-1 compared with wild-type and transheterozygous flies. The internal control is the mRNA level of {alpha}-tubulin. (C-E'') Follicle cells with homozygous clones for DDB15-1 at early stage 10B (C-C'') and 10B (D-D'') and for EY01408 at 10B (E,E''). Arrows indicate homozygous mutant cells (dashed circles) with normal Dup patterns; dashed arrows indicate reduced Dup focal signals; arrowheads indicate undetectable Dup signals; and open arrows indicate elevated Dup levels (see key in F for summary). Hoechst 33342 staining is in blue. Scale bars: 10 µm. (F) Statistics for Dup expression patterns (1-4, see key on right) in homozygous DDB15-1, EY01408 and corresponding twin-spot controls at stage 10B-11. Results are shown as mean ± s.e.m. of two experiments. Number of cells: DDB15-1, 192; twin-spot DDB1+, 238; EY01408, 305; and twin-spot WT, 349.

 

We then examined Dup expression in follicle cells homozygous for EY01408 or DDB15-1 during gene amplification stages. In early stage 10B, the Dup focal pattern was almost normal, without detectable accumulation in DDB1 mutant cells for both alleles tested (Fig. 4C and not shown). By late stage 10B or stage 11, the normal Dup pattern still persisted in a large fraction of mutant cells (arrows in Fig. 4D-E', and quantification in Fig. 4F). Dup nuclear accumulation was seen in small fractions of DDB1 mutant cells (open arrows in Fig. 4D,E, and 16.3% and 9.5% in DDB15-1 and EY01408, respectively). Surprisingly, some DDB1 mutant follicle cells had either diminished (dashed arrows in Fig. 4C-E) or no Dup signals (arrowheads) in gene amplification foci (27.4% in total for DDB15-1). The reduction in Dup focal signals was detected at a very low frequency (2.5%) in Cul4 mutant cells before the nuclear accumulation (right open arrow in Fig. 3A' and quantification in Fig. 3D). The detection of reduced Dup signals at gene amplification foci, if it occurs in Cul4 mutant cells, would be masked by efficient accumulation of Dup in the nucleoplasm. Alternatively, Cul4 and DDB1 might have distinct functions in regulating Dup focal localization. Comparison of the fractions of Cul4 and DDB1 mutant cells that accumulated Dup (cf. Fig. 3D; Fig. 4F) and the stage of onset in Dup accumulation, these results are consistent with the possibility that in addition to DDB1, Cul4 might function through other pathways to regulate nuclear Dup protein levels during gene amplification stages.

BrdU incorporation patterns are altered in Cul4 and DDB1 mutants
It has been suggested that replication stresses can cause Dup accumulation in follicle cells during gene amplification (May et al., 2005Go). Therefore, to examine possible DNA replication defects in Cul4 mutants, we performed BrdU incorporation assay in follicle cells during gene amplification stages. Ovaries carrying Cul4G1-3 mutant clones in follicle tissues were treated for a BrdU pulse at 40-48 hours after clone induction. Two to five BrdU foci, one at DAFC-66D, DAFC-7F and DAFC-62D each, and two at DAFC-30B, were observed in synchronized wild-type follicle cells at stage 10B, and the major BrdU focus DAFC-66D lasted into stage 11 (GFP-positive cells in Fig. 5A-C) (Claycomb et al., 2002Go). During stages 10B-11, whereas there are 32.8% of Cul4G1-3 mutant cells undergoing apoptosis (asterisks in Fig. 5), 11.6% of Cul4G1-3 cells displayed the normal BrdU focal pattern (arrows) and 30.9% mutant cells had reduced or were even devoid of BrdU signals (Fig. 5D, dashed arrows and arrowheads, respectively). If apoptotic cells are disregarded, about 46% of the surviving cells displayed reduced BrdU focal patterns, indicating that Cul4 is required in many follicle cells for BrdU incorporation at focal sites during gene amplification stages.


Figure 5
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  		Fig. 5. Defective BrdU incorporation in Cul4 mutant cells during chorion gene amplification. (A-C'') Follicle cells carrying GFP-negative homozygous mutant clones for Cul4G1-3 (A-B'') or KG02900 (C-C'') show GFP signal (green), incorporated BrdU (red) and nuclear Hoechst 33342 signals (blue). On average, two to four BrdU foci are present in wild-type cells at stage 10B (A-A') and only one main focus is present at stage 11. Complex BrdU patterns are detected in Cul4 mutant cells. Asterisk, apoptotic cells with no BrdU signals but with characteristic condensed DNA distinguished by Hoechst 33342 staining; open arrow, abnormal genomic replication with elevated nuclear BrdU signals; arrowhead, no BrdU signal; dotted arrow, >50% reduction in BrdU focal signals; and arrow, normal BrdU signal (see key in D for summary). Scale bars: 10 µm. (D) Statistics of various BrdU patterns (1-5, see key on right) in Cul4G1-3 or KG02900 mutant and respective twin-spot WT cells at stage 10B-11 are shown as percentages (mean ± s.e.m.) of four (Cul4G1-3) or three (KG02900) experiments. Number of cells: Cul4G1-3, 287; twin-spot Cul4+, 532; KG02900, 492; and twin-spot cells, 531.

 
Notably, Cul4G1-3 cells also displayed abnormal genomic replication phenotypes characteristic of defects in origin specificity, as shown in mutants for Rad3 and the E2F components (Aggarwal and Calvi, 2004Go; Royzman et al., 1999Go). The abnormal genomic replication can be detected by BrdU incorporation in genomic regions outside the chorion amplification focus. We found that 24.7% of Cul4G1-3 mutant cells displayed ectopic BrdU incorporation. The magnitude of this defect varied, ranging from weak to mass BrdU incorporation (open arrows in Fig. 5A,B). Most mutant cells with weak genomic replication phenotypes also had reduced BrdU signals in chorion foci (open arrow in Fig. 5B') and are classified as abnormal genomic replication (Fig. 5D). Strong nuclear BrdU signals in those cells could mask the focal signals (open arrow in Fig. 5A'). Therefore, the reduction in BrdU incorporation at the focal sites might be greater than the percentage we estimated (Fig. 5D).

To examine the effect of the Cul4 gene dosage in chorion gene amplification and suppression of aberrant genomic replication, we then performed the BrdU incorporation assay for follicle cell clones homozygous for the KG02900 chromosome. These hypomorphic Cul4 cells were also normal in their DNA contents as judged from Hoechst labeling (arrowheads in Fig. 5C''), largely excluding the possible defect in the endocycle stage preceding gene amplification. Although KG02900 cells showed a similar percentage of normal BrdU incorporation as Cul4G1-3 cells, apoptosis and abnormal genomic replication were largely suppressed (Fig. 5C-C'',D). Interestingly, the percentage of mutant cells that had incorporated either reduced or no BrdU at chorion gene foci increased to 71.8% and ectopic genomic replication was not detected in these cells. Thus, Cul4 activity is required for proper BrdU incorporation at gene amplification foci, consistent with the notion that Cul4 is involved in the gene amplification process in follicle cells.

We also performed BrdU incorporation assays in DDB1 mutant cells. Similarly to Cul4G1-3 mutant cells, phenotypes of ectopic, diminished or no BrdU incorporation were also detected in DDB15-1 mutant cells (Fig. 7A). The combination of cells with reduced or absent BrdU signals accounts for 48% of the mutant cells, and reaches 63% when apoptotic cells are excluded. In addition, some cells (9.5%) displayed abnormal genomic replication (Fig. 7C). From the analysis of BrdU incorporation patterns, the DDB15-1 allele behaves similarly to the null Cul4G1-3 but not the hypomorphic KG02900 allele. The phenotypic similarity in BrdU incorporation is consistent with the notion that Cul4 and DDB1 function in the same pathway to regulate gene amplification.


Figure 7
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  		Fig. 7. Altered BrdU and Orc2 patterns in DDB15-1 mutants. (A-B'') Follicle cells at stage 10B carrying homozygous mutant clones for DDB15-1 showing signals of GFP (green), Orc2 (red in A,A'), BrdU (red in B,B'), and Hoechst 33342 (blue). In A', asterisks indicate apoptotic cells; open arrow, elevated nuclear BrdU signal; dashed arrow, >50% reduction in BrdU focal signals; and arrow, normal BrdU signal. Diminished Orc2 focal signals in DDB15-1 mutant cells (dashed arrows in B'). Scale bars: 10 µm. (C,D) Quantification of different patterns of BrdU incorporation from two experiments (C) or Orc2 focal localization from four experiments (D) in DDB15-1 mutant and twin-spot wild-type cells showing percentages (mean ± s.e.m.). The numbers of bars represent phenotypes in BrdU incorporation (C) and Orc2 localization patterns (D) as described for Fig. 5D and Fig. 6D, respectively. Numbers of cells scored: 249 DDB15-1 and 270 twin WT cell in BrdU incorporation and 396 DDB15-1 mutant and 547 twin WT cells in Orc2 staining.

 

Orc2 localization at chorion gene amplification foci are altered differentially by Cul4 and DDB1 mutations
Abnormal genomic replication in mutant follicle cells usually signifies a defect in Orc protein localization during chorion gene amplification foci (Aggarwal and Calvi, 2004Go; Cayirlioglu et al., 2001Go; Royzman et al., 1999Go). Next, we examined the localization of the Orc2 protein at gene amplification foci in Cul4 mutants. In wild-type follicle cells, the Orc2 protein is uniformly distributed in nuclei during stage 9 and loads onto replication foci during stage 10A (Royzman et al., 1999Go; Whittaker et al., 2000Go). In late stage 10B, one prominent Orc2 focus was detected in each wild-type follicle cell (GFP-positive in Fig. 6A-B'') (Royzman et al., 1999Go; Whittaker et al., 2000Go). Similarly to what was observed in the BrdU incorporation assay, 27.6% of Cul4G1-3 cells underwent apoptosis (Fig. 6A,D). Orc2 signals in most of mutant follicle cells (42.6%) were either reduced or absent (Figs 6A-B'',D), whereas a very low percentage of cells (6.6%) exhibited low-level nuclear Orc2 accumulation (arrows in Fig. 6A). To avoid the interference of apoptosis or other defects due to the complete absence of Cul4 activity, we analyzed Orc2 localization in homozygous follicle cells for the hypomorphic KG02900 allele (Fig. 1B). Reduction or absence of Orc2 focal signals accounted for most mutant follicle cells (59%) (Fig. 6C,D), indicating that Cul4 is required for normal Orc2 localization at gene amplification foci of follicle cells. We also carried out immunostaining for Orc2 in DDB1 mutant cells. Similarly to what was observed in Cul4G1-3 mutant cells, DDB15-1 cells displayed normal, reduced or no Orc2 localization, in addition to the fraction of apoptotic cells (Fig. 7B,D). However, the percentage of mutant cells with a reduction or absence of Orc2 focal signals accounted for only 22% of cells, and the accumulation of Orc2 protein that was evident in Cul4G1-3 mutant cells was almost undetectable in DDB15-1 cells. These comparisons suggest that Cul4 functions separately to DDB1 in the regulation of Orc2 localization at chorion gene amplification foci.


Figure 6
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  		Fig. 6. Alteration of Orc2 localization at chorion gene foci in Cul4 mutants. (A-C'') Follicle cells carrying GFP-negative homozygous mutant clones for Cul4G1-3 (A-B'') or KG02900 (C-C'') immunostained for GFP (green), Orc2 (red) and Hoechst 33342 (blue). One major focal Orc2 signal is detected at the 10B stage. Asterisk, apoptotic cells with no Orc2 focal signals; open arrow, diffused nuclear or accumulated Orc2 signals (even with reduced focal signals); arrowhead, no Orc2 signals; dashed arrow, reduction in Orc2 focal signals by more than 50%; and arrow, normal Orc2 focal signal (see key in D for summary). Scale bars: 10 µm. (D) Statistics of various Orc2 patterns (1-5, see key on right) in Cul4G1-3 or KG02900 mutant cells at stage 10B-11 shown as percentages (mean ± s.e.m.) of four (Cul4G1-3) or three (KG02900) experiments. Numbers of cells scored: Cul4G1-3, 340; twin-spot for Cul4G1-3: 399; KG02900, 381; and twin-spot for KG02900, 378.

 


   Discussion
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 Summary
 Introduction
 Results
 Discussion
Materials and Methods
 References
 
In this study, we have isolated Cul4 and DDB1 mutants which were larval lethal with growth arrest in the second instar stage, similarly to previous results (Hu et al., 2008Go). We further showed that Cul4 mutant clones in developing wing discs were defective in proliferation and had a reduced number of S-phase cells. To focus on the role of Cul4 during DNA replication and bypass the requirement of Cul4 in G1-S transition (Higa et al., 2006bGo), we analyzed mutant follicle cells during gene amplification stages in follicle cells. We showed that the Dup protein level and Orc2 focal localization were regulated by Cul4 and, differentially, by DDB1. In addition, BrdU focal patterns were defective in Cul4 and DDB1 mutant follicle cells.

Dup accumulation in Cul4 mutant cells during gene amplification
Previous studies have shown the replication-dependent degradation of human and Xenopus Cdt1 by the Cul4-DDB1 E3 ligase, and the replication-coupled recruitment of the DDB1 to chromatin in Xenopus cells (Arias and Walter, 2006Go; Nishitani et al., 2006Go; Senga et al., 2006Go). In this study, we show the requirement of Cul4 for the suppression of Dup protein levels during gene amplification in Drosophila follicle cells. Comparison of Dup nuclear accumulation in Cul4 and DDB1 mutant follicle cells, however, reveals substantial differences (Figs 3 and 4). Almost all Cul4 mutant cells, except those undergoing apoptosis, accumulated Dup in the nucleoplasm in stage 10B, consistent with Cul4 being a dedicated component of the E3 ligase in promoting Cdt1 degradation. By contrast, accumulation of Dup levels was observed in much smaller fractions of two DDB1 alleles we analyzed. These analyses have not excluded the involvement of DDB1 in downregulating Dup protein levels. Compensatory or parallel Cul4-mediated Dup degradation pathways might be present in addition to the Cul4/DDB1-mediated Dup degradation. In agreement with our speculation, a recent study has shown that nuclear accumulation of cyclin D1 during the S phase promotes human Cdt1 stabilization and triggers DNA re-replication. This Cdt1 stabilization could be suppressed by the overexpression of Cul4A and Cul4B but not DDB1 or Cdt2, also an adaptors for the Cul4 ligases, implying the involvement of other adaptors in mediating the Cul4-dependent degradation of Cdt1 (Aggarwal et al., 2007Go).

Involvement of Cul4 and DDB1 in chorion gene amplification
We found that BrdU incorporation at chorion gene amplification foci was reduced or even absent in about half of Cul4G1-3 mutant follicle cells, if apoptotic cells that cannot be scored for their capability in BrdU incorporation were excluded. Similarly, 40% of Cul4G1-3 mutant cells displayed ectopic BrdU incorporation (the abnormal genomic replication group). These cells were not scored for their BrdU incorporation at focal sites because of overall strong nuclear signals. Therefore, the effect of Cul4 on the chorion gene amplification might be underestimated in this Cul4-null allele. Using the Cul4 hypomorphic allele KG02900 in which both the fractions of apoptotic cells and abnormally BrdU-incorporated cells were reduced, the combined percentages for reduced and absent BrdU incorporation combined at chorion gene amplification foci reached more than 70% (Fig. 5D). DDB15-1 mutant follicle cells also displayed a severe phenotype in the BrdU incorporation assay (Fig. 7C). When apoptotic cells were disregarded, cells with a reduction or absence of BrdU incorporation accounted for more than 60% of DDB15-1 mutant cells. Therefore, these BrdU incorporation analyses lend support to the notion that certain processes in chorion gene amplification require both Cul4 and DDB1.

These BrdU foci represent DNA amplification of chorion genes within 100 kb of origins, and the phenotype of absence or reduction in BrdU incorporation could reflect a failure in the initiation of DNA replication, reduced processivity in DNA synthesis or fewer rounds of gene amplification (Claycomb et al., 2002Go). To further support the idea that Cul4 is involved in gene amplification, we took advantage of the dominant-negative Cul4KR mutant in which the neddylation site has been mutated. Acute expression of Cul4KR suppressed BrdU incorporation in almost all follicle cells (supplementary material Fig. S1A,B). When assayed by quantification PCR, gene amplification at chorion foci was strongly suppressed (supplementary material Fig. S1C), supporting a role of Cul4 in the chorion gene amplification process.

Abnormal genomic replication, as inferred from ectopic BrdU incorporation, was observed in Cul4 mutant follicle cells in both Cul4 mutant alleles tested. The percentage of cells with such a defect was reduced in the cells with the hypomorphic mutant allele, indicating that the low level of Cul4 activity partially suppresses this defect (Fig. 5D). Abnormal genomic replication was also detected in DDB1 mutant follicle cells with a lower frequency than in Cul4-null mutants (Fig. 7C). The phenotype of ectopic genomic replication is less likely to be a retarded developmental process in the previous endocycle stage, because these cells with ectopic BrdU signals show normal DNA contents as estimated by Hoechst staining. Ectopic genomic replication might require some prerequisite steps in DNA replication, such as Orc2 localization at replication origins, which is defective in Cul4 mutant cells, thus blocking abnormal DNA replication throughout the genome.

Role Cul4 in controlling Orc2 protein localization
We examined the localization of Orc2, a component of the pre-RC, at chorion gene foci during gene amplification. Mutations in Cul4 caused reduced or no Orc2 localization at gene amplification foci, a prominent phenotype in both G1-3 (59%) and KG02900 alleles (60%) when apoptotic cells were discounted. Failure of proper Orc2 focal localization might represent defects in the initial loading of Orc2 or the maintenance of Orc2 localization at amplification foci. Such Orc2 localization defects were not prominent in DDB1 mutant follicle cells (Fig. 7D).

Consequently, defective Orc2 localization at regular gene amplification foci might evoke ectopic genomic replication in Cul4 mutant follicle cells. Upon co-labeling for Orc2 and BrdU in Cul4 mutant cells, the absence of or reduction in Orc2 signals was found in conjunction with a reduction in BrdU incorporation at gene amplification loci or with abnormal BrdU incorporation throughout the genome. In some cells, normal Orc2 loading was accompanied with abnormal BrdU incorporation (supplementary material Fig. S2). The decoupling of both phenotypes therefore suggests that Cul4 functions distinctively in Orc2 localization and in suppression of abnormal BrdU incorporation during chorion gene amplification.

Many interesting questions remain to be answered regarding how genetic loci are selected for amplification, how the pre-RC is assembled only in specific loci and how other genomic regions are kept silent. Previous evidence suggests that high transcriptional activity of specific loci controls replication origin firing during the gene amplification stage (Claycomb and Orr-Weaver, 2005Go; Tower, 2004Go). Mutants for transcription factors such as E2f2, Dp, Rbf, Myb and Mip130 show increased mRNA and protein levels of replication factors, such as components of the Orc and MCM complexes, and ectopic genomic replication during the gene amplification stage (Bosco et al., 2001Go; Cayirlioglu et al., 2001Go; Cayirlioglu et al., 2003Go; Royzman et al., 1999Go). Mutant follicle cells for Cul4 displayed both Orc2 localization defects and abnormal genomic replication, implying that Cul4 is probably involved in both processes by modulating the transcriptional activity in DNA replication. Interestingly, a recent study suggests that Cul4 targets degradation of the transcriptional activator E2F1 during S phase (Shibutani et al., 2008Go). However, Drosophila E2F1 proteins became abundant in the nucleus (Sun et al., 2008Go) and rich at ACE3 origin DNA during gene amplification (Bosco et al., 2001Go). How this developmental regulation of E2F is involved in Cul4 activity needs further investigation. Some studies suggest that Cul4 functions in histone modification and heterochromatin maintenance (Higa et al., 2006aGo; Hong et al., 2005Go; Jia et al., 2005Go; Kapetanaki et al., 2006Go; Wang et al., 2006Go). We speculate that the Cul4 E3 complex also functions in regulating Orc2 origin localization through a local remodeling of the chromatin structure on ACE3 and Ori-β.


   Materials and Methods
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 Summary
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 Results
 Discussion
 Materials and Methods
References
 
Fly stocks and genetics
GE17790 (renamed G2935 from GenExel fly stocks) located 79 bp upstream of Cul4 5'UTR was used for P-element imprecise excision to generate three independent null alleles, Cul4G3-5, Cul4L2-1 and Cul4G1-3, with deletions of amino-terminal 190, 396 or 340 amino acids, respectively. The Cul4JJ11 allele, with a substitution of W199 by a stop codon, was isolated from an EMS screen and fails to complement the lethality of the Cul4L2-1 allele. These Cul4G1-3, Cul4G3-5, Cul4L2-1 and Cul4JJ11 alleles also failed to complement to each other and Df(2R)Exel6056. DDB15-1 was obtained by P-element imprecise excision form EY01408 (Bloomington Stock Center). The deletion includes 45 bp of DDB1 5'UTR and N-terminal 208 amino acids.

Genotypes for generating mosaic mutant clones in wing discs and follicle cells are hs-FLP122; FRT42D Cul4G1-3 (or other Cul4 alleles)/FRT42D [ubi-nlsGFP] and hs-FLP122; FRT82B DDB15-1 (or EY01408)/FRT82B [ubi-nlsGFP]. Genotypes for generating MARCM (Lee and Luo, 2001Go; Lee et al., 2000Go) clones are: (1) hs-FLP122; FRT42D tub-GAL80/FRT42D[pconD]; tub-GAL4 mCD8GFP; (2) hs-FLP122; FRT42D tub-GAL80/FRT42D Cul4L2-1; tub-GAL4 mCD8GFP; and (3) hs-FLP122; FRT42D tub-GAL80/FRT42D Cul4L2-1; tub-GAL4 mCD8GFP/UAS-p35. To make mosaic mutant clones in imaginal wing discs, 0- to 24-hour-old eggs were collected, heated shocked at 37°C for 30 minutes over 72-96 hours. For making mutant follicle cell clones, 0- to 5-day-old yeast-fed females were incubated at 37°C for 30 minutes, and transferred back to 25°C for 40-48 hours before dissection.

Immunolabeling and microscopy
Imaginal discs were fixed and stained as described previously (Chen and Chien, 1999Go). Ovaries were dissected in Schneider's Insect Medium (GIBCO) and fixed in 200 µl of 4% formaldehyde saturated with 600 µl heptane and 0.25% NP40 for 20 minutes. Rabbit anti-Cul4 polyclonal antibodies were generated against amino acids 1-19 of Cul4 (QCB Co.), and were used at 1:100 dilution for immunostaining and 1:1000 for western blots. Other primary antibodies used in this study were: anti-Dup [1:500-1000 (Whittaker et al., 2000Go)], anti-ORC2 [1:500 (Royzman et al., 1999Go) and anti-{alpha}-tubulin (1:100,000; Sigma). Cy3- and Cy5-conjugated secondary antibodies (1:500-1:1000, Jackson Immuno Research) were used. Images were acquired by the Zeiss LSM510 Meta confocal microscope and processed by LSM510 software.

BrdU incorporation assays
BrdU incorporation assays in wing discs were performed at 66-92 hours after clone induction as previously described (Baker and Rubin, 1992Go). Ovary BrdU labeling was performed as previously described (Lilly and Spradling, 1996Go). Ovaries were collected in Schneider's Insect Medium (GIBCO) on ice, incubated in medium containing 30 µg/ml BrdU (Sigma) at room temperature for 1 hour, and washed with PBS for 5 minutes. These BrdU-incorporated ovaries were fixed in 200 µl of 4% formaldehyde saturated with 600 µl heptane and 0.25% NP40 for 20 minutes, washed with PBT (phosphate-buffered saline containing 0.3% Triton X-100) for 5-15 minutes, and fixed again in 600 µl Buffer B (100 mM KH2PO4/K2HPO4, pH 6.8, 450 mM KCl, 150 mM NaCl, 20 mM MgCl2) + 4% formaldehyde (1:5) for 20 minutes, then washed in PBT three times. These samples were treated with 2N HCl in PBT for 30 minutes, neutralized with 100 mM sodium borate for 5 minutes, and washed three times with PBT for 5-15 minutes. For BrdU detection, these samples were incubated in PBT including 10% normal donkey serum at room temperature for 1 hour and then with the BrdU antibody (1:50-100, BD) at 4°C overnight. After PBT washes, the Cy5-conjugated secondary antibody (Jackson Immuno Research) was added at 1:1000 at room temperature for 2 hours. To double stain with BrdU antibody, a second fixation was performed and the titer of antibodies used was Orc2, 1:500 and BrdU, 1:100.


   Footnotes
 
Supplementary material available online at http://jcs.biologists.org/cgi/content/full/122/14/2393/DC1

We would like to thank Terry L. Orr-Weaver for Dup antibody; Stephen P. Bell for Orc2 antibody; Drosophila Bloomington Stock Center for fly stocks, Ya-Ru Chan for generation of Cul4KR construct; Tzu-Ting Lai for help with EMS screen of Cul4 mutants, Min-Han Chen for microinjection, and Yu-Chin Lin for help with figure preparation, and Margaret Su-chun Ho and Hai Wei Pi for comments on the manuscript. B.C.-M.T. was supported by the National Science Council (NSC). C.-T.C. is supported by grants from NSC and Academia Sinica of Taiwan.


   References
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