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Vol. 18, Issue 10, 3733-3740, October 2007
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Institute of Genetics (240), University of Hohenheim, 70599 Stuttgart, Germany
Submitted March 21, 2007;
Revised July 3, 2007;
Accepted July 10, 2007
Monitoring Editor: Carl-Henrik Heldin
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Ruiz i Altaba, 1999). In Drosophila, a large body of work shows that several of these pathways act in concert in the coordination of cell survival and death. For example, overexpression of Notch causes vast overgrowth, whereas inhibition of Notch activity by overexpression of its antagonist Hairless results in tissue loss and apoptosis (Artavanis-Tsakonas et al., 1999; Müller et al., 2005). Indeed, the combined activity of Hedgehog (Hh), Decapentaplegic (Dpp), and Notch is required to promote reentry into the cell cycle after a developmentally regulated G1 arrest in the eye anlagen of Drosophila larvae (for review, see Thomas, 2005). Moreover, it was shown that Hh signaling promotes cellular growth by transcriptional activation of G1 cyclins Cyclin D and Cyclin E (Duman-Scheel et al., 2002). However, to this end, the understanding of the molecular principles that connect these pathways to either control of cell cycle or apoptosis remains largely fragmentary (Thomas, 2005).
Cell cycle entry requires the activity of G1-S cyclins that eventually activate dE2F1, a transcription factor that induces transcription of downstream genes required, e.g., for DNA replication (Duronio et al., 1995). In Drosophila, transcriptional activation of replication-related genes encoding, for example, proliferating cell nuclear antigen (PCNA) or DNA-polymerase 
subunit involves also DNA replication-related element factor (DREF) that recognizes DNA replication-related element (DRE) response elements (Hirose et al., 1993, 2001; Hyun et al., 2005). DREF can be part of a macromolecular complex including TRF2, a TATA-binding protein-related factor that binds to a subset of selected promoters, including one promoter in the PCNA gene (Hochheimer et al., 2002). TRF2 has been isolated from several different organisms, where it is required for transcription of replication-related genes and key developmental genes as well (Hochheimer and Tjian, 2003). The TRF2/DREF complex consists of more than a dozen proteins, including several known chromatin-remodeling components. Three of them confer chromatin activation, whereas two others, including p160, resemble regulators of insulator function (Hochheimer et al., 2002; Hochheimer and Tjian, 2003). Interestingly, p160 was recently found to enhance position effect variegation and hence chromatin silencing and to be associated with interband regions of polytene chromosomes (Eggert et al., 2004). To this end, the biochemical activity and functional specificity of most of the proteins within the TRF2-complex, i.e., their role in transcriptional activation or in chromatin remodeling, however, remain elusive (Hochheimer and Tjian, 2003).
We have isolated the Zn-finger protein p160 as a genetic suppressor of Hairless activity, prompting our interest in its role during Drosophila development and especially during Notch signaling. In vivo RNA interference resulted in tiny larvae and developmental delay, which is why we named the corresponding gene putzig (pzg). We present the first in vivo evidence that pzg is essential for fly survival by regulating cell cycle entry and progression. In addition, we show that pzg encodes a key regulator of the Notch signaling pathway and that it is involved in histone modification and chromatin activation. Interestingly, this activity is independent of DREF, suggesting context dependence of Pzg activity.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Nagel et al., 2002). To induce strong RNAi activation, crosses were performed at 25°C, with the exception of da-Gal4, which was analyzed at 18°C due to early larval death. All stocks used in this study are described in the Supplemental Material.
A pzg full-length cDNA clone (LD15904) was obtained from Open Biosystems (Heidelberg, Germany). A 699-base pair polymerase chain reaction (PCR) product including the C-terminal part of pzg (base pairs of cDNA 2005-2704) was used for the generation of the RNAi-construct; the cloning strategy was according to Nagel et al. (2002), with the exception that pUAST instead of pUASTdsGFP was used. Transgenic flies were generated according to standard methods. Phenotypic analyses were performed with transgenes located on different chromosomes. Most of the data shown were obtained with a second chromosomal insertion line, which behaved qualitatively and quantitatively comparable with all other lines tested. To visualize the expression domain in some of the experiments, UAS-pzg-RNAi (II.) was recombined with enGFP-Gal4.
Antibody Generation and Verification of pzg-RNAi
Guinea pig and rat anti-Pzg antibodies were raised against the C-terminal part of Pzg (base pairs of cDNA 2005-2991) fused to maltose binding protein (PINEDA, Berlin, Germany). Reduction of Pzg protein levels by pzg-RNAi was measured by Western analyses of protein extracts from da-Gal4 > UAS-pzg-RNAi larvae versus da-Gal4 > UAS-lacZ control larvae: 100 first instars were frozen in liquid nitrogen and shaken rigorously. They were transferred to a new tube, homogenized in 30 µl of SDS-loading buffer (250 mM Tris-HCl, pH 7.6, 0.001% bromphenol blue, 5% [vol/vol] SDS, 5% [vol/vol] 2-mercaptoethanol, and 40% [vol/vol] glycerol), and immediately boiled for 10 min. After 10-min centrifugation, 15 µl of the supernatant was run on standard 10% polyacrylamide gel. Blots were probed with a mixture of guinea pig anti-Pzg (1:1000) and rabbit anti-actin (1:250; Sigma-Aldrich, St. Louis, MO) antibodies. Secondary antibodies, coupled to alkaline phosphatase (1:500), were obtained from Jackson Laboratories (Dianova, Hamburg, Germany). The signals were quantified by densitometry of dry membranes using the WinCAM software program (Cybertech, Berlin, Germany) and normalized with actin control.
Immunoprecipitation, Cross-linked Chromatin Immunoprecipitation (XChIP), and Semiquantitative PCR Analysis
Immunoprecipitations were essentially performed according to Nagel et al. (2005) by using protein extracts of 
500 wild-type embryos and 100 wild-type imaginal discs, respectively. For immunoprecipitations, we used guinea pig anti-Pzg antibodies at 1:100 dilutions; and for detection, we used rat anti-Pzg at 1:500 and rat anti-Ci 2A1 at 1:2 dilutions (Motzny and Holmgren, 1995).
Three independent immunoprecipitations of formaldehyde cross-linked chromatin (XChIP) from 0- to 14-h embryos were done according to Cavalli et al. (1999). Antibodies used were mouse anti-DREF (1:100; Hirose et al., 1996), guinea pig anti-Pzg (1:100), and guinea pig preimmune sera (1:25) as mock control. The precipitated DNA was dissolved in 20 µl of H2O double distilled water, and 1 µl was used for PCR reaction. Primer pairs from the promoter regions of Enhancer of split m8 [E(spl) m8], Notch, Suppressor of Hairless [Su(H)], and vestigial were used in this study. Additionally, primer pairs amplifying the Polymerase gene promoter were used as positive control, whereas the 3' untranslated region (UTR) of E(spl) m8 was chosen as an unrelated negative control. The number of cycles used for amplification was 35, which was within the linear range of the amplification judged by different number of cycles and the amount of input DNA used. We took 7-µl samples every two cycles from the 31st to the 35th cycle to determine the linear range of amplification. Signals were quantified using the histogram function of ImageJ software (http://rsb.info.nih.gov/ij/).
For chromatin immunoprecipitations of first instars (14–15 h after hatching), we used the ChIP Assay Kit according to the manufacturer's protocol (Upstate Biotechnology, Lake Placid, NY). For precipitation, we used rabbit ab850 anti-trimethylated H3-K4 (1:100; Abcam, Cambridge, United Kingdom). Rabbit preimmune sera (1:25) served as mock control. Of the precipitated DNA, 1.5% was used per PCR reaction.
The primer sequences and PCR conditions used to amplify XChIP DNA are described in Supplemental Material.
Antibody Stainings of Tissues and Documentation of Phenotypes
Antibody staining was performed as described in Müller et al. (2005). Antibodies used in this study are listed in Supplemental Material. 5-Bromo-2'-deoxyuridine (BrdU) labeling was for 45 min and standard protocols were followed thereafter (de Nooij et al., 1996). Immunostaining of polytene chromosomes was performed according to Salvaing et al. (2003), by using mouse anti-DREF (1:50) and guinea pig anti-Pzg (1:1000). Secondary antibodies coupled to fluorescein, Cy3, or Cy5 were purchased from Jackson Laboratories. Adult wings were dehydrated in ethanol and mounted in Euparal (Roth, Karlsruhe, Germany).
Wing size was determined using ImageJ software for pixel measurements. For phenotypic comparisons, experiments were done in parallel under identical conditions, and they were repeated at least twice. To test statistical significance, p values were calculated according to Student's t tests (http://www.physics.csbsju.edu/stats/t-test.html). Pictures were taken with Nomarski optics on a Zeiss Axiophot (Carl Zeiss, Jena, Germany). Fly heads were pictured on a WILD stereomicroscope equipped with a Pixera camera (Optronics, Goleta, CA) by using the Pixera Viewfinder, version 2.0, software. Confocal images were acquired with a Zeiss Axioskop linked to a Bio-Rad MRC1024 scanhead by using Bio-Rad Laser Sharp 3.1 software (Bio-Rad, Hercules, CA). Figures were compiled using Corel Photo Paint and Corel Draw software (Corel, Ottawa, ON, Canada).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). This positive effect was not restricted to the eye, because it was likewise observed during wing development. Moreover, cell growth and proliferation induced by an enforced Notch signal was significantly enhanced (20%) by a combined overexpression with EP756. Tissue specific overexpression of EP756 caused a very mild enlargement of the respective tissues on its own. These data suggest a more general role of EP756 in the control of cell proliferation as well as an intimate interaction with Notch signaling. They are shown in Supplemental Figure S1, A–P and W.
EP756 Drives the Expression of putzig, Which Is Essential for Developmental Growth
The EP756 element is inserted 0.1 kb upstream of transcription unit CG7752, and it drives its expression analyzed by in situ hybridization (data not shown). CG7752 encodes a Zn-finger protein. To investigate the function of CG7752 during Drosophila development, we designed an inverted RNAi-construct under UAS-control, allowing us to precisely ablate CG7752 function in a spatially and temporally controlled manner (Nagel et al., 2002). Ubiquitous down-regulation of CG7752 activity, starting already during embryonic development, resulted in late embryonic/early larval death at 25°C. At 18°C, larval development was significantly delayed with respect to growth and developmental timing compared with control larvae. Eventually, the animals died at early pupal stages (Figure 1A). Owing to this influence on growth of the larvae, we renamed the CG7752 gene putzig (pzg), which is German for tiny/droll.
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160 kDa, which is considerably higher than the predicted molecular weight of 110 kDa (Figure 1F). This is in agreement with other reports (e.g., Hochheimer et al., 2002). In animals depleted for pzg by RNAi, we observed an 80% reduction of endogenous protein levels compared with the wild-type, indicating that our RNAi-construct caused a strong hypomorphic situation (Figure 1, E–G).
pzg Is Involved in Cell Cycle Regulation
Pzg was recently coisolated with a protein complex including TRF2 and DREF, which is involved in the transcriptional activation of replication-related genes (Hochheimer et al., 2002; Hyun et al., 2005). Because a direct involvement of pzg in up-regulation of replication-related genes has not been demonstrated yet, we investigated whether pzg is required for cell cycle entry and progression. Indeed, depletion of pzg activity interfered strongly with DNA replication compared with untreated tissue (Figure 2, A–A'). Moreover, the number of cells entering mitosis was strongly reduced within the affected area (Figure 2, B–B'). In support of a rather direct requirement of Pzg for cell cycle control, expression of S phase-promoting factors such as dE2F1 (Figure 2, C–C'), Cyclin A (Figure 2, D–D'), or Cyclin D (data not shown) was likewise diminished. These observations indicate that, in the absence of pzg activity, cells are hampered to enter the cell cycle.
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pzg Is Required for Notch Target Gene Activation
Originally, we had isolated pzg as a positive genetic modifier of Notch signaling. Accordingly, pzg could regulate Notch-mediated growth control, supported by the observation that overexpression of pzg not only suppresses tissue loss caused by a reduction but also enhances tissue overgrowth caused by a gain of Notch signaling activity (Supplemental Figure S1, A–P, W). Moreover, pzg-RNAi strongly increased both the penetrance (from 55 to 100%) and the severity of the haploinsufficient wing phenotype of N5419/+ mutants (Figure 3, A–A'). If the role of pzg were to positively regulate Notch signaling, loss of pzg should be bypassed by an overactive Notch signal. Indeed, introducing either a gain-of-function Notch allele (NAxM1) or a loss-of-function allele of the Notch antagonist Hairless (H1), both rescued the growth defects caused by pzg-RNAi to near wild-type morphology and size (Figure 3, B–D). Thus, subtle enhancement of Notch signaling activity in a pzg mutant background yields a growth signal that is sufficient to overcome the respective cell cycle defects. These genetic experiments suggest that pzg acts upstream of Notch. In this case, we would expect a change of Notch activity in pzg-RNAi mutant tissues. Therefore, we examined the expression of several Notch target genes in pzg-RNAi mutant tissues. In wing imaginal discs, cut, Enhancer of split m8, and vestigial are expressed along the prospective wing margin, and their expression depends directly on Notch activity (Lecourtois and Schweisguth, 1995; Neumann and Cohen, 1996). We found that cut, m8-lacZ, and vgBE-lacZ expression are strongly reduced or even abolished in pzg-RNAi mutant cells (Figure 3, E–J'). In summary, Pzg positively regulates Notch signaling, and loss of pzg activity results in a lowered Notch signal.
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; Giraldez and Cohen, 2003). These results are compatible with a model, whereby the major function of pzg is to up-regulate G1-S cyclins through the stimulation of Notch signaling. In this case, loss of tissue caused by an inhibition of Notch activity, e.g., through overexpression of the Notch antagonist Hairless, should be suppressed to the same degree no matter whether Pzg, Cyclin D, or Cyclin E is overexpressed. As noted above, pzg was isolated as a suppressor of Hairless-mediated tissue loss. We observed a significantly better rescue with Pzg compared with either Cyclin D or Cyclin E, and no rescue with String, indicating that the effects of Pzg on proliferation cannot be explained solely through activation of Notch. The phenotypic enhancement of pzg-RNAi caused by the reduction of Cyclin E dose (Figure 2E) can be rescued by a concurrent reduction of Hairless gene dose (Supplemental Figure S2, D–K). These results can be explained by a dual input of Pzg, activation of Notch signaling as well as activation of cell cycle regulation.
Pzg Regulation of Notch Signaling Is Independent of DREF
Our data show that Pzg is required for proper expression of Notch target genes (Figure 3, E–J'). As a component of the TRF2/DREF complex, Pzg might exert its function by direct transcriptional activation. The TRF2/DREF complex is targeted to DNA via DRE elements located in the proximity of target promoters. There are potential, albeit variant DRE recognition sites in the promoter regions of Notch, Su(H), members of the E(spl) complex as well as of vestigial (vg) (Supplemental Figure S3). To address a potential binding of DREF to these sites in vivo, we performed XChIP with anti-DREF and anti-Pzg as well as unrelated antibodies (as negative controls). As an additional control, we tested the 3' flanking region of E(spl)m8 that lacks DRE-like sequences. We found no evidence for DREF binding on any of the potential DRE elements in our tested genes nor on the 3' UTR of E(spl)m8 (Figure 4A and Supplemental Figure S3), whereas we could easily confirm DREF and Pzg presence on the promoter of DNA-Pol gene (Hirose et al., 1993) (Figure 4A and Supplemental Figure S3). Interestingly, we were able to precipitate chromatin of the regulatory region of vg and of E(spl)m8 with anti-Pzg antibody, in agreement with a DREF-independent binding of Pzg at the promoter regions. Apparently, Pzg is involved in the activation of these Notch target genes independently of DREF (Figure 4A and Supplemental Figure S3). To confirm this conclusion, we analyzed the expression levels of the Notch target genes cut, E(spl)m8 and vg after DREF knockdown in the developing wing. We made use of two UAS-DREF-RNAi lines of different origin that had been functionally analyzed earlier and that behaved largely identically in our assays (Yoshida et al., 2004; Hyun et al., 2005). In contrast to the lowered Notch target gene expression observed in pzg mutant tissues, we did not detect any changes in the DREF mutant territories (Figure 4, E and F). Moreover, we compared the distribution of DREF and Pzg on polytene chromosomes of salivary glands. Both, DREF and Pzg, are associated with hundreds of sites (Hart et al., 1999; Eggert et al., 2004), but they do not completely overlap. Overlap of Pzg and DREF was detected, e.g., at the Pol locus (Melov et al., 1992; Hirose et al., 1993; Hyun et al., 2005), but not at the E(spl)m8 or vg loci (Figure 4, B–D''), underscoring our hypothesis that Notch regulation is independent of DREF. No ectopic expression of Notch target genes was observed as a consequence of overexpression of Pzg during larval wing development. Therefore, we conclude that Pzg is required but not sufficient for the activation of Notch signaling (Supplemental Figure S4, A and B).
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; Xiao et al., 2001; Badenhorst et al., 2002; Hochheimer et al., 2002). Several recent studies reveal a close functional connection between chromatin remodeling/modifying complexes and signal transduction pathways including Notch (Bray et al., 2005; Eissenberg et al., 2005; Müller et al., 2005). Because pzg was copurified in the same complex, we reasoned that it might have a further role in chromatin activation, thereby positively regulating Notch signaling. In this case, loss of Pzg should interfere with hallmarks of activated chromatin, such as trimethylation of histone H3 on lysine 4 (H3K4me3), which is typically found on transcriptionally active loci (Lachner and Jenuwein, 2002). In fact, pzg down-regulation caused a strong reduction of the H3K4me3 signal (Figure 5, B, D, and D'). This was independent of apoptosis, because H3K4me3 levels were unaltered in a DIAP1 gain-of-function background. Notably, we could not detect any changes in H3K4me3 distribution in DREF-RNAi mutant cells (Figure 5, B and C), arguing for a DREF-independent role of pzg in chromatin activation.
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promoter (data not shown). These results strongly argue for an important role of Pzg in chromatin activation at Notch target genes.
pzg Is Not Involved in Hh-mediated Cell Growth and Proliferation
Because our data suggest that pzg activates Notch signaling via chromatin activation, we might expect a positive influence on other signaling pathways as well. We decided to investigate Pzg's requirement for the Hh signaling pathway. In Drosophila, Hh signaling can induce vast overproliferation during imaginal development, and it promotes the expression of Cyclin E and Cyclin D (Duman-Scheel et al., 2002). Hence, Pzg might as well act upstream of Hh signaling, similar to that seen for Notch. In this case, we would expect that removal of pzg activity interferes with the expression of hh, the activation of its downstream effector Cubitus interruptus (Ci), or the target gene dpp, which is not the case (Figure 3, K–N' and Supplemental Figure S4, H–H''). Alternatively, Pzg might serve as cofactor for Ci-mediated growth regulation, as suggested from a large-scale yeast two-hybrid assay (Hermjakob et al., 2004; http://pim.hybrigenics.com). However, we failed to confirm a direct protein interaction in vivo by coimmunoprecipitation experiments by using embryonic and larval protein extracts (data not shown). Although Pzg seems not to be a molecular interaction partner of Ci, one might envisage scenarios in which Pzg activity is still essential for Ci function, e.g., by enhancing Ci-effects on responsive promoters. In this case, loss of pzg might interfere with Ci-activity. Overexpression of Ci results in a considerable overproliferation, e.g., of the adult wing, which is largely independent of pzg. The observed overproliferation involves transcriptional up-regulation of both Cyclin D and Cyclin E, which, however, remained unaltered in a pzg-RNAi mutant background (Supplemental Figure S4, E–G). Hence, our results show that pzg activity is not required for Hh signal transduction and growth regulation.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). TRF2 allows transcription initiation from selected promoters independently of TFIID (Hochheimer et al., 2002; Hochheimer and Tjian, 2003). DREF is a positive transcriptional regulator of cell cycle and replication-related genes, and it may guide TRF2 to the PCNA and DNA-polymerase promoters (Hochheimer et al., 2002). Assuming promoter recognition or binding requires Pzg contained within the TRF2/DREF complex, depletion of Pzg might destroy the complex or reduce its activity, easily explaining the dramatic proliferation defects. However, we note that only a subset of promoters containing DREF binding sites involves activation through TRF2, suggesting that DREF can act independently of TRF2 (Hochheimer et al., 2002). Moreover, we find Pzg activity independently of DREF, indicating that TRF2/DREF complex components can act either alone or in conjunction with other factors.
The TRF2/DREF complex contains several proteins involved in chromatin remodeling (Hochheimer et al., 2002). Notably, Pzg and one other TRF2/DREF component p190 are reminiscent of factors implicated in insulator function (Bell et al., 2001). In accordance, Pzg activity has been associated with position effect variegation and chromatin silencing (Eggert et al., 2004). In contrast, our assays reveal an essential function of Pzg in retaining robust K4-trimethylation of histone H3, which is directly associated with open chromatin structures (Lachner and Jenuwein, 2002). In accordance with our findings, EP756 was recently identified as a suppressor of the cut allele ctK. This cut mutation is caused by the insulator activity of a gypsy retrotransposon, which can be relieved by EP756 overexpression (Krupp et al., 2005). We show here that EP756 drives Pzg expression in support of the notion that Pzg's epigenetic activity overcomes gypsy insulator function.
Three of the proteins found in the TRF2/DREF complex have been identified previously in the nucleosome-remodeling factor NURF, which consists in total of four subunits (Tsukiyama and Wu, 1995). NURF is associated with chromatin activation by facilitating transcription of chromatin in vivo (Tsukiyama and Wu, 1995; Xiao et al., 2001). In fact, mutations in Drosophila ISWI, the catalytic subunit of NURF, and other nucleosome remodeling complexes caused phenotypes that are very reminiscent of pzg-RNAi–induced defects (Deuring et al., 2000). Because DREF down-regulation has no effect on trimethylation of H3K4, it seems unlikely that the TRF2/DREF complex as a whole is involved in chromatin activation. Instead, Pzg may be part of a NURF-like chromatin-remodeling complex, depending on the developmental context.
Apart from a role in proliferation, we have uncovered an important role for Pzg as positive regulator of Notch signaling. Interestingly, we found that Pzg binds to chromatin in the regulatory region of the Notch target genes E(spl)m8 and vg. This regulation is independent of DREF: albeit DREF binding sites are common to Drosophila promoters (Ohler et al., 2002), neither Notch nor Notch target genes that we have investigated are transcriptional targets of DREF. Thus, reduced transcriptional activity of Notch target genes in pzg-RNAi mutant cells is due to a DREF-independent role of Pzg. Alternatively, Pzg could facilitate formation of the transcriptional activator complex that is assembled on Notch target promoters involving intracellular Notch itself (Schweisguth, 2004; Nam et al., 2006; Wilson and Kovall, 2006). By using the yeast two-hybrid system, we have tested several Notch pathway members; however, we failed to detect any binding to Pzg (our unpublished data). We propose that Pzg has a dual function that is effected differently. On one hand, it activates proliferation-related genes in conjunction with TRF2/DREF, and on the other hand, it activates Notch signaling by chromatin activation independently of DREF.
Several lines of evidence support the idea that Notch signaling is particularly susceptible to chromatin remodeling. For example, Notch transcriptional activity requires the histone-modifying enzyme dBre1 that is indirectly required for K4-methylation of histone H3 (Bray et al., 2005). Moreover, chromatin-modifiers were also shown to potentiate Notch activity during Drosophila wing development (Eissenberg et al., 2005; Gause et al., 2006). Finally, general transcriptional regulators and chromatin remodeling factors were found in several independent genetic screens to influence Notch signaling, indicating to a role of pzg in linking Notch to chromatin remodeling (Eissenberg et al., 2005; Müller et al., 2005). The bimodal activity of Pzg onto both cell cycle genes and Notch signaling provides further insight into the complex interplay between cell proliferation and differentiation in the fly.
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ACKNOWLEDGMENTS |
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Footnotes |
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The online version of this article contains supplemental material at MBC Online (http://www.molbiolcell.org). 
Address correspondence to: Anja C. Nagel (anjnagel{at}uni-hohenheim.de)
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