Epithelial renewal in the Drosophila intestine is orchestrated by Intestinal Stem Cells (ISCs). Following damage or stress the intestinal epithelium produces ligands that activate the epidermal growth factor receptor (EGFR) in ISCs. This promotes their growth and division and, thereby, epithelial regeneration. Here we demonstrate that the HMG-box transcriptional repressor, Capicua (Cic), mediates these functions of EGFR signaling. Depleting Cic in ISCs activated them for division, whereas overexpressed Cic inhibited ISC proliferation and midgut regeneration. Epistasis tests showed that Cic acted as an essential downstream effector of EGFR/Ras signaling, and immunofluorescence showed that Cic’s nuclear localization was regulated by EGFR signaling. ISC-specific mRNA expression profiling and DNA binding mapping using DamID indicated that Cic represses cell proliferation via direct targets including string (Cdc25), Cyclin E, and the ETS domain transcription factors Ets21C and Pointed (pnt). pnt was required for ISC over-proliferation following Cic depletion, and ectopic pnt restored ISC proliferation even in the presence of overexpressed dominant-active Cic. These studies identify Cic, Pnt, and Ets21C as critical downstream effectors of EGFR signaling in Drosophila ISCs.
EGFR/Ras/MAPK signaling has diverse functions in regulating cell proliferation, growth, differentiation and survival in most animal cells [1]. Abundant studies also indicate that epidermal growth factor receptor (EGFR) activation is a causal driver of many cancers, including breast, lung, brain, and colorectal cancer [2]. Similarly, activating mutations in KRAS and BRAF, which are essential downstream effectors of the EGFR, are among the most common mutations found in a very wide range of human cancers [3,4]. However, despite much study, many questions remain to be answered to fully understand the impact of EGFR and its downstream effectors during normal cell function and in carcinogenesis. As many epithelial cancers arise through dysregulation of the stem cell self-renewal and homeostatic maintenance of the epithelium [5], understanding the precise functions of EGFR signaling in epithelial homeostasis is very important. The Drosophila midgut is an outstanding model system to study the basis of epithelial homeostasis due to its simple structure, similarity to the mammalian intestine, and powerful genetics. As in the mammalian intestine, epithelial turnover in the fly midgut is carried out through a dynamic process mediated by intestinal stem cells (ISCs). ISCs undergo cell division to renew themselves and give rise to transient cells called enteroblasts (EBs), which can further differentiate into either absorptive enterocytes (ECs) or secretory enteroendocrine (EE) cells. When damaged or aged cells are lost from the fly’s gut epithelium, ISCs respond by dividing to replenish the epithelium [6,7, 8]. During this response multiple Drosophila EGFR ligands, namely spitz (spi), vein (vn), and keren (krn) are induced in progenitor cells (EBs and ISCs), visceral muscle (VM) and ECs respectively. Thereby, the EGFR signaling pathway is activated in ISCs. This promotes ISC growth, division and midgut epithelial regeneration [9,10,11]. ISCs defective in EGFR signaling cannot grow or divide, are poorly maintained, and are unable to support midgut epithelial replenishment after enteric infection by the bacteria Pseudomonas entomophila (P. e.) [11] or Erwinia carotovora carotovora 15 (ECC15) [12]. Interestingly, the critical role of EGFR signaling in the Drosophila intestine is consistent with its role during mammalian gut homeostasis and colorectal cancer development [10,11,12,13]. EGFR signaling is required for murine ISC growth [14,15], and the deletion of Lrig1, a negative feedback regulator of EGFR signaling, causes excessive ISC proliferation [16]. Furthermore, adenoma formation in Apcmin/+ mice was severely impaired in a genetic background with partial loss of function of EGFR (Egfrwa2) [17]. Despites its importance, the mechanism by which EGFR/Ras/MAPK signaling promotes ISC proliferation is poorly understood in this cell type. Indeed, despite decades of intensive study, the precise linkage between EGFR/Ras/MAPK signaling and cell growth and division is surprisingly obscure for animal cells in general [3]. Textbook models highlight a prevailing model in which EGFR/Ras signaling controls cell proliferation via a Ras-Myc-CyclinD-Rb pathway [18,19]. While this may have relevance in some human cancers it is clearly not the case in normal Drosophila cells, and so other mechanisms should be sought and characterized. One potentially important downstream effector of EGFR signaling is the HMG-box transcriptional repressor Capicua (Cic). This highly conserved DNA binding factor has been shown to act downstream of receptor tyrosine kinase (RTK) /Ras/MAPK signaling in Drosophila eye and wing imaginal discs, embryos, and ovaries [20,21,22,23] where it regulates diverse RTK-dependent processes including cell proliferation, specification, and pattern formation. Cic orthologs from invertebrate and vertebrate species share two well-conserved regions: the HMG-box, presumed to mediate DNA binding at target promoters [21] and a C-terminal domain [24]. The C-terminal region of Drosophila Cic contains a “C1” motif important for repressor activity, and a “C2” motif that functions as a MAPK docking site responsible for downregulation of Cic following the activation of RTK signaling [25]. It has been proposed that MAPK phosphorylates Cic in its C2 motif, and that phosphorylated Cic is either degraded or re-localized to the cytoplasm [25]. Cic downregulation controlled by Torso and EGFR signaling varies in different Drosophila tissues [24]. For example, Torso RTK signaling, which also works via the Ras/Raf/MAPK pathway, apparently increases the rate of Capicua degradation by promoting its accumulation in the cytoplasm [26]. EGFR signaling has been reported to regulate Cic protein in distinct ways in different tissues. Wing and eye discs cell clones mutant for Egfr or Ras showed elevated levels of Cic protein [20,27]. In the ovary, in contrast, Cic protein localized to the cytoplasm in cells in which EGFR signaling was active, but in nuclei in cells in which EGFR signaling was inactive [25]. A recent study suggested that Cic actually undergoes a two-step process in releasing its target gene repression: slower changes in nuclear localization occur after a faster reduction of Cic repressor activity [28]. In cultured human cells, EGF stimulated dissociation of human CIC from importin-α4 (also known as KPNA3), an adaptor required for the nuclear import of many proteins. But full length GFP-CIC was nuclear even after EGF stimulation, and the N-terminal half of the CIC protein was found to be nuclear, even though it does not bind to importin-α4. Hence the biological significance of the CIC: importin association remains unclear [29]. CIC, the human homolog of Drosophila Cic, has been implicated in several human diseases including spinocerebellar ataxia type 1 (SCA1) neuropathology, oligodendroglioma (OD) [30] and Ewing-like sarcoma [31]. Human CIC is frequently mutated in samples from cancer genome studies such as The Cancer Genomic Atlas (TCGA) (S1 Fig) [32,33]. For instance CIC mutation was reported in 6 out of 7 brain tumors [30], 3 out of 11 breast cancers [34] and 6 out of 72 colorectal cancers [35]. The Drosophila work suggests that in these cases CIC loss might have the same downstream consequences (e. g. cell transformation) as oncogenic activation of the EGFR, RAS or BRAF, but this has not been rigorously evaluated. During RNAi screening we discovered that depletion of Cic in Drosophila’s intestinal stem cells (ISCs) activates these cells for rampant proliferation [11]. Based on previous studies in other fly organs we hypothesized that Cic might act as an obligate repressor downstream of EGFR signaling, itself a central driver of normal ISC proliferation in both flies and mice, as well as in many human colorectal cancers, which are frequently mutant for RAS, BRAF, or CIC. However, until now this hypothesis had not been tested and the underlying mechanisms via which Cic might control ISC proliferation remained undefined. In this report we demonstrate that Cic acts as a critical negative downstream regulator of EGFR signaling to control ISC proliferation. We show that EGFR/Ras activity controls Cic nuclear localization, and we present RNA-Seq and DamID-Seq datasets that together constitute a genome-wide survey of potential Cic target genes in Drosophila ISCs. Our analysis indicated that Cic not only directly regulates cell cycle regulators such as string (cdc25) and Cyclin E, but also the ETS transcription factors pnt and Ets21C, all of which must be de-repressed to activate ISCs for growth and division.
To investigate a potential role for Cic in regulating ISC proliferation, we used the esg-Gal4-UAS-2XEYFP; Su (H) GBE-Gal80, tub-Gal80ts system (henceforth referred as esgts; Su (H) -Gal80) to express UAS-cic-RNAi specifically in ISCs. After 4 days of cic-RNAi induction, a dramatic increase in the number of YFP positive cells (Fig 1A and 1B) and a large increase in ISC mitoses were observed (Fig 1C). Most of the PH3+ cells were YFP+ [YFP+, PH3+ cells = 99. 37% (nmidguts = 10 midguts, ncells = 994), YFP-, PH3+ cells = 0. 63% (n = 10, ncells = 7) ], indicating that Cic regulates ISC proliferation cell autonomously. When we used another ISC-specific driver Dlts (tub-Gal80ts UAS-GFP; Dl-Gal4) to knock down cic in ISCs specifically, we not only detected the same overporoliferation phenotype (S3A, S3B and S3E Fig) but also found that most of mitotic cells were GFP+ (S3F Fig). Increased GFP+ cells and mitoses were also noticed when the esgGal4 UAS-GFP tub-Gal80ts system (henceforth referred as esgts) was used to express UAS-cic-RNAi in ISCs and their undifferentiated daughters, the EBs (S2A–S2B and S3 Figs). To further validate the specificity of this RNAi experiment, GFP-marked ISC clones homozygous for the loss-of-function allele cicfetU6 [22] were generated using the MARCM system [36] (S2C–S2H Fig). The size of marked ISC clones was quantified at intervals after clone induction by measuring GFP-labeled clone areas. cic mutant clones were larger than control clones at all time points assayed (Fig 1D). In addition, the numbers of cells per clone were increased in the cic mutant clones (Fig 1E). To further confirm Cic’s function in the midgut, we generated viable transheterozygotes using three different loss-of-function alleles of cic. cicfetE11 is a P-element insertion mutant, while both cicfetT6 and cicfetU6 are homozygous lethal EMS alleles [22]. In addition to the EGFR-related extra wing vein phenotype reported previously [27], these transheterozygote mutants showed increased mitoses in their midguts (Fig 1I). As the ISCs are the predominant dividing cell type in Drosophila midguts, these data further indicate a role for Cic as an obligate repressor of ISC proliferation. To investigate the respective requirements of Cic in the ISC and EB cell types, the EB-specific driver Su (H) ts [Su (H) -Gal4, UAS-CD8-GFP; tub-Gal80ts] was used to knock down cic in EBs. Increased mitoses were observed after depleting cic in EBs (S3C–S3E Fig). However, in this case only a few GFP+ EBs were observed in mitosis, while most of the dividing cells marked by PH3 were GFP-negative (S3F Fig). These GFP-negative mitotic cells are likely ISCs. These data indicated that Cic has both cell autonomous and non-cell autonomous functions in regulating ISC proliferation. In this study we followed up on Cic’s cell autonomous effects on ISC proliferation, and the non-cell autonomous effect was not investigated further. To determine whether increased Cic function yields a phenotype similar to that of EGFR loss-of-function, we generated transgenic flies harboring UAS-cicΔC2-HA or UAS-cic-HA. CicΔC2 is a Cic derivative carrying a deletion of the MAPK docking site-C2 motif, and has been shown to be a dominant repressor that escapes inactivation by MAPK [25]. Either cic or cicΔC2 were over-expressed in progenitor cells using esgts, and then the flies were fed Pseudomonas entomophila (P. e.) for 12 hours to generate an enteric infection. ISCs from control midguts, which expressed GFP only, showed regeneration-associated proliferation [8]. In contrast both cic and cicΔC2 overexpressing midguts displayed an inhibition of regeneration after 12 hours P. e. infection (Fig 1J). To test if cic or cicΔC2 overexpression could influence turnover of the midgut epithelium we used the esgts F/O system (esg-Gal4; tubGal80ts Act>Cd2>Gal4 UAS-flp UAS-GFP) [11] to mark all the ISC progeny produced during 12 days of cic overexpression. Normally, the posterior midgut epithelium renews it self within about 12 days [8]. Therefore, control midgut epithelia were almost completely replaced by large GFP+ clones that formed during 12 days. However, in the gain-of-function Cic conditions, growth of GFP-marked clones was significantly decreased, indicating that gut epithelial renewal was greatly suppressed (Fig 1H–1J). EGFR activates ISCs for growth and division via Ras/Raf/MAPK signaling. When an activated form of the EGFR (λTOP) [37] or activated Ras (RasV12G) [38] is ectopically expressed in progenitor cells, ISC division is dramatically induced. Conversely, EGFR suppression by inducing Egfr-RNAi, Ras-RNAi, or MEK-RNAi in progenitor cells almost completely inhibits ISC division and growth [11,12]. Furthermore, inhibition of EGFR signaling suppresses the activation of ISC divisions after P. e. infection [10,11]. As demonstrated above, Cic knockdown and overexpression phenocopy these EGFR overexpression or knockdown phenotypes, respectively, suggesting that Cic may act as a downstream effector in the EGFR signaling in ISCs. To test the function of Cic in EGFR signaling we performed epistasis tests. After 2 days of clone induction with the esgts F/O system, control midguts generated only 2-cell clones, whereas clones overexpressing an activated variant of the EGFR, (λtop), grew very large and showed increased ISC division. However, when cic or cicΔC2 was co-overexpressed along with λtop, clone sizes and ISC mitoses were significantly reduced (Fig 2A–2C and 2I). Overexpression of Cic or CicΔC2 could also partially inhibit the ISC growth effects of RasV12S35, an activated allele that can activate RAF/MAPK signaling but not PI3K signaling [38] (Fig 2D, 2H, and 2J). Furthermore, we used esgts to induce Egfr-RNAi, or Ras-RNAi in combination with cic-RNAi. The cic, Egfr or cic, Ras double RNAi animals exhibited increased ISC mitosis relative to controls expressing Ras-RNAi or Egfr-RNAi only (Fig 2E–2G and 2K), indicating that reduced ISC proliferation caused by the inactivation of EGFR signaling can be restored by Cic knock-down. These epistasis data further support the hypothesis that Cic acts as a negative downstream effector of EGFR to regulate ISC proliferation. To understand how EGFR signaling controls Cic in ISCs, we expressed HA-tagged Cic or CicΔC2 protein in midgut progenitor cells (ISCs and EBs). As expected, HA-tagged Cic or CicΔC2 proteins were only detected in nuclei under normal conditions (Fig 3A–3A’ and 3B–3B’). However, HA-tagged Cic protein accumulated nearly exclusively in the cytoplasm when RasV12S35 was co-expressed with it (Fig 3E–3E’). In contrast, CicΔC2 remained in the nucleus even following ectopic RasV12S35 expression (Fig 3F–3F’). A similar but milder re-localization of Cic protein from the nucleus to the cytoplasm was observed following P. e. infection (Fig 3C and 3C’), a treatment known to increase MAPK signaling in the gut [11]. It is interesting to note that CicΔC2 did not completely suppress RasV12S35 induced ISC proliferation, even though it remained localized to nuclei in RasV12S35 expressing cells (Fig 2H and 2J). However, nuclear CicΔC2 lost its characteristic punctate localization in the presence of RasV12S35 expression, and became more diffusely localized in the nucleoplasm (Fig 3F–3F’). These results suggest that, although EGFR signaling controls Cic nucleo-cytoplamic localization via the C2 motif, there may be a second MAPK-dependent mechanism to regulate Cic repressor activity, involving dissociation from chromatin, that is C2-independent. Cic has been studied in several cell types from both Drosophila and humans. In human melanoma cells, CIC represses mRNA expression of the PEA3 subfamily of ETS transcription factors, namely ETV1, ETV4 and ETV5 [29]. In early Drosophila development post-transcriptional down-regulation of Cic by the Torso and EGFR pathways regulates terminal and dorsal-ventral patterning, respectively, by allowing expression of Cic target genes such as huckebein (hkb), intermediate neuroblasts defective (ind), and argos (aos) [39]. However, a genome-wide mapping of Cic target genes has not yet been reported. To identify Cic target genes involved in ISC growth and proliferation we profiled Cic binding throughout the genome using the “TaDa” (Targeted DamID) ” technique. The TaDa method involves low-level expression of a GAL4-inducible Dam methylase-fusion protein in a specific cell type, enabling cell-specific profiling without cell isolation [40,41]. Here, we induced a low level of Dam-only or Dam-Cic fusion protein in progenitor cells (ISC & EB) using the esgts system and a 24-hour induction. Genomic DNA was extracted from isolated midguts, digested with Dpn I, which cuts only methylated GATCs, and amplified. The amplified gDNA fragments were subjected to high-throughput sequencing, rather than tiling microarrays as previously reported [40]. We identified 2279 binding sites that were highly enriched (log2 fold change > 3, false discovery rate<0. 01%) when comparing Dam-Cic to Dam alone samples (S1 Table). These sites were non-randomly distributed in the genome, and were significantly over-represented ~500 bp 5’ to Transcription Start Sites (TSS; Fig 4A). Cic DamID was also performed on progenitor cells from P. e. infected midguts. After a 24 hours induction of Dam or Dam-Cic transgenes via the esgts system, flies were fed P. e. bacteria for 16 hours. The number of highly enriched (log2 fold change > 3, FDR < 0. 1%) peaks was reduced to 1903. In addition, the fold change of peaks (Dam-Cic vs Dam-alone) after P. e. infection was significantly decreased (Figs 4B–4C and S4A). The frequency of peaks adjacent to TSS was also significantly reduced in the P. e. -infected midgut sample (Fig 4A). We believe that this decrease was due to the change of Cic localization from the nucleus to cytoplasm, which was caused by the activation of EGFR/Ras/MAPK signaling after infection. To further understand how Cic regulates ISC proliferation we performed gene expression profiling using amplified mRNA from FACS-sorted esg+ progenitor cells that expressed cic-RNAi, and controls. As a way to identify potentially direct target genes of Cic, the RNA-Seq and DamID-Seq data sets were cross-compared. Amongst 439 transcriptionally up-regulated genes (>1. 5 fold change, 90% CI) (S2 Table), a large fraction [134 genes, (S3 Table) ] had Cic binding sites as defined by DamID (Fig 4E). We next examined the enrichment of the DamID peaks in the transcriptionally induced genes, ranked by absolute expression change in cic knockdown progenitor cells (see Materials and Methods). Cic binding peaks that were significantly reduced upon P. e. infection (< 2 fold change) were enriched in up-regulated genes from the RNA-Seq dataset (Fig 4F). Hence, the set of genes present in the overlapping set are likely to be direct target genes of Cic. Many cell cycle regulators and genes involved in DNA replication were upregulated in Cic-depleted progenitor cells (Fig 4D). In addition, a large portion of cell cycle control genes that were upregulated upon cic-RNAi, including string (stg, Cdc25) and Cyclin E (CycE), had Cic binding sites (Fig 4D, 4G, and 4H). To further assess the reliability of this approach we examined the occupancy of Cic on its previously characterized direct target gene-aos [39]. Our DamID-Seq data showed that aos contained two Cic binding sites within its enhancer, and that their occupancy was significantly reduced after P. e. infection (S4B Fig). The significant induction of aos transcription was verified both by RNA-Seq and qRT-PCR data from FACS-sorted progenitor cells expressing cic-RNAi (S4C Fig). Having confirmed the reliability of our approach for identifying genes that are repressed by Cic in ISCs, we focused on genes likely to contribute to ISC proliferation. We were interested in stg and CycE because they are transcriptionally induced in proliferating ISCs [42], required for ISC divisions, and also sufficient to induce sustained ISC division when co-overexpresssed [42]. To further test whether Cic regulates the transcription of stg and CycE we measured their normalized expression ratios in gain- or loss-of-function Cic midguts via RT-qPCR (Fig 4I–4K). The stg and CycE mRNAs were significantly increased in Cic-depleted midguts, and decreased in midguts expressing the dominant active CicΔC2. Strong inductions of stg and CycE were also observed in Cic-depleted progenitor cells or ISCs purified using FACS (Fig 4J). Moreover, both the stg and CycE loci had multiple strong Cic-Dam-ID binding peaks containing TGAATG (G/A) A motifs, and binding these peaks were reduced by P. e. infection (Fig 4G and 4H). Consistently, the induction of stg and CycE transcription upon P. e. infection was significantly repressed by CicΔC2 overexpression (Fig 4K). These data support the notion that Cic controls ISC cell cycle progression by directly repressing transcription of stg and CycE via binding sites in their regulatory regions. It has been suggested that Cic might regulate the transcription of certain members in a subfamily of ETS transcription factors [29,31]. Consistent with this, we identified the Drosophila ETS transcription factors pnt and Ets21C as potential Cic direct target genes by both RNA-Seq and DamID-Seq (Figs 5 and S5). These genes contain Cic binding sites, were highly expressed in midgut progenitor cells, and were significantly induced upon infection or cic depletion or mutation. Notably, induction of pnt and Ets21C was detected in FACS-sorted ISCs depleted of Cic (Fig 5B). Moreover, the induction of pnt and Ets21C expression by P. e. infection was suppressed when the dominant active form, CicΔC2 was overexpressed (Fig 5D). Similar effects were observed when Cic was either depleted or overexpressed in whole midgut samples (Figs 5C and S5C). These data suggest that Cic also regulates pnt and Ets21C transcription in Drosophila midgut ISCs, by directly binding to these loci. As in the case of stg and CycE, this regulation appeared to be modulated by P. e. infection, most likely in a MAPK-dependent manner. The HMG box of Human Cic binds to TGAATG (G/A) A octamers in vitro [31]. This motif was also verified as a Cic binding sequence in several Cic target genes in Drosophila embryos and wing discs [39]. Notably, the TGAATG (G/A) A motif was observed in 692/2279 Cic binding sites in our DamID-Seq dataset (p-value = 3. 045475× 10−11). Each of the four Cic target genes discussed above contained more than one TGAATG (G/A) A motifs in its Cic binding sites. Moreover, TGAATGAA motifs found in the pnt locus also mapped to Cic binding sites that we determined from Drosophila embryo ChIP-Seq (Fig 5E). This suggests that Cic may bind to the pnt locus via TGAATGAA octamers, and that the occupancy of Cic at the pnt locus may also be conserved in different Drosophila cell types. To further evaluate this hypothesis we performed electrophoretic mobility shift assay (EMSA). Cic showed specific binding to two DNA fragments from the pnt locus that were identified as prominent in vivo Cic binding peaks by DamID-Seq and ChiP-Seq (Fig 5F and 5G). Importantly, the EMSA interaction was lost when the HMG-box in Cic was mutated, or when the TGAATGAA motifs were mutated. These data strongly support the idea that Cic directly regulates pnt transcription by directly binding to TGAATGAA motif in pnt locus. Pnt is believed to be a downstream effector of EGFR signaling in developing Drosophila eyes [43,44,45]. The pnt locus produces two alternative transcripts that encode two different protein isoforms: PNTP1 and PNTP2 [44]. PNTP1 was proposed to be a constitutive activator of transcription, whereas PNTP2 has a PNT (pointed) domain that was reported to be phosphorylated by MAP kinase in vitro [45]. The mutant protein, PNTP2T151A, which cannot be phosphorylated in vitro, was unable to rescue pnt phenotype in eyes but instead enhanced the mutant phenotype, suggesting that the PNT domain is an auto-inhibitory domain that can be inactivated by MAPK-dependent phosphorylation [45]. Furthermore PNTP2 is thought to induce transcription of PNTP1, which might thereby encode the final nuclear effector of the EGFR pathway in eye discs [43]. In the midgut, we found an interesting interaction between Pnt and Cic: pntP1 and pntP2 were both induced when Cic was depleted, and both decreased when Cic was overexpressed (Figs 5B, 5C, and S5A). The expression of transcripts encoding both isoforms was also increased in P. e. infected guts (Figs 5D and S5B). This raises the possibility that pnt might be an important downstream effector of Cic in controlling ISC proliferation. To test this we over-expressed either pntP1 or pntP2 in progenitor cells using the esgts or Dlts driver systems. After 4 days of transgene induction a dramatic increase in ISC division was evident in response to either pntP1 or pntP2 (Figs 6A–6B, 6I, and S6A–S6B). Conversely, mutant clones that were generated using a pnt null allele (pnt Δ88) [46] did not grow past the 2-cell stage (S6D Fig). Moreover, when we depleted pnt in progenitor cells by expressing a pnt-RNAi that recognizes both isoforms, or generated homozygous pnt null mutant ISCs via MARCM, ISC proliferation after P. e. infection was suppressed (Figs 6C–6D, 6J, and S6E). Next, we investigated the functional significance of the inhibition of pnt expression by Cic. Whereas loss of Cic function induced massive ISC proliferation, inhibiting both isoforms of pnt in this context suppressed this over-proliferation (Figs 6G–6H, 6K and S6F–S6G). Conversely, when we over-expressed either pntP1 or pntP2 in ISCs that also overexpressed CicΔC2, the inhibitory effect of CicΔC2 on proliferation was bypassed and the cells divided (Figs 6F, 6L and S6C). Hence, a significant fraction of the ISC over proliferation caused by Cic knockdown can be attributed to Cic’s effects on pntP1 and pntP2 Interestingly, mutant clones generated using a pntP1 specific mutant allele, pnt Δ33 [45,47], or a pntP2 specific mutant allele, pnt Δ78 [45,47], grew normally. However ISCs mutant for the pnt null allele pnt Δ88 did not expand (S6D Fig). In addition, pnt Δ33 and pnt Δ78 homozygous clones in which cic was depleted by RNAi had similar numbers of cells to cic-depleted control clones (i. e. they overgrew), whereas pnt Δ88 null mutant clones contained significantly fewer cells (S6G Fig). These data not only support our conclusion that pnt is required for ISC proliferation as a target of Cic, but show that PNTP1 and PNTP2 have redundant function in regulating ISC proliferation. Furthermore, pntP2 homozygous mutant ISCs did not appear to have any defect in proliferation upon P. e. infection (S6E Fig). Overall these results indicate that pntP2, the isoform proposed to be activated directly by MAKP phosphorylation [45], is not specifically required in ISC proliferation. Pnt is the Drosophila ortholog of the human ETS2 transcription factor and has a conserved ETS-type DNA binding domain, while Ets21C is the Drosophila ortholog of the human proto-oncogene ERG. In addition to having Cic binding sites, RT-PCR and RNA-Seq data showed that Ets21C was highly induced upon P. e. infection (Figs 5A and S5C). Moreover RNAi mediated depletion experiments indicated that Ets21C was also required for ISC proliferation in response to P. e. infection (Fig 6J). Over-expression of Ets21C caused a strong increase of ISC division (Fig 6E and 6I) suggesting that transcriptional induction of Ets21C could promote ISC proliferation. Furthermore, ectopic expression of Ets21C in progenitor cells could bypass the strong growth-suppressive effect of depleting MEK (Fig 6M). These data indicated that Cic controls ISC proliferation in part by regulating Ets21C transcription. Finally, we tested whether Yan, an inhibitory ETS type transcription factor, reported to be MAPK responsive and to compete with Pointed for binding to common sites on the DNA [45,48,49], had an opposite function in ISCs. Although yan mRNA is expressed in the midgut (Fig 5A), yan depletion from ISCs did not produce a detectable effect (S6F Fig). Two independent yan-RNAi lines were used, both of which were proven to be effective by qRT-PCR (S6H Fig). In summary these observations suggest that EGFR signaling controls ISC growth and division by regulating the activity of Cic, Pnt and Ets21C but not Yan, and that Cic directly represses pntP1, pntP2 and Ets21C in this context.
It is well established that EGFR signaling is essential to drive ISC growth and division in the fly midgut [10,11,12]. However, the precise mechanism via which this signal transduction pathway activates ISCs has remained a matter of inference from experiments with other cell types. Moreover, despite a vast literature on the pathway and ubiquitous coverage in molecular biology textbooks, the mechanisms of action of the pathway downstream of the MAPK are not well understood for any cell type. From this study, we propose a model summarized in Fig 7. Multiple EGFR ligands and Rhomboid proteases are induced in the midgut upon epithelial damage, which results in the activation of the EGFR, RAS, RAF, MEK, and MAPK in ISCs. MAPK phosphorylates Cic in the nucleus, which causes it to dissociate from regulatory sites on its target genes and also translocate to the cytoplasm. This allows the de-repression of target genes, which may then be activated for transcription by factors already present in the ISCs. The critical Cic target genes we identified include the cell cycle regulators stg (Cdc25) and Cyclin E, which in combination are sufficient to drive dormant ISCs through S and M phases, and pnt and Ets21C, ETS-type transcriptional activators that are required and sufficient for ISC activation. Although we found more than 2000 Cic binding sites in the ISC genome, not all of the genes associated with these sites were significantly upregulated, as assayed by RNA-Seq, upon Cic depletion. One possible explanation for this is that Cic binding sites from DamID-Seq were also associated with other types of transcription units (miRNAs, snRNAs, tRNAs, rRNAs, lncRNAs) that were not scored for activation by our RNA-Seq analysis. Indeed a survey of the Cic binding site distributions suggests this (S5 Table). This might classify some binding sites as non-mRNA-associated. However, it is also possible that many Cic target genes may require activating transcription factors that are not expressed in ISCs. Such genes might not be strongly de-repressed in the gut upon Cic depletion. In other Drosophila cells MAPK phosphorylation is thought to directly inactivate the ETS domain repressor Yan, and to directly activate the ETS domain transcriptional activator Pointed P2 (PNTP2) [45,50]. In fact Pnt and Yan have been shown to compete for common DNA binding sites on their target genes [45,48,49]. Thus, previous studies proposed a model of transcriptional control by MAPK based solely on post-translational control of the activity of these ETS factors. However, we found that Pnt and Ets21C were transcriptionally induced by MAPK signaling, and could activate ISCs if overexpressed, and that depleting yan or pntP2 had no detectable proliferation phenotype. In addition, overexpression of PNTP2 was sufficient to trigger ISC proliferation, suggesting either that basal MAPK activity is sufficient for its post-translational activation, or that PNTP2 phosphorylation is not obligatory for activity. Moreover, pntP2 loss of function mutant ISC clones had no deficiency in growth (S6D Fig) even after inducing proliferation by P. e. infection, which increases MAPK signaling (S6E Fig). These observations indicate that the direct MAPK→PNTP2 phospho-activation pathway is not uniquely or specifically required for ISC proliferation. Or results suggest instead that transcriptional activation of pnt and Ets21c via MAPK-dependent loss of Cic-mediated repression is the predominant mode of downstream regulation by MAPK in midgut ISCs. In addition to activating ISCs for division, EGFR signaling activates them for growth. Previous studies showed loss of EGFR signaling prevented ISC growth and division, and that ectopic RasV12 expression could accelerate the growth not only of ISCs but also post-mitotic enteroblasts [11]. Similarly, our study shows that loss of cic caused ISC clones to grow faster than controls, by increasing cell number as well as cell size (Figs 1H and S2C–S2H). For instance, increased size of GFP+ ISCs and EBs was observed when cic-RNAi was induced by the esgts or esgtsF/O systems (Figs 1B, 6G and S2B). Therefore, in our search for Cic target genes we specifically checked probable growth regulatory genes such as Myc, Cyclin D, the Insulin/TOR components InR, PI3K, S6K and Rheb, Hpo pathway components, and the loci encoding rRNA, tRNAs and snRNAs. We found that Cic bound to the InR, Akt1, Rheb, Src42A and Yki loci. However, of these only InR mRNA was significantly upregulated in Cic-depleted progenitor cells (S4 Table). In surveying the non-protein coding genome, we found that Cic had binding sites in many loci encoding tRNA, snRNA, snoRNA and other non-coding RNAs (S5 Table), though not in the 28S rRNA or 5S rRNA genes (S4 Table). Due to the method we used for RNA-Seq library production, our RNA expression profiling experiments could not detect expression of these loci, and so it remains to be tested whether Cic may regulate some of those non-coding RNA’s transcription to control cell growth. It is also possible that Cic controls cell growth regulatory target genes indirectly, for instance via its targets Ets21C and Pnt, which are also strong growth promoters in the midgut (Figs 6A–6B, 6E and S6A–S6B). But given that no conclusive model can be drawn from our data regarding how Cic restrains growth, one must consider the possibility that ERK signaling stimulates cell growth via non-transcriptional mechanisms, as proposed by several studies [51,52,53,54]. The critical role of Cic as a negative regulator of cell proliferation in the fly midgut is consistent with its tumor suppressor function in mammalian cancer development (S1 Fig). Also consistent with our findings are the observations that the ETS transcription factors ETV1 and ETV5 are upregulated in sarcomas that express CIC-DUX, an oncogenic fusion protein that functions as a transcriptional activator [31], and that knockdown of CIC induces the transcription of ETV1, ETV4 and ETV5 in melanoma cells [29]. Moreover the transcriptional regulation by ETS transcription factors is important in human cancer development (S7 Fig). Their expression is induced in many tumors and cancer cell lines. For example, ERG, ETV1, and ETV4 can be upregulated in prostrate cancers [55], and ETV1 is upregulated in post gastrointestinal stromal tumors [56] and in more than 40% of melanomas [57]. In addition, the mRNA expression of these ETS genes was correlated with ERK activity in melanoma and colon cancer cell lines with activating mutations in BRAF (V600E), such that their expression decreased upon MEK inhibitor treatment [58]. Furthermore, overexpression of the oncogenic ETS proteins ERG or ETV1 in normal prostate cells can activate a Ras/MAPK-dependent gene expression program in the absence of ERK activation [59]. These cancer studies imply that there is an unknown factor that links Ras/Mapk activity to the expression of ETS factors, and that some of the human ETS factors might act without MAPK phosphorylation, as does Drosophila PntP1. Combining our knowledge of Cic with what was previously known about CIC in tumor development, we propose that CIC is the unknown factor that regulates ETS transcription factors in Ras/MAKP-activated human tumors. In summary, our study has elucidated a mechanism wherein Cic controls the expression of the cell cycle regulators stg (Cdc25) and Cyclin E, along with the Ets transcription factor Pnt, and perhaps also Ets21C, by directly binding to regulatory sites in their promoters and introns. Using genetic tests we show that these interactions are meaningful for regulating stem cell proliferation. Therefore, we suggest that human CIC may also lead to the transcriptional induction of cell cycle genes and ETS transcription factors in RAS/MAPK activated- or loss-of-function-CIC tumors such as brain or colorectal cancers.
esgts: esg-Gal4/Cyo; tubGal80ts UAS-GFP/TM6B [60] esgts F/O: esg-Gal4 tubGal80ts UAS-GFP/Cyo; UASflp>CD2>Gal4/TM6B [8] Tubts: tub-Gal4; tubGal80ts/TM3, ser [61] (provided from Valeria Cavaliere lab) esgts; Su (H) -Gal80: esg-Gal4-UAS-2XEYFP; Su (H) GBE- Gal80, tub-Gal80ts (Gift from Steven Hou’s lab) UAS-λTOP/FM7 [37] UAS-RASv12s35 [38] UAS-Ras RNAi [11] UAS-Egfr RNAi [11] UAS-cic-RNAi/Cyo (VDRC KK103805) UAS-cic-RNAi/Cyo (VDRC KK103012) UAS-pnt. P1 (Bloomington Drosophila Stock Center 869) UAS-pnt. P2 (Bloomington Drosophila Stock Center 399) UAS-pnt-RNAi (Bloomington Drosophila Stock Center 31936) UAS-pnt-RNAi (Bloomington Drosophila Stock Center 35808) UAS-yan-RNAi (Bloomington Drosophila Stock Center 26759) UAS-yan-RNAi (Bloomington Drosophila Stock Center 34909) UAS-yan-RNAi (Bloomington Drosophila Stock Center 35404) UAS-Ets21C-RNAi (VDRC KK103211) FRT82B cicfetu6 / TM3, Sb, Se (gift from Jimenez lab, Barcelona) w; cicfetT6 /TM3, Ser (gift from Nilson lab, Canada) w; cicfetE11 / TM6b (gift from Nilson lab, Canada) w; +; UAS-cic-HA w; UAS-cic-HA; + w; +; UAS-cic ΔC2-HA w; UAS-cic ΔC2-HA; + FRT82B pnt Δ33 [45,47] (gift from Joseph Bateman lab, Wolfson Centre for Age-Related Diseases) FRT82B pnt Δ78 [45,47] (gift from Joseph Bateman lab, Wolfson Centre for Age-Related Diseases) FRT82B pnt Δ88[45,47] (gift from Joseph Bateman lab, Wolfson Centre for Age-Related Diseases) The cic ΔC2 was amplified from the pCasper4—cic ΔC2 plasmid. The cic or cic ΔC2 cDNAs were inserted into pUASg-attB-HA [62] vector and used to generate transgenic flies. To generate UAS-cicDam transgenic flies, Cic was amplified from a cDNA library prepared from midgut. This cic cDNA was inserted into the pUASTattB-LT3-NDam plasmid (from Andrea brand lab), and transgenics were produced. Ectopic expression of transgenes in the adult midgut was achieved using the temperature sensitive inducible UAS-Gal4 system [63], TARGET. Crosses were set up and maintained at 18°C, the permissive temperature. 3–7 day old flies were shifted to 29°C for different times as indicated. Gut infections were performed by feeding flies live P. e. in 5% sucrose on Whatman filter paper and yeast paste at 29°C. The MARCM system was used to generate ISC clones. In order to reduce heat shock dependent stress, the clones were induced by heat shocking 3–5 days old flies at 34°C for 20 minutes. The heat shocked flies were then kept at 25°C. Clone size was measured after 10,20,30 days of clone induction. The size of the clones was quantified by Fiji software measuring GFP+ area from z-projected confocal microscopy images. Female adult flies were dissected in 1×PBS. Midguts were fixed in 1×PBS with 4% paraformaldehyde for 30 minutes at room temperature. Samples were washed in PBS with 0. 1% X-100 (PBST) for 3x10 minutes each. Then the tissues were blocked in PBS with 0. 1% X-100,2. 5%BSA, 10% NGS for at least 30 min at room temperature. All samples were incubated with primary antibody overnight at the following dilutions: rat anti-HA (1: 200; Roche), guinea pig anti-Cic (1: 1000, generated by author), rabbit anti-PH3 (1: 1000, Millipore). After washing 3 times 10 minutes each in PBST, samples were incubated with secondary antibodies for at least 2 hours at room temperature at a dilution of 1: 1000. DNA was visualized with DAPI (0. 1mg/ml, Sigma), diluted 1: 200. Images of Figs 1A–1B and 2E–2H were acquired by Delta vision microscope and the rest of the fluorescence images were taken by Leica SP5 confocal microscope. Images were then processed using Fiiji and Adobe Photoshop software. RNA was extracted from 10–12 female midguts using the RNAeasy kit (QIAGEN). RNA isolation from sorted cells was performed as previously described [64] and 100ng RNA (non-amplifed) used for reverse transcription. cDNA was synthesized by QuantiTect reverse transcription kit (QIAGEN). RT-qPCR was performed on a Light Cycler 480 II using SYBR Green I (Roche). Experiments were performed in biological triplicate. Relative fold differences in expression level of target genes were calculated as ratios to the mean of the reference genes rp49 [65] and tubulin [23]. Primer sequences are given in Supplementary Material and Methods. RNA isolation and amplification from sorted cells was performed as previously described [64]. Four independent biological replicates were used for sequencing. Raw reads were checked for quality using Fastqc and subsequently aligned using Tophat2, version 2. 0. 9, against the Flybase genome version 6. Mapped reads were counted using HTSeq-count version 0. 5. 4p5 [66] with mode „union“. Genes showing a cpm value below 1 in four samples per treatment were considered as poorly expressed and filtered out before conducting differential expression analysis using edgeR, version 3. 2. 4 [67]. Since our replicates were generated independently, we used a paired design and corrected the resulting p-values by the Benjamini-Hochberg method [68]. Subsequently, genes with a fold change of 1. 5 and an adjusted p-value lower than 0. 1 were considered as significantly deregulated. Rp49 –Forward: TCGATATGCTAAGCTGTC Rp49 –Reverse: GGCATCAGATACTGTCCCTTG β-tubulin-Forward: ACATCCCGCCCCGTGGTC β-tubulin-Reverse: AGAAAGCCTTGCGCCTGAACATAG pnt-Forward: ACGCCCTATGATGCTCAATC pnt-Reverse: TATCCAGACCCAAGGTGCTC pntP1-Forward: CGACTGCGAACAATCTGGT pntP1-Reverse: TTGCTGGTGTTGTAGCCTGT pntP2-Forward: TTAGCCAATTGAACGGCATC pntP2-Reverse: GCACAGATCCTTGCATCCAT Ets21C-Forward: CCGGGCACTCAGGTACTACT Ets21C-Reverse: CATACTGGAGGCCGGATCT aos-Forward: AGAACCCATGGCTTACATGC aos-Reverse: CGTCGCGGGTGTTAAGTTAC yan-Forward: CTGCTGGTCATCGTGCTTAG yan-Reverse: GACCTCAGTGTGAGCAGCAA stg-Forward: CAGCATGGATTGCAATATCAGTA stg-Reverse: CAACGTCGTCGTCGTAGAAC CycE-Forward: ACAAATTTGGCCTGGGACTA CycE-Reverse: GGCCATAAGCACTTCGTC
Studies suggest that epidermal growth factor receptor (EGFR) signaling activation is a causal driver of many stem cell-derived epithelial cancers, including colorectal cancer. As in the human intestine, epithelial renewal in Drosophila intestine is orchestrated by intestinal stem cells (ISCs). EGFR signaling also plays an important role in regulating ISC proliferation in flies. However, the mechanism by which EGFR/Ras/MAPK signaling promotes ISC proliferation is poorly understood. Here we demonstrate that the transcriptional repressor, Capicua (Cic), mediates these functions of EGFR signaling. We found that the critical role of Cic as a negative regulator of cell proliferation in the fly midgut is consistent with its tumor suppressor function in mammalian cancer development. The direct target genes of Cic were identified by ISC-specific mRNA expression profiling and DNA binding mapping (DamID) method. Cic represses cell proliferation via regulating string (stg), Cyclin E (CycE), and the ETS domain transcription factors Ets21C and pointed (pnt). Using genetic tests we show that these interactions are meaningful for regulating stem cell proliferation. Combining our knowledge of Cic with what was previously known about CIC in tumor development, we propose that human CIC may regulate Ets transcription factors and cell cycle genes in Ras/MAKP-activated tumors.
EGFR/Ras Signaling Controls Drosophila Intestinal Stem Cell Proliferation via Capicua-Regulated Genes