Arginine methylation-dependent reader-writer interplay governs growth control by E2F-1

E2F-1 undergoes arginine methylation by PRMT1 and PRMT5

PRMT5 symmetrically methylates E2F-1 on arginine (R) residues 111 and 113 (Cho et al., 2012). We evaluated whether asymmetric arginine methylation also occured on E2F-1 by immunoprecipitating endogenous E2F-1 followed by immunoblotting with anti-asymmetric and anti-symmetric methylation-specific antibodies; both modifications were detected on E2F-1, and absent when cells were treated with E2F-1 siRNA before the immunoprecipitation step (Figure 1a). Moreover, when the E2F-1 immunocomplex was assessed for symmetrically acting PRMTs, an interaction was detected between E2F-1 and PRMT1 in addition to PRMT5 (Figure 1b). The asymmetric arginine methylation mark on E2F-1 was dependent on PRMT1 because it was no longer detectable on E2F-1 immunoprecipitated from PRMT1 siRNA treated cells (Figure 1c); a similar effect was apparent on the symmetrical methylation modification of E2F-1 in cells treated with PRMT5 siRNA (Figure 1c). Furthermore, PRMT1 was able to methylate E2F-1 in vitro, as PRMT1 immunopurified from cells (with undetectable PRMT5) was capable of in vitro methylating E2F-1 (SI Figure 1a). Ectopic PRMT5 behaved in a similar fashion, as immunopurified PRMT5 (with undetectable PRMT1) could methylate E2F-1 in vitro (SI Figure 1b). These results were recapitulated with recombinant proteins, as both GST-PRMT1 and GST-PRMT5 methylated E2F-1 in vitro (not shown). Thus, the interaction between E2F-1 and PRMT1 and PRMT5 reflected the presence of the relevant arginine methylation mark on E2F-1 in cells, namely asymmetric (by PRMT1) or symmetric (by PRMT5) respectively.

We investigated the residues in E2F-1 methylated by PRMT1 using a panel of single residue substitutions expressed as GST-E2F-1 fusion proteins (R to K; Figure 1d). Each purified GST-E2F-1 fusion protein was incubated with either ectopic PRMT1 or PRMT5 and the capacity as methylation substrates measured. As expected, wild-type GST-E2F-1 could be in vitro methylated by PRMT1 and PRMT5 (Figure 1e and f). Substitution at either R111 or R113 (single or in combination) failed to alter the methylation of E2F-1 by PRMT1, in contrast to mutating R109, as the R109K mutant was poorly methylated by PRMT1 (Figure 1e and f). In contrast, PRMT5 efficiently methylated R109K but failed to methylate R111K and R113K (Figure 1e and f). Expressing the E2F-1 mutants in transfected cells indicated that asymmetric arginine methylation (ADMA) occurred on R111K and R113K, but not R109K or the triple KKK mutant derivatives (Figure 1g). However, the R109K mutant had readily detectable levels of symmetric methylation (SDMA; Figure 1g). An analysis by tandem mass spectrometry of E2F-1 purified from cells provided further support for dimethylation at R109 (SI Figure 4). These results establish therefore that R109 is the major site of asymmetric arginine methylation on E2F-1 in cells, and identify PRMT1 as the enzyme responsible for the methylation event.

Competition between PRMT1 and PRMT5 for E2F-1 methylation

Since PRMT1 and PRMT5 bind to and methylate a similar region in E2F-1 (Figure 1d), we reasoned that competition between the two enzymes may occur. We tested this idea by immunoprecipitating E2F-1 from cells treated with either PRMT1 or PRMT5 siRNA, and measured the interaction between E2F-1 and the other PRMT enzyme. The interaction between E2F-1 and PRMT5 was enhanced when PRMT1 was depleted and conversly the interaction between PRMT1 and E2F-1 increased in conditions of PRMT5 depletion (Figure 1c). Significantly, the level of E2F-1 methylation coincided with PRMT binding. Thus, enhanced PRMT1 binding under PRMT5 siRNA treatment conditions reflected increased E2F-1 asymmetric arginine methylation and, conversely, enhanced levels of PRMT5 upon PRMT1 siRNA treatment reflected greater symmetric methylation (Figure 1c). Further, ectopic PRMT1 expression in a stable cell line caused an enhanced asymmetric and decreased symmetric mark on E2F-1 (SI Figure 5a).

We next examined whether the methylation mark mediated by each enzyme was able to interfere with methylation by the other enzyme. We used chemically synthesised E2F-1 peptides where individual arginine residues within the R-rich motif were either asymmetrically or symmetrically modified to reflect the methylation mark in cells (Figure 1h). A peptide in which each R residue had been changed to a K residue, KKK, could not be methylated by either PRMT1 or PRMT5 (Figure 1h). PRMT1 methylated the E2F-1 peptide only when R109 was in the unmodified state, since the asymmetrically methylated R109 (R109-ADMA) peptide was poorly methylated (Figure 1h). Similarly, PRMT5 methylated the E2F-1 peptide, but not when the peptide had symmetric methylation at R111 and R113 (R111/R113-SDMA; Figure 1h). Interestingly, R111/R113-SDMA had an impact on R109 methylation by PRMT1, as methylation by PRMT1 of R111/R113-SDMA was reduced when compared to the unmodified E2F-1 peptide and R109-ADMA affected the methylation of R111 and R113 by PRMT5, as PRMT5 methylation of R111/R113-SDMA was also reduced (Figure 1h). These results define a biochemical antagonism between the methylation events on E2F-1 carried out by PRMT1 and PRMT5.

We established that the methylated peptide results were relevant to what happens in cells by studying the properties of the E2F-1 mutant derivatives. The R109K mutant had reduced levels of asymmetric methylation but exhibited higher levels of symmetric methylation compared to wild-type E2F-1 (Figure 1g). Conversely, mutation of R111 or R113 resulted in reduced symmetric methylation, and a correspondingly increased level of asymmetric modification compared to wild-type E2F-1 (Figure 1g). Combined with the methylated peptide results (Figure 1h), these results are consistent with the idea that methylation by PRMT1 or PRMT5 impacts on methylation by the other PRMT enzyme, namely methylated R109 hinders symmetric methylation at R111 and R113, and methylation of R111 and R113 reduces asymmetric methylation at R109.

We were interested to study the role of PRMT1 methylation of E2F-1, and identify any biochemical differences that might occur through the arginine residues targeted by each enzyme. We therefore compared the half-life of R109K to wild-type E2F-1, R111/113K and KKK. The R109K mutant had a much shorter half-life compared to wild-type E2F-1, and an even greater difference in half-life was apparent when R109K was compared to R111/113K (1h compared to 6h; Figure 2 a, b and c). In cells treated with PRMT1 siRNA (Figure 2d), E2F-1 had a shorter half-life than wild-type E2F-1 and an extended half-life was observed in cells depleted of PRMT5 (Figure 2 e, f and g), which is consistent with the stability differences apparent with the R to K mutants. Therefore, opposite biochemical consequences occur upon asymmetric and symmetric arginine methylation of E2F-1, as PRMT1 methylation increases E2F-1 half-life, in contrast to PRMT5 methylation which reduces E2F-1 half-life.

Impact of E2F-1 methylation on chromatin binding and transcription

We further investigated whether there were any differences in the effect of PRMT1 and PRMT5 on E2F-1 promoter binding activity by chromatin immunoprecipitation (ChIP), using a selection of E2F target genes connected with different cell fates (SI Figure 2). PRMT1 depletion led to reduced E2F-1 ChIP activity across the set of genes examined, contrasting with PRMT5 depletion, where enhanced ChIP activity was apparent (SI Figure 2;(Cho et al., 2012)). To establish that the differences in promoter binding did not only result from the altered level and half-life of E2F-1 protein caused by manipulating PRMT1 and PRMT5, we evaluated the properties of individual E2F-1 mutant derivatives under conditions of equivalent protein expression (SI Figure 2). The R109K mutant exhibited reduced ChIP activity relative to wild-type E2F-1, contrasting with mutation of R111 or R113, or both together, where enhanced E2F-1 ChIP activity was evident (SI Figure 2). Under conditions of either combined PRMT1 and PRTM5 siRNA treatment or expression of the KKK mutant, it was R111 and R113 that provided the dominant effect (SI Figure 2). These results highlight the opposing effects of PRMT1 and PRMT5 on E2F-1 chromatin binding activity.

We investigated the influence of PRMT1 and PRMT5 on the transcriptional activity of E2F target genes using a panel of luciferase reporters (SI Figure 3). Depleting PRMT1 with siRNA caused reduced transcriptional activity relative to the control siRNA treated cells (SI Figure 3). In contrast, PRMT5 siRNA resulted in an increase in transcription (SI Figure 3;(Cho et al., 2012)). In conditions of the combined depletion of PRMT1 and PRMT5, enhanced levels of transcription were retained (SI Figure 3). The transcription properties of the E2F-1 mutants were also assessed where, at equivalent levels, the activity of the R109K was compromised compared to wild-type E2F-1 (SI Figure 3). In contrast, mutating either R111, R113 or both residues together resulted in enhanced activity (SI Figure 3). Thus, asymmetric and symmetric arginine methylation have opposite effects on E2F-1 activity.

In order to determine the influence of PRMT1 and PRMT5 on endogenous E2F target genes, we measured RNA levels in siRNA treated cells (Figure 2h and SI Figure 1c). Depleting PRMT1 reduced the expression of a variety of E2F target genes, with the greatest effect apparent on genes such as APAF1, E2F-1 and p73, contrasting with PRMT5 depletion, which caused an increase in the RNA level of the same genes (Figure 2h). The combined depletion of PRMT1 and PRMT5 retained the increased expression profile characteristic of PRMT5 siRNA alone (Figure 2h). Thus, PRMT1 is a positive regulator of some E2F-1 target genes, in contrast to PRMT5 which is a negative regulator. Furthermore, genes connected with apoptosis (p73, APAF1 and E2F-1) are particularly sensitive to control by arginine methylated E2F-1.

Arginine methylation in E2F-1-dependent growth control

We assessed the role of PRMT1 in regulating cell growth in a colony formation assay, where PRMT1 depletion resulted in an increased growth rate compared to the control treatment (Figure 3a and SI Figure 5b and c). The increased growth was dependent on E2F-1, because it was lost when PRMT1 and E2F-1 were depleted together (Figure 3a). This was also apparent when viable cells were quantitated by measuring cellular ATP (as a measure of metabolically active cells), where cell number increased upon PRMT1 depletion relative to the control siRNA treatment, and this effect was again reduced when E2F-1 was co-depleted (Figure 3b). The influence of PRMT1 on cell growth was in sharp contrast to PRMT5, where growth inhibition was apparent again in a fashion dependent upon E2F-1 activity (Figure 3b; (Cho et al., 2012)). In cells depleted of both PRMT1 and PRMT5, growth inhibition was apparent which again was mediated through E2F-1 (Figure 3a and 3b). The growth regulating properties of the E2F-1 mutants were assessed in similar assays. R109K caused less growth inhibition than wild-type E2F-1, which was even more striking when R109K was compared to R111/R113K, where enhanced growth inhibition was evident (Figure 3c). Similar outcomes were evident when cellular ATP was measured, which again verified the significant growth advantage of R109K relative to wild-type E2F-1, and growth inhibition by the R111/113K mutant (Figure 3d).

We continued the analysis of PRMT1 and PRMT5 by measuring apoptosis, specifically the level of cleaved PARP and sub-G1 cells (Figure 3e and f). PRMT1 siRNA reduced the level of apoptotic (sub-G1) cells, contrasting with PRMT5 siRNA where a greater level of apoptosis was evident and both effects of PRMT1 and PRMT5 were dependent upon E2F-1 (Figure 3e). Similarly, opposite effects of PRMT1 and PRMT5 siRNA were apparent on PARP cleavage, where PRMT1 siRNA caused a modest decrease contrasting with PRMT5 siRNA where an increased level of cleaved PARP was apparent (Figure 3f). Altogether, these results establish opposite roles for asymmetric and symmetric methylation marks on E2F-1 in regulating cell growth and apoptosis.

Regulation of arginine methylation under DNA damage condition

E2F-1 is a DNA damage responsive protein, and in DNA damaged cells drives apoptosis (La Thangue, 2003; Polager and Ginsberg, 2008; Stevens et al., 2003). It was of interest to establish the role of PRMT1 and PRMT5 in E2F-1 control under DNA damage conditions. In cells treated with etoposide the level of E2F-1 increased (Figure 4a). Under these DNA damage conditions, PRMT1 siRNA reduced the level of E2F-1 which coincided with decreased PARP cleavage and sub-G1 cells compared to the control siRNA treatment (Figure 4a and b). In contrast, PRMT5 siRNA enhanced the level of E2F-1, which reflected increased PARP cleavage and sub-G1 cells (Figure 4a and b; (Cho et al., 2012)).

Both asymmetric and symmetric arginine methylation modifications occur on E2F-1 in unperturbed cells (Figure 1a and c). In DNA damaged cells (treated with either doxorubicin or etoposide) the enhanced level of E2F-1 coincided with decreased interaction with PRMT5 and a corresponding reduction in symmetric arginine methylation (Figure 4c; SDMA). In contrast, an enhanced interaction with PRMT1 and associated increase in asymmetric arginine methylation occurred (Figure 4c; ADMA), and the greater methylation by PRMT1 and reduced methylation by PRMT5 reflected increased E2F-1 levels and expression of E2F target genes (SI Figure 1d). Thus, arginine methylation of E2F-1 mediated by PRMT1 and PRMT5 is regulated by DNA damage. The competitive binding events between PRMT1 and PRMT5 noted earlier (Figure 1), might contribute to the regulated methylated events.

We then investigated the effect of PRMT1 on E2F-1 under DNA damage conditions in colony formation assays. As expected, fewer colonies were evident in etoposide treated cells compared to unperturbed cells, and inhibition of colony formation was enhanced upon treatment with PRMT5 siRNA (Figure 4d). Remarkably, however, PRMT1 siRNA enhanced colony formation activity and overcame the growth inhibitory activity of etoposide relative to the control siRNA treatment (Figure 4d). The enhanced colony formation apparent upon PRMT1 siRNA treatment and the reduced colony formation upon PRMT5 siRNA co-treatment were dependent on E2F-1 activity (Figure 4e). However, the ability of PRMT1 siRNA to overcome DNA damage induced growth inhibition was lost upon co-depletion with PRMT5 (Figure 4d and e), arguing that the colony formation activity seen under PRMT1 siRNA treatment was mediated through PRMT5 and symmetric methylation of R111 and R113. The enhanced symmetrical arginine methylation and increased PRMT5 binding upon PRMT1 depletion (Figure 1c) is consistent with the observed increase in colony activity under DNA damage from PRMT1 siRNA, as PRMT5-dependent symmetric modification of E2F-1 would be expected to suppress its apoptotic activity.

Arginine methylation and cyclin A binding to E2F-1

The cyclin A/cdk2 binding motif in E2F-1 is located in the region from residue 87 to 95, containing the core consensus cyclin binding motif RRL (Adams et al., 1996). We reasoned that the juxta-position of this motif with the R-rich motif might result in a level of competitive interplay between cyclin A and PRMT1 and/or PRMT5. We tested this idea by treating cells with cyclin A siRNA, and monitored the effect on E2F-1 methylation and interaction with PRMT1 and PRMT5. An interaction was apparent between E2F-1 and cyclin A, and PRMT5 could be detected in the same immunoprecipitated complex (Figure 5a). In cells treated with cyclin A siRNA, increased levels of E2F-1 occurred, with a concomitant increase in the level of PRMT1 and reduced PRMT5 in the E2F-1 immunocomplex (Figure 5a). Furthermore, changes in the level of arginine methylation mirrored these binding events as asymmetric arginine methylation of E2F-1 was higher in cyclin A siRNA treated cells, in contrast to symmetric arginine methylation which was highest in the control siRNA treated cells (Figure 5a). The absence of cyclin A binding to E2F-1 therefore enabled enhanced PRMT1 and reduced PRMT5 interaction, which thus resembles the profile of interaction and methylation events that occur in DNA damaged cells (Figure 4c).

When the interaction between E2F-1 and cyclin A was examined in DNA damaged cells, a decreased interaction with cyclin A was observed compared to undamaged cells (Figure 5b). Moreover, the decreased cyclin A binding in DNA damaged cells coincided with enhanced PRMT1 and reduced PRMT5 binding to E2F-1, together with enhanced asymmetric and reduced symmetric respectively (Figure 5b). Thus, cyclin A binding hinders PRMT1 which favours PRMT5 interacting with E2F-1, and DNA damage causes a decreased interaction with cyclin A, which allows a stronger interaction with PRMT1 with reduced PRMT5 binding.

The increased binding of PRMT1 to E2F-1 under conditions of decreased cyclin A binding and the coincidental increase in asymmetric methylation would, based on the earlier results, be expected to facilitate E2F-1-dependent apoptosis (by virtue of PRMT1 methylation hindering PRMT5 methylation; Figure 1c). We tested this idea using a cyclin A binding defective E2F-1 derivative, in which the cyclin A binding domain had been removed (▵87-95; Figure 5c). The E2F-1▵87-95 mutant exhibited an enhanced interaction with PRMT1 and reduced PRMT5 interaction, and an associated increase in asymmetric but not symmetric methylation (Figure 5c). Significantly, E2F-1▵87-95 reduced colony formation (under normal and DNA damage conditions) more effectively than wild-type E2F-1, similar to the level seen with R111/113K mutant (Figure 5d), and by flow cytometry ▵87-95 displayed increased apoptotic activity relative to wild-type E2F-1 (SI Figure 1e). Thus, cyclin A binding to E2F-1 impacts on the type of arginine methylation mark that occurs on E2F-1, which thereby controls cell cycle progression by E2F-1 activity.

p100-TSN reads E2F-1 arginine methylation

Because the methylation status of R111/R113 impacts on E2F-1-dependent apoptosis, and since the effect of R109 methylation is in part mediated through regulating the methylation status of R111/R113, it was of interest to gather an understanding of proteins that read the symmetrical arginine methylation mark. We prepared biotinylated E2F-1 peptides that were either unmodified or symmetrically modified (SDMA) at R111 and R113, and used the fluorescently labelled streptavidin conjugate to screen the chromatin-associated domain array (CADOR; (Kim et al., 2006; Yang et al., 2010)). CADOR is a protein array platform developed to identify protein domains that bind to modified peptides, and includes the vast majority of reader domains involved with chromatin and transcriptional control (Yang et al., 2010). We identified a single hit in the screen, p100-TSN (TDRD11), which bound only to R111/113 SDMA peptide and not the unmethylated E2F-1 peptide (Figure 6a). p100-TSN is composed of a single extended Tudor domain (TD) and five staphylococcal nuclease (SN)-like domains (Figure 6b) (Shaw et al., 2007). The Tudor domain is a member of the Royal family of protein domains that bind to methyl-lysine and methyl-arginine residues (Chen et al., 2011), and p100-TSN has documented roles in RNA processing and transcriptional regulation (Shaw et al., 2007; Valineva et al., 2005).

We established that the interaction between p100-TSN and E2F-1 was arginine methylation-dependent using in vitro and cell-based approaches. In a peptide binding assay, only R111/113 SDMA and not the unmethylated or R109 ADMA E2F-1 peptide bound to p100-TSN (Figure 6c and SI Figure 1f), confirming the specificity of p100-TSN for symmetric arginine methylation. A kinetic biophysical analysis by biolayer interferometry of the binding between the p100-TSN Tudor domain and R111/113 SDMA peptide determined the dissociation constant (Kd) to be in the order of 12μM (Figure 6d), which is similar to the Kd for other reader domains of arginine methylation marks (Liu et al., 2010); there was no detectable binding to the unmodified peptide. Moreover, the integrity of the Tudor domain in p100-TSN was required for the interaction because the Tudor domain inactivating mutant E770K failed to bind to the R111/113 SDMA peptide (Figure 6c), which is consistent with the role of the Tudor domain in recognising methylated arginine residues (Liu et al., 2010). In both unperturbed and transfected cells, an interaction was apparent between E2F-1 and p100-TSN (Figure 6e, f and h). The interaction was dependent on symmetric arginine methylation because p100-TSN failed to interact with the R111/113K mutant, contrasting with wild-type E2F-1 (Figure 6f), and the interaction between E2F-1 and p100-TSN was enhanced by the ectopic expression of PRMT5 (Figure 6g) and reduced upon PRMT5 siRNA treatment (Figure 6h). Furthermore, the interaction between p100-TSN and E2F-1 affected the half-life of E2F-1. Thus, an extended half-life was apparent in p100-TSN siRNA treated cells, which resembled the effect of PRMT5 siRNA on E2F-1 half-life (Figure 6i, j, k and l). In cells treated with both p100-TSN and PRMT5 siRNA, E2F-1 half-life remained similar to the single siRNA treatments (Figure 6l), suggesting that p100-TSN is involved in mediating the effect of the SDMA mark on E2F-1.

The ability of p100-TSN to bind to the form of E2F-1 that has reduced apoptotic activity (namely R111/113 SDMA) suggested that in cells the interaction could influence cell viability. We tested this idea by depleting p100-TSN with siRNA, which resulted in increased levels of E2F-1 and an associated increase in PARP cleavage (Figure 7a). Significantly, co-depletion of p100-TSN and E2F-1 overcame PARP cleavage (Figure 7a), indicating that the effect of p100-TSN on apoptosis was mediated through E2F-1. A similar relationship between p100-TSN and E2F-1 was evident when sub-G1 cells were measured by flow-cytometry (Figure 7b). Further, in a colony formation assay p100-TSN siRNA decreased colony formation, which was no longer apparent when p100-TSN and E2F-1 were co-depleted (Figure 7c).

To resolve the mechanism involved in greater detail, we addressed if p100-TSN could bind to the promoters of E2F target genes. By ChIP, p100-TSN was detected on E2F-1, Cyclin E, Cdc6 and DHFR genes, contrasting with the APAF-1 and p73 genes, where E2F-1 was not detectable (Figure 7d). The presence of p100-TSN on E2F target genes was dependent on E2F-1, since in E2F-1 siRNA treated cells p100-TSN ChIP activity was lost (Figure 7d). Similarly, p100-TSN ChIP activity was absent in PRMT5 siRNA treated cells (Figure 7d). Furthermore, enhanced p100-TSN ChIP activity was apparent in PRMT1 siRNA treated cells (Figure 7d), which is compatible with the enhanced SDMA mark on E2F-1 upon PRMT1 siRNA treatment (Figure 1c) and recognition of SDMA by p100-TSN (Figure 6c). These results indicate that p100-TSN reads the SDMA mark on E2F-1, and further suggest that the interaction regulates cell viability. This occurs, in part, through p100-TSN regulating E2F-1 stability and targeting a group of E2F responsive genes.

Cell Culture

U2OS and SAOS2 cell lines were obtained from the American Type Culture Collection (ATCC). They were maintained in Dulbecco's Modified Eagle Medium (DMEM) (GIBCO®) supplemented with 10% (v/v) Foetal Calf Serum (FCS) (Biosera) and 1% (v/v) Penicillin-Streptomycin (Pen-Strep) (Gibco®). Stable Tet-On cell lines expressing inducible PRMT5 were prepared as previously described (Jansson et al., 2008). They were maintained in DMEM supplemented with 5% (v/v) Tet-negative FCS, 100μg/ml G418 (Clontech), 0.3% hygromycin (Clontech) and 1% (v/v) Pen-Strep. 1μg/ml doxycline was used to induce PRMT5 expression. They were maintained in DMEM with 10% (v/v) FBS, 1% Pen/Strep (v/v) and 2μg/ml puromycin (Clontech).

Antibodies

p73 antibody was from Abcam. PARP-1 antibody was from BD Pharmingen. Cleaved PARP, PRMT1 and PRMT5 antibody were from Cell Signaling. HA antibody was from Covance. Asymmetric dimethyl-arginine and symmetric dimethyl-arginine antibodies were from Merck Millipore. APAF1, Cdc6, Cyclin E, E2F-1 and GAPDH antibodies were from Santa Cruz. Flag and β-actin antibodies were from Sigma. The p100-TSN antibody was from Bethyl Laboratories. Mouse and rabbit secondary antibodies were from GE Healthcare. Trueblot® secondary antibodies were from eBioscience. HA or Flag antibody-coupled agarose beads were from Sigma.

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Supplementary Materials