- Email: [email protected]
Setting the Stage for S Phase
Molecular Cell
Previews Setting the Stage for S Phase Angus C. Wilson1,* 1
Department of Microbiology and NYU Cancer Institute, New York University School of Medicine, 550 First Avenue, New York, NY 10016, USA *Correspondence: [email protected] DOI 10.1016/j.molcel.2007.07.001
In a recent issue of Molecular Cell, Tyagi et al. (2007) show that E2F1, a positive regulator of S phase entry, recruits cofactor HCF-1 and associated hSet1/MLL histone H3 lysine 4 methyltransferase complex, facilitating the activation of genes required for proliferation. E2F transcription factors control the expression of numerous genes involved in the G1/S cell-cycle transition, initiation of DNA synthesis, and mitosis and also link DNA-damage recognition pathways to key cell-cycle checkpoints (reviewed in Cam and Dynlacht [2003]). Individual E2F proteins can be classified as activators (E2F1, E2F2, and E2F3a) or repressors (E2F3b, E2F4, and E2F5) and interact with members of the retinoblastoma (Rb) pocket protein family. In serumarrested cells, the majority of E2Fresponsive promoters are occupied by E2F4 together with the p130 pocket protein and a histone deacetylase (HDAC)-containing corepressor complex (Figure 1A). As cells reenter G1 phase of the cell cycle, E2F4 is replaced by E2F1, E2F2, or E2F3a. Subsequent release of the pocket protein through the action of a cyclindependent kinase leads to gene activation and initiation of S phase. This switch is accompanied by the appearance of activating marks including histone H3 and H4 acetylation and methylation of histone H3 lysine 4 (H3K4). Regulation of E2F activity by pocket proteins is fairly well understood, but less is known about mechanisms for activation of E2F target genes. A study by Tyagi et al. reported in a recent issue of Molecular Cell shines light on this more dimly lit corner of an important field. The new work builds on reports that noticed a short sequence (D/EHxY, where x can be any residue) known as the HCF-binding motif (HBM), in E2F1 and E2F4 (Luciano and Wilson, 2003, Knez et al., 2006). Pull-down and mutagenesis experiments firmly established the interac-
tion between host cell factor-1 (HCF-1) and both E2F proteins (Knez et al., 2006). The prototype HBM is found in the herpes simplex virus transactivator VP16, which uses HCF-1 to assemble an activator complex on viral immediate-early gene promoters (reviewed in Wysocka and Herr [2003]). Similar HBM sequences occur in other DNAbinding proteins, coactivators, and corepressors, and HCF-1 has been detected in multiple complexes with histone modifying activity. One wellcharacterized example consists of HCF-1 (or its relative HCF-2), a Set domain-containing protein (hSet1 or MLL), and the three accessory subunits Ash2, WDR5, and RBP5 (Wysocka et al., 2003). Set domains catalyze dimethylation and trimethylation of H3K4, and the promoter regions of active genes are often enriched for these modifications. From the outset, the idea that HCF-1 serves as an E2F cofactor is very appealing. It has been known for more than a decade that loss of HCF-1 function causes a reversible cellcycle arrest in G1 phase and, more recently, that the arrest can be relieved by expression of adenovirus E1A or simian virus 40 large T antigen, viral oncoproteins that counter the repressive function of pocket proteins (Reilly et al., 2002). Using nuclear extracts and chromatin from synchronized cells, Tyagi and colleagues have made further strides in fleshing out the story. They begin by showing that HCF-1 interacts with E2F1, E2F3a, and E2F4, each of which contains the HBM. Appropriately, an interaction was not seen with E2F2, which lacks a candidate sequence. The E2F-HCF
176 Molecular Cell 27, July 20, 2007 ª2007 Elsevier Inc.
interaction is regulated in a cellcycle-dependent manner even though the overall abundance of HCF-1 does not change. Binding of HCF-1 to E2F1 is first detected in late G1 phase, increasing at the G1/S boundary and remaining high as cells progress into S phase. Association with E2F4 shows a reciprocal profile consistent with its pattern of promoter occupancy and role as an antagonist of gene activation and suggests that HCF-1 participates in both the activation and repression of E2F-responsive genes. Next, chromatin immunoprecipitation (ChIP) was used to show that HCF-1 is indeed present at the promoters of E2F-responsive genes, with maximal levels in early G1, coincident with occupancy by E2F4, and then again at the G1/S transition when E2F1 is predominant. Recruitment of HCF-containing activator complexes represented by hSet1, MLL, and WDR5 tracked closely with E2F1 and with elevated levels of H3K4 trimethylation. Depletion of HCF-1 using siRNA did not substantially alter occupancy by E2F1 but significantly decreased recruitment of the HMT complex, with concordant decreases in H3K4 trimethylation and mRNA synthesis. Collectively, these data suggest that the stage-specific recruitment of the H3K4 methyltransferase complex to E2F-responsive genes is largely dependent on HCF-1. The role of HCF-1 in E2F4 repression is less clear. HCF-1 is known to associates with a repressor complex that includes mSin3A/B and HDAC1/2 (Wysocka et al., 2003), and in keeping, mSin3A can be detected after sequential immunoprecipitation of
Molecular Cell
Previews
Figure 1. HCF-1 Facilitates Recruitment of Coactivator and Corepressor Complexes in a Temporal Sequence to E2F-Regulated Genes (A) As cells progress from early G1 into S phase, promoters of E2F-regulated genes are sequentially occupied by E2F4 (or E2F5) and E2F1 (or E2F2 and E2F3). Recruitment of hSet1 and MLL histone methyltransferases (HMT) to E2F1 requires HCF-1 and leads to H3K4 histone methylation and gene transcription. (B) The location of the HCF-binding motif (HBM) differs in activator E2F1 compared to repressor E2F4.
HCF-1 followed by E2F4 from early G1 extracts (Tyagi et al., 2007). Recruitment of an mSin3/HDAC complex via HCF-1 could account for the hypoacetylated state of E2F-regulated genes in early G1; however, other mechanisms are probably in play. It is known that recruitment of HDAC1 to a subset of promoters is dependent on p130 (Rayman et al., 2002), and there is no evidence that inactivation of HCF-1 causes derepression of E2F genes in noncycling cells. The notion that the same adaptor might be used to reverse the active state at the end of S phase is intriguing and raises questions of selectivity. What determines the recruitment of a given HCF-containing complex to a particular E2F protein at the appropriate time? It is striking that HCF-1 binding parallels occupancy of either E2F1 or E2F4 on DNA, hinting that HCF-containing complexes preferentially recognize the DNA-bound E2Fs and that proximity to DNA or promoter context is important. It is certainly
clear that pocket protein release is not the signal for HCF-1 binding because sequential immunoprecipitation from G1/S extracts detects E2F1 in a complex with both HCF-1 and pRB. Could it be that an extended complex is built first and then activated by pocket protein phosphorylation? Selectivity may also relate to HBM position. The basic arrangement of functional domains in E2F1 and E2F4 is similar, and yet the HBMs are in completely different positions (Figure 1B). In E2F1 and E2F3a, the motif lies toward the N terminus, whereas in E2F4 it overlaps with the pocket protein-binding site in the C terminus. Steric hindrance or protein conformation could influence the choice of HCF-1containing complex or direct the assembly of a complex de novo. In terms of preference, E2F1 resembles the archetypal activator VP16, which selects the HCF-containing hSet1/MLL complex, leading to elevated H3K4 methylation on the viral IE promoters (Huang et al., 2006; Wysocka et al., 2003).
This elegant study provides an attractive mechanism for the establishment of activated chromatin on E2F-responsive genes at the G1/S transition and helps explain why cells lacking functional HCF-1 arrest. Almost certainly the use of an HCF adaptor to couple E2F proteins to various histone-modifying complexes is not limited to mammalian cells. The genomes of all multicellular animals encode one or two HCF-like proteins. In Drosophila, there are two E2F proteins and it is satisfying to see that both contain the requisite HBM sequence and can be coimmunoprecipitated with an antibody to dHCF (Tyagi et al., 2007). Thus, equivalent mechanisms are likely to be at work in invertebrates, and indeed dHCF, Ash2, and H3K4 trimethylation are found together on Drosophila polytene chromosomes (Beltran et al., 2007). It seems a safe bet that a subset of these highlighted loci correspond to dE2Fresponsive genes.
REFERENCES Beltran, S., Angulo, M., Pignatelli, M., Serras, F., and Corominas, M. (2007). Genome Biol. 8, R67. Cam, H., and Dynlacht, B.D. (2003). Cancer Cell 3, 311–316. Huang, J., Kent, J.R., Placek, B., Whelan, K.A., Hollow, C.M., Zeng, P.Y., Fraser, N.W., and Berger, S.L. (2006). J. Virol. 80, 5740–5746. Knez, J., Piluso, D., Bilan, P., and Capone, J.P. (2006). Mol. Cell. Biochem. 288, 79–90. Luciano, R.L., and Wilson, A.C. (2003). J. Biol. Chem. 278, 51116–51124. Rayman, J.B., Takahashi, Y., Indjeian, V.B., Dannenberg, J.H., Catchpole, S., Watson, R.J., te Riele, H., and Dynlacht, B.D. (2002). Genes Dev. 16, 933–947. Reilly, P.T., Wysocka, J., and Herr, W. (2002). Mol. Cell. Biol. 22, 6767–6778. Tyagi, S., Chabes, A.L., Wysocka, J., and Herr, W. (2007). Mol. Cell 27, 107–119. Wysocka, J., and Herr, W. (2003). Trends Biochem. Sci. 28, 284–304. Wysocka, J., Myers, M.P., Laherty, C.D., Eisenman, R.N., and Herr, W. (2003). Genes Dev. 17, 896–911.
Molecular Cell 27, July 20, 2007 ª2007 Elsevier Inc. 177
Previews Setting the Stage for S Phase Angus C. Wilson1,* 1
Department of Microbiology and NYU Cancer Institute, New York University School of Medicine, 550 First Avenue, New York, NY 10016, USA *Correspondence: [email protected] DOI 10.1016/j.molcel.2007.07.001
In a recent issue of Molecular Cell, Tyagi et al. (2007) show that E2F1, a positive regulator of S phase entry, recruits cofactor HCF-1 and associated hSet1/MLL histone H3 lysine 4 methyltransferase complex, facilitating the activation of genes required for proliferation. E2F transcription factors control the expression of numerous genes involved in the G1/S cell-cycle transition, initiation of DNA synthesis, and mitosis and also link DNA-damage recognition pathways to key cell-cycle checkpoints (reviewed in Cam and Dynlacht [2003]). Individual E2F proteins can be classified as activators (E2F1, E2F2, and E2F3a) or repressors (E2F3b, E2F4, and E2F5) and interact with members of the retinoblastoma (Rb) pocket protein family. In serumarrested cells, the majority of E2Fresponsive promoters are occupied by E2F4 together with the p130 pocket protein and a histone deacetylase (HDAC)-containing corepressor complex (Figure 1A). As cells reenter G1 phase of the cell cycle, E2F4 is replaced by E2F1, E2F2, or E2F3a. Subsequent release of the pocket protein through the action of a cyclindependent kinase leads to gene activation and initiation of S phase. This switch is accompanied by the appearance of activating marks including histone H3 and H4 acetylation and methylation of histone H3 lysine 4 (H3K4). Regulation of E2F activity by pocket proteins is fairly well understood, but less is known about mechanisms for activation of E2F target genes. A study by Tyagi et al. reported in a recent issue of Molecular Cell shines light on this more dimly lit corner of an important field. The new work builds on reports that noticed a short sequence (D/EHxY, where x can be any residue) known as the HCF-binding motif (HBM), in E2F1 and E2F4 (Luciano and Wilson, 2003, Knez et al., 2006). Pull-down and mutagenesis experiments firmly established the interac-
tion between host cell factor-1 (HCF-1) and both E2F proteins (Knez et al., 2006). The prototype HBM is found in the herpes simplex virus transactivator VP16, which uses HCF-1 to assemble an activator complex on viral immediate-early gene promoters (reviewed in Wysocka and Herr [2003]). Similar HBM sequences occur in other DNAbinding proteins, coactivators, and corepressors, and HCF-1 has been detected in multiple complexes with histone modifying activity. One wellcharacterized example consists of HCF-1 (or its relative HCF-2), a Set domain-containing protein (hSet1 or MLL), and the three accessory subunits Ash2, WDR5, and RBP5 (Wysocka et al., 2003). Set domains catalyze dimethylation and trimethylation of H3K4, and the promoter regions of active genes are often enriched for these modifications. From the outset, the idea that HCF-1 serves as an E2F cofactor is very appealing. It has been known for more than a decade that loss of HCF-1 function causes a reversible cellcycle arrest in G1 phase and, more recently, that the arrest can be relieved by expression of adenovirus E1A or simian virus 40 large T antigen, viral oncoproteins that counter the repressive function of pocket proteins (Reilly et al., 2002). Using nuclear extracts and chromatin from synchronized cells, Tyagi and colleagues have made further strides in fleshing out the story. They begin by showing that HCF-1 interacts with E2F1, E2F3a, and E2F4, each of which contains the HBM. Appropriately, an interaction was not seen with E2F2, which lacks a candidate sequence. The E2F-HCF
176 Molecular Cell 27, July 20, 2007 ª2007 Elsevier Inc.
interaction is regulated in a cellcycle-dependent manner even though the overall abundance of HCF-1 does not change. Binding of HCF-1 to E2F1 is first detected in late G1 phase, increasing at the G1/S boundary and remaining high as cells progress into S phase. Association with E2F4 shows a reciprocal profile consistent with its pattern of promoter occupancy and role as an antagonist of gene activation and suggests that HCF-1 participates in both the activation and repression of E2F-responsive genes. Next, chromatin immunoprecipitation (ChIP) was used to show that HCF-1 is indeed present at the promoters of E2F-responsive genes, with maximal levels in early G1, coincident with occupancy by E2F4, and then again at the G1/S transition when E2F1 is predominant. Recruitment of HCF-containing activator complexes represented by hSet1, MLL, and WDR5 tracked closely with E2F1 and with elevated levels of H3K4 trimethylation. Depletion of HCF-1 using siRNA did not substantially alter occupancy by E2F1 but significantly decreased recruitment of the HMT complex, with concordant decreases in H3K4 trimethylation and mRNA synthesis. Collectively, these data suggest that the stage-specific recruitment of the H3K4 methyltransferase complex to E2F-responsive genes is largely dependent on HCF-1. The role of HCF-1 in E2F4 repression is less clear. HCF-1 is known to associates with a repressor complex that includes mSin3A/B and HDAC1/2 (Wysocka et al., 2003), and in keeping, mSin3A can be detected after sequential immunoprecipitation of
Molecular Cell
Previews
Figure 1. HCF-1 Facilitates Recruitment of Coactivator and Corepressor Complexes in a Temporal Sequence to E2F-Regulated Genes (A) As cells progress from early G1 into S phase, promoters of E2F-regulated genes are sequentially occupied by E2F4 (or E2F5) and E2F1 (or E2F2 and E2F3). Recruitment of hSet1 and MLL histone methyltransferases (HMT) to E2F1 requires HCF-1 and leads to H3K4 histone methylation and gene transcription. (B) The location of the HCF-binding motif (HBM) differs in activator E2F1 compared to repressor E2F4.
HCF-1 followed by E2F4 from early G1 extracts (Tyagi et al., 2007). Recruitment of an mSin3/HDAC complex via HCF-1 could account for the hypoacetylated state of E2F-regulated genes in early G1; however, other mechanisms are probably in play. It is known that recruitment of HDAC1 to a subset of promoters is dependent on p130 (Rayman et al., 2002), and there is no evidence that inactivation of HCF-1 causes derepression of E2F genes in noncycling cells. The notion that the same adaptor might be used to reverse the active state at the end of S phase is intriguing and raises questions of selectivity. What determines the recruitment of a given HCF-containing complex to a particular E2F protein at the appropriate time? It is striking that HCF-1 binding parallels occupancy of either E2F1 or E2F4 on DNA, hinting that HCF-containing complexes preferentially recognize the DNA-bound E2Fs and that proximity to DNA or promoter context is important. It is certainly
clear that pocket protein release is not the signal for HCF-1 binding because sequential immunoprecipitation from G1/S extracts detects E2F1 in a complex with both HCF-1 and pRB. Could it be that an extended complex is built first and then activated by pocket protein phosphorylation? Selectivity may also relate to HBM position. The basic arrangement of functional domains in E2F1 and E2F4 is similar, and yet the HBMs are in completely different positions (Figure 1B). In E2F1 and E2F3a, the motif lies toward the N terminus, whereas in E2F4 it overlaps with the pocket protein-binding site in the C terminus. Steric hindrance or protein conformation could influence the choice of HCF-1containing complex or direct the assembly of a complex de novo. In terms of preference, E2F1 resembles the archetypal activator VP16, which selects the HCF-containing hSet1/MLL complex, leading to elevated H3K4 methylation on the viral IE promoters (Huang et al., 2006; Wysocka et al., 2003).
This elegant study provides an attractive mechanism for the establishment of activated chromatin on E2F-responsive genes at the G1/S transition and helps explain why cells lacking functional HCF-1 arrest. Almost certainly the use of an HCF adaptor to couple E2F proteins to various histone-modifying complexes is not limited to mammalian cells. The genomes of all multicellular animals encode one or two HCF-like proteins. In Drosophila, there are two E2F proteins and it is satisfying to see that both contain the requisite HBM sequence and can be coimmunoprecipitated with an antibody to dHCF (Tyagi et al., 2007). Thus, equivalent mechanisms are likely to be at work in invertebrates, and indeed dHCF, Ash2, and H3K4 trimethylation are found together on Drosophila polytene chromosomes (Beltran et al., 2007). It seems a safe bet that a subset of these highlighted loci correspond to dE2Fresponsive genes.
REFERENCES Beltran, S., Angulo, M., Pignatelli, M., Serras, F., and Corominas, M. (2007). Genome Biol. 8, R67. Cam, H., and Dynlacht, B.D. (2003). Cancer Cell 3, 311–316. Huang, J., Kent, J.R., Placek, B., Whelan, K.A., Hollow, C.M., Zeng, P.Y., Fraser, N.W., and Berger, S.L. (2006). J. Virol. 80, 5740–5746. Knez, J., Piluso, D., Bilan, P., and Capone, J.P. (2006). Mol. Cell. Biochem. 288, 79–90. Luciano, R.L., and Wilson, A.C. (2003). J. Biol. Chem. 278, 51116–51124. Rayman, J.B., Takahashi, Y., Indjeian, V.B., Dannenberg, J.H., Catchpole, S., Watson, R.J., te Riele, H., and Dynlacht, B.D. (2002). Genes Dev. 16, 933–947. Reilly, P.T., Wysocka, J., and Herr, W. (2002). Mol. Cell. Biol. 22, 6767–6778. Tyagi, S., Chabes, A.L., Wysocka, J., and Herr, W. (2007). Mol. Cell 27, 107–119. Wysocka, J., and Herr, W. (2003). Trends Biochem. Sci. 28, 284–304. Wysocka, J., Myers, M.P., Laherty, C.D., Eisenman, R.N., and Herr, W. (2003). Genes Dev. 17, 896–911.
Molecular Cell 27, July 20, 2007 ª2007 Elsevier Inc. 177