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Understanding the function of histone modifications across inducible genes in mammalian cells requires quantitative, comparative analysis of their fate during gene activation and identification of enzymes responsible. We produced high-resolution comparative maps of the distribution and dynamics of H3K4me3, H3K36me3, H3K79me2 and H3K9ac across c-fos and c-jun upon gene induction in murine fibroblasts. In unstimulated cells, continuous turnover of H3K9 acetylation occurs on all K4-trimethylated histone H3 tails; distribution of both modifications coincides across promoter and 5′ part of the coding region. In contrast, K36- and K79-methylated H3 tails, which are not dynamically acetylated, are restricted to the coding regions of these genes. Upon stimulation, transcription-dependent increases in H3K4 and H3K36 trimethylation are seen across coding regions, peaking at 5′ and 3′ ends, respectively. Addressing molecular mechanisms involved, we find that Huntingtin-interacting protein HYPB/Setd2 is responsible for virtually all global and transcription-dependent H3K36 trimethylation, but not H3K36-mono- or dimethylation, in these cells. These studies reveal four distinct layers of histone modification across inducible mammalian genes and show that HYPB/Setd2 is responsible for H3K36 trimethylation throughout the mouse nucleus.
Histone modifications, which include acetylation, phosphorylation, methylation and ubiquitination, play key roles in gene regulation (reviewed in Shilatifard, 2006; Berger, 2007; Bernstein et al, 2007; Kouzarides, 2007; Li et al, 2007). They are studied by chromatin immunoprecipitation (ChIP) to define their location, by genetic and biochemical analyses to identify enzymes involved and by identification of binding proteins or domains to determine their molecular function. ChIP is widely used to map histone modifications across genes and chromosomes, but their quantification and interpretation is contentious (reviewed in Clayton et al, 2006). In particular, high-resolution quantitative comparative studies in synchronised mammalian cells tracking changes in histone modification during transcriptional activation are currently unavailable.
The immediate-early (IE) genes c-fos and c-jun are rapidly induced in mammalian cells via ERK or p38 MAP kinase cascades. These kinases activate the downstream kinase MSK1/2, which phosphorylates histone H3 and HMGN1 (reviewed in Clayton and Mahadevan, 2003). Phosphorylated histone H3 is subject to continuous dynamic acetylation, creating phosphoacetylated H3 (Barratt et al, 1994), which is seen at c-fos and c-jun upon gene induction (Cheung et al, 2000; Clayton et al, 2000). Although they are colocalised at these genes, these two modifications are independently regulated by distinct mechanisms (Thomson et al, 2001). More recently, we reported that all K4-trimethylated H3 in these cells is also subject to extremely dynamic, continuous turnover of acetylation (Hazzalin and Mahadevan, 2005). ChIP with phosphoacetylated (S10phK9ac) or K4me3-specific antibodies and re-ChIP experiments showed that all three modifications can occur on the same nucleosomes (Hazzalin and Mahadevan, 2005). Because K4me3 was continuously detectable, whereas K9ac and S10ph were transient, we favoured a model whereby K4me3 was the key pre-existing modification, which attracts both dynamic acetylation and stimulus-dependent S10 phosphorylation (reviewed in Clayton et al, 2006). Such complex models can only be confirmed by detailed quantitative comparative mapping of these modifications across genes. H3 methylated at lysines 4, 36 or 79 is generally associated with active/permissive chromatin, whereas lysine 9 or 27 methylation is associated with repression (reviewed in Shilatifard, 2006). Within genes, K4me3, K9ac and K14ac occur around start sites of active genes (Liang et al, 2004; Bernstein et al, 2005; Pokholok et al, 2005; Barski et al, 2007; Guenther et al, 2007; Mikkelsen et al, 2007). K4me2 has been reported at start sites of active genes in metazoans (Liang et al, 2004; Schneider et al, 2004), but is spread throughout coding regions in yeast (Pokholok et al, 2005). In contrast, K4me1 is reported towards the 3′ end of active genes (Kouskouti and Talianidis, 2005; Pokholok et al, 2005), as have both K36me2 and me3 (Bannister et al, 2005; Farris et al, 2005; Morillon et al, 2005; Pokholok et al, 2005; Vakoc et al, 2006; Barski et al, 2007; Guenther et al, 2007; Mikkelsen et al, 2007). An exception to this is a transient post-initiation peak of K36me2 and me3 at the yeast MET16 promoter (Morillon et al, 2005). In Drosophila, K79me2 occurs throughout coding regions of active genes (Schübeler et al, 2004). Likewise, in yeast, K79me3 was found in coding regions, but there was no correlation with gene activity (Pokholok et al, 2005). At active mammalian genes, K79me2 and me3 are reported to peak towards the 5′ end of the coding region (Kouskouti and Talianidis, 2005; Vakoc et al, 2006), although another study found K79me2 at both promoter and coding regions of active genes (Im et al, 2003).
Complications to all such studies include quantitation and comparability of ChIP recovery using different antibodies and the possibility of microheterogeneity at any position, whereby distinct histone modifications, deriving from different cells within a population, are detectable at the same position along a gene. An obvious source of microheterogeneity is the transient dynamic nature of some histone modifications, for example, H3 acetylation described above. A second relates to the fact that some histone-modifying enzymes travel with elongating RNA polymerase II (Pol II), producing elongation-dependent variation across coding regions. In yeast, the K4 methyltransferase SET1/COMPASS complex (Briggs et al, 2001; Krogan et al, 2002) coimmunoprecipitates with Pol II phosphorylated at Ser 5 in its CTD (Ng et al, 2003), a form associated with early elongating complexes (Komarnitsky et al, 2000). In contrast, Set2, the yeast K36 methyltransferase (Strahl et al, 2002), associates with Pol II phosphorylated at Ser 2 in its CTD (Li et al, 2002; Krogan et al, 2003; Schaft et al, 2003; Xiao et al, 2003; Kizer et al, 2005), a form of Pol II found increasingly towards the 3′ end of transcribed genes (Komarnitsky et al, 2000). K36me then recruits a histone deacetylase (HDAC) complex via interaction with the chromodomain of Eaf3 (Carrozza et al, 2005; Joshi and Struhl, 2005; Keogh et al, 2005), which is required for suppression of internal transcript initiation in the STE11 and FLO8 genes (Carrozza et al, 2005). In higher eukaryotic cells, the location of K36me at 3′ coding regions of active genes (Bannister et al, 2005; Carrozza et al, 2005), the observation that it is dependent upon the CTD kinase p-TEFb (Zhou et al, 2004; Eissenberg et al, 2007) and sensitive to the transcriptional inhibitor DRB (Vakoc et al, 2006) all suggest an elongation-dependent mechanism similar to that in yeast. Potential enzymes include nuclear receptor-binding SET domain-containing protein 1 (NSD1), Smyd2 and the Huntingtin-interacting protein HYPB/SETD2, all of which methylate K36 in vitro (Rayasam et al, 2003; Sun et al, 2005; Brown et al, 2006), but questions of which enzyme is relevant in vivo and whether it catalyses mono-, di- and/or trimethylation in intact cells are unanswered.
Here, we have comprehensively mapped the distribution of histone H3 K9ac, K4me3, K36me3 and K79me2 across c-fos and c-jun at high resolution, and demonstrate rapid elongation-dependent changes in K4me3 and K36me3 within the coding regions upon gene activation. These studies reveal four distinct layers of histone modification that may be distinguished across these genes. Using siRNA to knock down the relevant enzymes, we show here that Setd2, the mouse homologue of human Huntingtin-interacting protein HYPB, is responsible for virtually all detectable K36me3, but not K36me2 or K36me1, throughout the nucleus. Consistent with its proposed interaction with RNA Pol II, this enzyme mediates EGF-induced transcription-dependent increase of H3K36 trimethylation at the 3′ ends of c-fos and c-jun. This represents the first unequivocal demonstration in mammalian cells that a single histone methyltransferase (HMTase) enzyme may be entirely responsible for the totality of a specific state of methylation, that is, trimethylation but not mono- or dimethylation, throughout the nucleus.
We first carried out a comprehensive mapping study to determine the distribution of histone modifications across c-fos and c-jun in control, EGF-stimulated (Figure 1) or anisomycin-stimulated (Supplementary Figure S1) mouse fibroblasts. A modified ChIP protocol, using MNase instead of sonication to generate predominantly mononucleosomal chromatin (MacDonald et al, 2005) was used to produce high-resolution quantitative comparative maps of the distribution and dynamics of H3K4 trimethylation (K4me3), K36 trimethylation (K36me3), K79 dimethylation (K79me2) and K9 acetylation (K9ac) across these genes (Figure 1 and Supplementary Figure S1). Each antibody was first titrated in ChIP to arrive at concentrations that recovered virtually all available epitopes, leaving little or none in the unbound fraction as monitored by western blotting. Peaks of distribution of each modification across these genes are shown graphically (Figure 1B and Supplementary Figure S1B) and as bar charts with error bars (Figure 1C and Supplementary Figure S1C). This yielded highly reproducible maps with striking differences in the amounts of each modification recovered. Their interpretation is extremely complex because of the continuous dynamic turnover of acetylation at these genes, the fact that methylation may occur transiently with the traverse of RNA pol II, and the phenomenon of microheterogeneity of histone modifications at any single position. Evaluating these maps in overview, common trends applying to both c-fos and c-jun emerge. First, K9ac (Figure 1B, panel i) and K4me3 (panel ii) have remarkably similar overlapping distributions across the start site of both genes. For both modifications, there is a striking dip precisely at the start site of both genes, and the positions of peaks of modification largely coincide. The stability of this colocalisation at all points analysed is consistent with sequential ChIP assays showing these modifications coexist on the same nucleosomes at these positions (Hazzalin and Mahadevan, 2005). The sharp dip in K4me3 and K9ac at the start site on both genes appears to be due to this position lacking a nucleosome and being more accessible, as indicated by MNase sensitivity maps across these genes (Figure 2 and Supplementary Figure S2). These show that the start sites of c-fos and c-jun are considerably more sensitive to MNase digestion with <5% (c-fos −79, c-jun −57) and <10% (c-jun −396) of the corresponding genomic DNA signal remaining after digestion (Figure 2 and Supplementary Figure S2).
In contrast, K36me3 and K79me2 are found exclusively within the coding regions of c-fos and c-jun (Figure 1B, panels iii and iv). Although they do overlap, their precise distributions appear distinct. K36me3 peaks towards the 3′ end of the coding regions but drops off before the gene's terminus (Figure 1B, panel iii), whereas K79me2 peaks over the 5′ end of the coding region before declining towards the 3′ end (Figure 1B, panel iv). The presence of both these modifications in unstimulated control cells may be attributed either to their being deposited here in previous rounds of transcription prior to quiescence, or to their being part of a transcription-independent pattern of pre-existing, possibly epigenetic, mechanism; there is no evidence either way at present.
Finally, as a negative control, we monitored all four modifications at a site beyond the end of each gene (Figure 1B; c-fos +5848; c-jun +5000) and found none of these modifications were present there. Stimulation with EGF leads to rapid increases in H3K9ac on c-fos across promoter and coding region, peaking at 15 min over the promoter and 30 min over the coding region (Figure 1B, panel i left-side graph). These increases are transient and decrease substantially by 60 min (Figure 1B, panel i), correlating well with kinetics of c-fos transcription (Thomson et al, 1999). For c-jun, basal level of K9ac is much higher than for c-fos, and smaller increases are seen upon EGF stimulation, peaking at 60 min (Figure 1B, panel i, right-side graph). The elevated basal level of K9ac on c-jun may be due to a low background level of c-jun transcription in control cells (Hazzalin and Mahadevan, 2005). K4me3 and K36me3 also increase within the coding regions on c-fos and c-jun upon stimulation (Figure 1B, panels ii and iii). Compared to K9ac, these increases are slower and more sustained, remaining after 2 h of stimulation (Figure 1B, panels ii and iii). Increases in K4me3 on c-jun are much smaller than on c-fos (Figure 1B, panel ii).
The entire mapping study was repeated using a different stimulus, anisomycin, which activates these genes predominantly via stress-activated protein kinases (SAPKs) as opposed to the ERK MAP kinase pathway, which EGF utilises (reviewed in Hazzalin and Mahadevan, 2002). This yielded extremely similar MNase sensitivity maps and patterns of distribution and quantitative levels of these modifications across each gene (Supplementary Figures S1 and S2), testifying to the high reproducibility of the methodology used here.
Finally, we address a concern that activation of these genes may produce marked changes of nucleosomes occupancy across the gene, thereby skewing the ChIP recovery data provided at various time points after induction. First, there is no indication of a great change in MNase sensitivity at various time points after stimulation (Figure 2 and Supplementary Figure S2), suggesting that average nucleosomal occupancy at any position did not vary greatly upon gene activation. Second, we carried out ChIP using anti-H3 antibodies directed to the unmodified C terminus across these genes (Supplementary Figure S3A and B, panel i), and this too showed no major changes in recovery upon gene activation, confirming indications from MNase sensitivity maps. As expected, there was good correspondence between the relative H3 occupancy and the MNase sensitivity maps at all points compared across the genes (Supplementary Figure S3A and B, panels ii and iii; see for example, the dip across the start site), indicating that MNase sensitivity seen here is a good approximation of average nucleosomal occupancy.
As shown in yeast (reviewed in Gerber and Shilatifard, 2003; Hampsey and Reinberg, 2003; Sims et al, 2004), stimulation-dependent increases in K4me3 and K36me3 across coding regions suggest a link with elongating Pol II, a possibility we tested with the transcriptional inhibitor DRB. Quantitative reverse transcription PCR (qRT–PCR) shows that DRB ablates induction of c-fos and c-jun (Figure 3A). This correlates with loss of Pol II across the coding region of both genes under these conditions, shown by ChIPs using anti-Pol II antibodies raised against its N terminus (Figure 3B). Similar results were obtained at several different positions within the coding regions of c-fos and c-jun (Supplementary Figure S4A), except for +444 of c-fos, which showed increased Pol II loading in response to DRB at later time points. This may be explained by the transcriptional pause site reported at this region (Mechti et al, 1991; Coulon et al, 1999).
EGF-induced increases in K4me3 and K36me3 within the coding regions of both genes were abolished by DRB pretreatment (Figure 3C, left and middle panels). Other coding region positions within c-fos and c-jun gave similar results (Supplementary Figure S4B and C). As expected, in the negative control hbb coding region, no significant change in either modification was observed in response to EGF or DRB (Figure 3C, right panel). Thus, EGF-induced increases in K4me3 and K36me3 within coding regions of these genes are dependent on transcription elongation.
In Saccharomyces cerevisiae, Schizosaccharomyces pombe and Neurospora crassa, Set2 is an H3K36 methyltransferase responsible for mono-, di- and trimethylation (Strahl et al, 2002; Adhvaryu et al, 2005; Morris et al, 2005; Chu et al, 2006). In mammals, several candidates have been proposed. The first discovered, NSD1, has in vitro HMTase activity towards H3K36 and H4K20 (Rayasam et al, 2003). Second, a SET domain protein that interacts with the Huntington's Disease gene (HD) product, HYPB/SETD2, also methylates H3K36 in vitro and, via its C-terminal Pol II interaction domain, can bind to phosphorylated Pol II (Sun et al, 2005). Finally, Smyd2 has HMTase activity towards H3K36 in vitro, and interacts with Sin3 HDAC complexes (Brown et al, 2006). None of these studies addressed their ability to methylate H3K36 in vivo, or the specific states (mono, di or tri) of methylation for which they might be responsible. Dimethyl-H3K36 antibodies were used in the NSD1 and Smyd2 studies, but capacity for mono- or trimethylation was not analysed.
Using NCBI protein BLAST search, we identified mouse homologues of yeast Set2 (data not shown). Setd2 (the mouse homologue of human HYPB/SETD2) and NSD1 (among others) had significant homology. In view of this homology, and the fact that HYPB/SETD2 has a Pol II interaction domain (Sun et al, 2005), we investigated Setd2 and NSD1 as possible H3K36 methyltransferases in mouse fibroblasts.
Two different siRNAs targeting Setd2 transcripts knocked down Setd2 mRNA levels by >60% (Setd2 0) and >80% (Setd2 3) (Figure 4A). siRNA targeting NSD1 transcripts knocked down NSD1 mRNA levels by >80% (Figure 4A). These were specific, as mock-transfected cells, or non-targeting siRNA, did not show similar effects (Figure 4A). Additionally, Setd2- or NSD1-specific siRNAs did not knock down mRNAs of other genes tested (Figure 4A and data not shown). We raised and characterised N- and C-terminal Setd2 antibodies to confirm that Setd2 protein levels were specifically diminished as a result of its mRNA knockdown (Figure 4B, panels i and ii; Supplementary Figure S5). We were not able to assess NSD1 protein levels, as a good antibody is currently unavailable. This became less relevant since the Setd2 knockdown produced a clear H3K36 trimethylation defect.
Total cellular K36me3 was virtually absent in the Setd2 knockdowns (Figure 4C, panel i), whereas K36me2 and me1 levels remain unaffected (Figure 4C, panels ii and iii). In contrast, NSD1 knockdown had no effect on mono-, di- or trimethylation of K36 (Figure 4C, panels i–iii). Additionally, total levels of K4me3, K9me3 or K79me2 were unchanged in either Setd2 or NSD1 knockdowns (Figure 4C, panels iv–vi). Therefore, Setd2 appears to be a non-redundant methyltransferase specific for trimethylation of H3K36. This is the first demonstration in intact cultured cells of a H3K36-specific methyltransferase whose knockdown results in abrogation of virtually all trimethylation, but not mono- or dimethylation. It is not possible, however, to distinguish from these experiments if HYPB/Setd2 can catalyse all states of modification culminating in K36 trimethylation, or just the final conversion of dimethylated K36 to the trimethylated state. Since there are several recent reports of a link between H3K36 methylation and histone deacetylation in yeast (Carrozza et al, 2005; Joshi and Struhl, 2005; Keogh et al, 2005), we also analysed global levels of various H3 and H4 acetyl modifications in Setd2 knockdown cells. No significant reproducible changes were observed for any acetyl modification (Figure 4C, panels ix–xv).
Next, we used ChIP to analyse all states of K36 methylation at c-fos- and c-jun coding regions (mid and 3′ end) after Setd2 knockdown. Four housekeeping genes were also analysed; glyceraldehyde-3-phosphate-dehydrogenase (gapdh), RNA-polymerase III subunit b (polr3b), glutaminyl-tRNA synthetase (glnrs) and cyclophilin b (cycb) (shown schematically in Figure 5A).
Setd2 knockdown produced almost complete loss of total cellular K36me3 in these cells (Figure 5B) and a marked reduction of K36me3 at c-fos- and c-jun coding regions (Figure 5C, panel i). Stimulation with EGF for 60 min resulted in a 2.7- to 3.4-fold increase in the K36me3 observed at these regions (black bars panel i), whereas in the Setd2 knockdown, induced levels of K36me3 were much smaller—1.3- to 1.8-fold (middle bars panel i). K36me3 was also decreased by >50% within the coding regions of gapdh, cycb, polr3b and glnrs after Setd2 knockdown (Figure 5D, panel i). Setd2 therefore appears to be responsible for both basal and inducible K36me3 at IE gene coding regions, and K36me3 levels at constitutively active genes.
For all points tested, the recovery of nucleosomes with H3 bearing mono- and dimethyl K36 was very much lower (>10-fold) than that of trimethyl K36 (Figure 5C). In untransfected cells, there is no increase in mono- and dimethylated K36 upon activation of c-fos and c-jun (Figure 5C, panels ii and iii), indicating no correlation between these modifications and transcription. In Setd2 knockdown cells, there is no decrease in these low levels of K36me2 or me1 within the coding regions of c-fos and c-jun, with the exception of the c-jun +1651 region where there is a small decrease in K36me2 (Figure 5C, panel ii). Thus, in general Setd2 does not appear to be responsible for the low levels of mono- or dimethyl K36 at these regions.
As described for c-fos and c-jun, recovery of K36me1 and me2 was also very much lower than that of K36me3 at coding regions of gapdh, cycb, polr3b and glnrs (Figure 5D). In fact, these levels are very similar to that seen at the inactive hbb coding region (Figure 5C, panels ii and iii bottom graphs). Knockdown of Setd2 resulted in slightly higher levels of K36me2 and K36me1 at these positions (Figure 5D, panels ii and iii) but recoveries remain relatively very low. Thus, K36me2 and K36me1 levels do not seem to correlate with active transcription as K36me3 does, and Setd2 does not appear to be responsible for these modifications. By their nature, knockdowns do not produce the complete loss of any enzyme and low residual activity of the remaining HYPB/Setd2 enzyme may explain the incomplete loss of H3K36 sometimes seen.
To investigate the potential role of K36me3 in transcriptional regulation, the effect of K36me3 loss on the transcription of IE and constitutively active genes was assessed by qRT–PCR (Figure 6). As shown previously, knock down of Setd2 mRNA (Figure 6A) led to virtually complete loss of all detectable K36me3 (Figure 6D). The gene induction kinetics of c-fos, c-jun and two other IE genes, core promoter element binding protein (CPBP) and MAP kinase phosphatase 1 (MKP1), was examined (Figure 6B). A recent report in yeast has shown that increased K36me leads to delayed induction of the HIS4 gene, whereas elimination of K36me accelerates HIS4 induction (Nelson et al, 2006). However, loss of K36me3 did not affect the kinetics of induction of any gene tested here (Figure 6B). Second, the steady-state mRNA levels of four constitutively active genes (cycb, polr3b, glnrs and gapdh) were assessed in the absence of K36me3, and no significant changes were observed for any transcript (Figure 6C). Furthermore, we did not observe any intragenic transcription from c-fos, c-jun or gapdh in K36me3-depleted cells (Figure 6E). This is in contrast to the observations in yeast where K36me appears to create a less permissive chromatin structure throughout yeast gene coding regions by recruiting HDAC complexes (Rpd3) via interaction with the chromodomain of Eaf3 (Carrozza et al, 2005; Joshi and Struhl, 2005; Keogh et al, 2005). Loss of K36me leads to increased acetylation, K4me3 and Pol II loading within specific gene coding regions and intragenic transcript initiation from the FLO8 and STE11 genes (Carrozza et al, 2005; Joshi and Struhl, 2005; Keogh et al, 2005; Kizer et al, 2005). Similar to the lack of any intragenic transcription in K36me3-depleted mouse fibroblasts (Figure 6E), we do not detect any increased H3 or H4 acetylation at 3′ regions of IE or housekeeping genes (Supplementary Figure S6), nor any change in Pol II occupancy at these genes (Supplementary Figure S7). These studies do not reveal a clear parallel at c-fos and c-jun in mammalian cells for the relationship between H3K36 methylation and histone deacetylation previously seen in yeast.
Four different layers of histone modifications can now be distinguished across inducible genes in mammalian cells. The first, which includes K4me3 across the start site as well as K79me2 and K36me3 in the coding region (Figure 1) are pre-existing, possibly epigenetic histone modifications detectable in quiescent G0 cells. The second relates to dynamic turnover of histone acetylation by the action of HATS and HDACs, which affects all K4me3-modified H3 in the mouse nucleus (Hazzalin and Mahadevan, 2005; reviewed in Clayton et al, 2006). Both of these are detectable in quiescent cells. Upon stimulation, a third signalling-dependent set, which includes H3 phosphorylation at IE genes (Chadee et al, 1999; Cheung et al, 2000; Clayton et al, 2000) or H4 acetylation at hsp70 (Thomson et al, 2004) becomes evident, also seen mostly across the start site. Finally, there are DRB-sensitive modifications restricted to coding regions of these genes that are dependent upon passage of Pol II, such as enhanced K4me3 and K36me3 (Figure 3C). Thus, a combination of epigenetic, dynamic, kinase- and Pol II-dependent histone modifications is required to fully explain their region and stimulus-specific appearance across these genes.
Enhancement of K36me3 peaks towards the 3′ end of the induced c-fos and c-jun genes. There is relatively low recovery of K36me2 and me1, and no increase upon stimulation. Further, mono- and dimethyl K36 states do not correlate with transcription as similar low levels (~1% input) are detectable at active or inactive genes (Figure 5C hbb, and 5D, panels ii and iii). This emphasises the importance of considering quantitative aspects of ChIP when evaluating reports in yeast (Krogan et al, 2003; Morillon et al, 2005; Rao et al, 2005) and metazoa (Zhou et al, 2004; Bannister et al, 2005; Miao and Natarajan, 2005) showing that both di- and trimethylated K36 are associated with active genes.
The distinctive distribution of K4me3 (peaking 5′) and K36me3 (peaking 3′) across these active genes (Figure 1B and Supplementary Figure S1B, panels ii and iii) may result from selective methyltransferase recruitment to differentially phosphorylated Pol II, as shown in yeast (reviewed in Hampsey and Reinberg, 2003). MLL1, a mammalian H3K4-specific methyltransferase, is recruited to the 5′ end of active genes (Milne et al, 2002; Nakamura et al, 2002; Dou et al, 2005; Guenther et al, 2005). MLL1 or components of MLL1-containing complexes interact with Pol II phosphorylated at the Ser 5 position of its CTD (Hughes et al, 2004; Milne et al, 2005), the form of Pol II associated with 5′ coding regions (Komarnitsky et al, 2000). HYPB/SETD2 has been shown to coimmunoprecipitate with Ser 2-phosphorylated Pol II (Sun et al, 2005), suggesting that like Set2, it may ‘piggyback' with Pol II across the gene to direct co-transcriptional trimethylation of K36 at the 3′ coding regions of c-fos and c-jun. Attempts to use our N- and C-terminal antibodies to show association of HYPB/Setd2 with these regions by ChIP were unsuccessful (data not shown), but as we do not have a positive control for these experiments, they are not conclusive.
In contrast to its homologue Set2, which is responsible for all three methyl states in S. cerevisiae, S. pombe and N. crassa (Adhvaryu et al, 2005; Morris et al, 2005; Chu et al, 2006), Setd2 is specifically responsible for K36 trimethylation in mouse cells (Figures 4C and 5B–D). This suggests a more complex multi-enzyme system for K36 methylation in higher eukaryotes, similar to that for K4 methylation. Set1 is the only enzyme shown to methylate K4 in yeast, whereas many enzymes exist in mammals, including hSet1, MLL1–4, Set7/9, SMYD2 and Meisetz (reviewed in Ruthenburg et al, 2007). Some of these are methyl-state specific. For example, Meisetz specifically catalyses trimethylation of K4 in vivo (Hayashi et al, 2005). Mouse NSD1 was previously shown to dimethylate K36 in vitro (Rayasam et al, 2003), but the mono- and trimethyl states were not analysed. In this study, NSD1 did not appear to be responsible for total nuclear mono-, di- or trimethyl K36 (Figure 4C, panels i–iii) nor that at c-fos and c-jun (data not shown). Other potential K36 mono- and/or dimethyltransferases include the closely related NSD2 and NSD3 proteins, or the Smyd2 protein recently found to dimethylate K36 in vitro (Brown et al, 2006). This study indicates an unexpectedly strict relationship between a specific state of H3K36 methylation and a single methyltransferase enzyme, a finding that may eventually be found relevant for other HMTases in mammals.
Anisomycin, TSA and DRB were supplied by Sigma and rhEGF by Promega. siRNAs were synthesised by Dharmacon. PCR primers and 5′ FAM 3′ TAMRA probes were synthesised by Eurogentec.
H3K36me3 (ab9050), H3K36me1 (ab9048), H3K18ac (ab1191) and unmodified H3 (ab7834 for western and ab1791 for ChIP assays) antibodies were from Abcam. H3K36me2 (07-369), H3K9me3 (07-442), H3K14ac (07-353), H3K79me2 (07-366), H4K8ac (06-760), H4K12ac (06-761) and H4K16ac (06-762) antibodies were from Upstate, and Pol II antibody (sc-899) was from Santa Cruz. Anti-pan-acetyl H4 was a gift from Dr Tim Hebbes (University of Portsmouth). C-terminal MLL1 antibody was a gift from Dr James Hsieh (Washington University, St Louis).
In-house antibodies were raised by PTU/BS (Pentlands Science Park, Penicuik, Midlothian, UK), using peptides from Alta Bioscience (University of Birmingham). Rabbit anti-H3K9ac and anti-H3K4me3 antisera were raised against the peptides ARTKQTARK(ac)STGGKAPRKQLC and ARTK(me3)QTARKSGGC, respectively. N-Setd2 and C-Setd2 antisera were raised in rabbit and sheep, against the peptides MERRGKYTSKLERESKRTSEC and RGTKRPDDRYDTPTSKKKVRC, corresponding to amino acids 1–20 and 1560–1579 of mouse Setd2 (NCBI accession no.: XP_900231), respectively. All peptides were synthesised with C-terminal cysteine for coupling to keyhole limpet haemocyanin (KLH).
C3H 10T1/2 mouse fibroblasts were cultured in DMEM containing 10% (v/v) fetal calf serum (FCS) and 2 mM glutamine. Cells (70–90% confluent) were quiesced by transferring to DMEM with 0.5% (v/v) FCS and 2 mM glutamine for 24–28 h prior to stimulation and treatment as described.
Extraction of histones, SDS–PAGE and western blotting were carried out essentially as described previously (Cano et al, 1995; Clayton et al, 2000). Setd2 and histones were resolved on 8 and 15% polyacrylamide gels, respectively. Antibodies were used at the following dilutions: H3K4me3 and H3K9ac, 1:10,000; H3K36me3, H3K36me2 and H3K79me2, 1:5000; H4K8ac, H4K12ac and H4K16ac, 1:4000; H3K14ac, MLL1, C-Setd2 and N-Setd2, 1:2500; H3K36me1, H3K18ac and unmodified H3, 1:2000; H3K9me3 and pan-acetyl H4, 1:1000.
Cells were cultured to 40–60% confluence before washing with, and transferring to OptiMEM® I medium (Invitrogen). Cells were transfected with siRNA (50 nM in final medium) using Lipofectamine 2000 (Invitrogen) according to the manufacturer's instructions. After 5 h, an equal volume of DMEM medium containing 20% (v/v) FCS and 2 mM glutamine was added. Total RNA was extracted 24 or 48 h after transfection as described below. For protein analysis, cells were harvested in lysis buffer (0.2% (v/v) Triton X-100, 10 mM HEPES pH 8, 0.25 M sucrose, 1.5 mM MgCl2, 10 mM NaCl, 10 mM sodium butyrate, 20 mM β-glycerophosphate, 100 μM sodium orthovanadate, 1 μM microcystin and protease inhibitors (Mahadevan and Bell, 1990)) 48 h after transfection. Crude nuclei were pelleted by centrifugation and resuspended in SDS–PAGE sample buffer. Cells were quiesced 24 h after transfection as described above. Crosslinked mononucleosomes were prepared 48 h after transfection as detailed below.
Total RNA was extracted using TRIzol® reagent (Invitrogen) according to the manufacturer's instructions. For qRT–PCR, contaminating DNA was removed using the TURBO DNA-free™ kit (Ambion), and 0.5 μg of RNA was reverse-transcribed using the SuperScript™ III First-Strand Synthesis kit (Invitrogen). cDNA was then quantified by real-time PCR. For northern analysis, 5 μg of RNA was resolved on formaldehyde/agarose gels, transferred to nylon membrane (Hybond-N+, Amersham) and hybridised with probes as detailed in Hazzalin et al (1997). c-fos (+2091 –3109) and c-jun (+1853 –2920) 3′ gene fragments for probes were amplified by PCR from cDNA, labelled by random priming (Amersham RediPrime II kit) and sequentially hybridised to membranes.
Formaldehyde crosslinked mononucleosomes were prepared as described in Macdonald et al (2005). Antibodies (10 μl H3K36me3/Pol II or 5 μl H3K36me2/H3K36me1/H3K79me2/H3K4me3 serum/H3K9ac serum/Pan-acetyl H4 serum) were added to chromatin aliquots (~25–30 μg DNA) and rotated for 2 h at 4°C. Protein A-sepharose beads (~5 mg) pre-blocked with 10 μg sonicated herring sperm DNA and BSA (0.02% final concentration after addition to chromatin) were then added and samples rotated for a further 2 h at 4°C. Beads were recovered by centrifugation, and unbound supernatant chromatin was retained. Beads were washed sequentially with 3 × RIPA buffer (Clayton et al, 2000) without phosphatase or protease inhibitors, 3 × RIPA containing 500 mM NaCl and 2 × TE (10 mM Tris–HCl pH 7.5, 1 mM EDTA). Input and antibody-bound chromatins were incubated at 65°C overnight to reverse crosslinks. Samples were treated with RNase A (100 μg/ml, 1 h, 37°C) followed by proteinase K (100 μg/ml, 3 h, 37°C) before phenol/chloroform extraction of DNA. Extracted DNA was ethanol precipitated and redissolved in 10 mM Tris–HCl pH 8.0, 0.1 mM EDTA. The presence of specific gene sequences was then quantified by real-time PCR.
Real-time PCR was performed using the ABI Prism 7000 sequence detection system (Applied Biosystems). All primers were designed with Primer Express 2.0 software (Applied Biosystems). All c-fos, c-jun, gapdh and hbb amplicons were detected by the 5′ nuclease (Taqman) assay using 5′ FAM 3′ TAMRA-labelled probes and qPCR Mastermix Plus (Eurogentec). All other amplicons were detected by the SYBR® Green assay using SYBR® Green PCR Master Mix (Applied Biosystems). Sequences of primers and probes and detailed conditions are available on request. PCR reactions were carried out in triplicate or duplicate.
For ChIP assays, standard curves (serially diluted sonicated genomic DNA) were included in every plate to ensure bound and input sample dilutions were within the linear range. Data were analysed by the CT method and plotted as % input recovered in the bound fraction for each amplicon, calculated by the following formula: % input recovery=[100/(input fold dilution/bound fold dilution)] × 2(input CT - bound CT), where no fold dilution=1. For qRT–PCR, data were analysed by relative quantification using the standard curve method (see Applied Biosystems user bulletin 2). All raw data from micrococcal nuclease sensitivity and ChIP experiments are available upon request.
During review of this manuscript, Bell et al (2007) reported that in Drosophila, dHypb is responsible for the trimethylated state of H3K36, in agreement with our findings in mammalian cells. However, the global increase in H4K16 acetylation reported upon dHypb knockdown in Drosophila is not seen in mammalian cells used here (Figure 4C).
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We thank Drs Fiona Myers and Alan Thorne (University of Portsmouth) for advice on real-time PCR in the early stages of this work, and Dr James Hsieh (Washington University, St Louis) for the MLL1 antibody. We thank all members of the Nuclear Signalling Laboratory for help and discussions throughout the course of this work. JWE is supported by an MRC:CASE award with Syngenta. This work was funded by a Wellcome Trust Programme Grant (065373/Z/01/Z).