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We report a function of human mRNA decapping factors in control of transcription by RNA polymerase II. Decapping proteins Edc3, Dcp1a and Dcp2 and the termination factor, TTF2, co-immunoprecipitate with Xrn2, the nuclear 5′-3′ exonuclease “torpedo” that facilitates transcription termination at the 3′ ends of genes. Dcp1a, Xrn2 and TTF2 localize near transcription start sites (TSSs) by ChIP-seq. At genes with 5′ peaks of paused pol II, knockdown of decapping or termination factors, Xrn2 and TTF2, shifted polymerase away from the TSS toward upstream and downstream distal positions. This re-distribution of pol II is similar in magnitude to that caused by depletion of the elongation factor Spt5. We propose that coupled decapping of nascent transcripts and premature termination by the “torpedo” mechanism is a widespread mechanism that limits bidirectional pol II elongation. Regulated co-transcriptional decapping near promoter-proximal pause sites followed by premature termination could control productive pol II elongation.
Decapping is a major control step in cytoplasmic mRNA degradation (Coller and Parker, 2004; Franks and Lykke-Andersen, 2008). In mammals this step is catalyzed by two unrelated pyrophosphatases. Dcp2 occurs in a complex with the co-factors Dcp1a/b, Edc3, Hedls and Rck/p54 (Lykke-Andersen, 2002; Wang et al., 2002) and is responsible for decapping a subset of mRNAs. The single subunit enzyme Nudt16 was recently found to be responsible for the majority of cytoplasmic decapping in some cells (Song et al., 2010). While Dcp2 is mainly cytoplasmic it has been detected in the nucleus (Parker and Song, 2004; Song et al., 2010) and has been implicated in turnover of misprocessed nuclear mRNA precursors in yeast (Kufel et al., 2004). Decapping initiates 5′-3′ mRNA degradation by exposing a 5′ phosphate substrate for the major cytoplasmic 5′-3′ RNA exonuclease Xrn1 (Muhlrad et al., 1994).
The conserved nuclear homologue of Xrn1 is the exonuclease Xrn2, Rat1 in yeast, which acts as a “torpedo” (Connelly and Manley, 1988) that facilitates poly(A) site-dependent termination by RNA polymerase II (pol II) downstream of genes (Kim et al., 2004; West et al., 2004). In yeast, Rat1 is also required for optimal termination of rDNA transcription by pol I (El Hage et al., 2008; Kawauchi et al., 2008). Just as decapping provides an entry point for Xrn1 on mature mRNAs in the cytoplasm, co-transcriptional poly (A) site cleavage provides a 5′ phosphate entry site for Xrn2/Rat1 that then degrades nascent pol II transcripts in the nucleus. The Xrn2/Rat1 “torpedo” is necessary but not sufficient to evict pol II from the template (Dengl and Cramer, 2009; Luo et al., 2006). Additional unknown factors probably cooperate with Xrn2 to elicit termination. One candidate is the DNA-dependent ATPase TTF2/lodestar, which can release pol II and RNA transcripts from a DNA template in vitro (Liu et al., 1998) and helps eliminate pol II from condensed mitotic chromosomes (Jiang et al., 2004). Interestingly, TTF2 physically interacts with the decapping factor Dcp1a (Leonard et al., 2003).
Control of the flux of elongating pol II through promoter-proximal pause sites is important for regulation of a large fraction of genes in multicellular organisms (Core et al., 2008; Muse et al., 2007; Nechaev and Adelman, 2011; Rahl et al., 2010; Zeitlinger et al., 2007). As a result of promoter-proximal pausing, high densities of pol II accumulate at the TSSs of most human genes with much lower densities downstream within genes. Promoter-proximal pausing is facilitated by the negative elongation factors NELF and DSIF (Spt4/5) and antagonized by the positive elongation factor P-TEFb, TFIIS and transcriptional activators (Chiba et al., 2010; Nechaev et al., 2010; Peterlin and Price, 2006). Control of the transition from pausing to productive elongation is widely thought to be a mechanism for rapidly up-regulating transcription in response to developmental and environmental cues (Bentley and Groudine, 1986; Core and Lis, 2008; Gilmour and Lis, 1986; Krumm et al., 1995; Levine, 2011; Rahl et al., 2010; Zeitlinger et al., 2007). However the fate of paused polymerases in vivo is a long-standing unresolved question. In principle these polymerases could resume productive elongation in response to an appropriate signal or they could terminate prematurely in a manner analogous to prokaryotic transcriptional regulatory mechanisms (Nudler and Gottesman, 2002).
Premature termination near 5′ ends has been detected within yeast genes where it is promoted by Nrd1, Nab3, and Sen1 (Gudipati et al., 2008; Lykke-Andersen and Jensen, 2007; Steinmetz et al., 2001; Vasiljeva et al., 2008). In addition, premature termination involving the Rat1 exonuclease has been detected in yeast mutants that are defective for mRNA capping (Jiao et al., 2010; Jimeno-Gonzalez et al., 2010). In these mutants premature termination by a torpedo mechanism is thought to serve a quality control function preventing production of full-length transcripts with defective cap structures. In mammalian cells premature termination of transcription has been demonstrated for SV40 and HIV viral genes and its mechanism is unknown (Hay et al., 1982; Kao et al., 1987; Toohey and Jones, 1989).
While there is only sparse evidence for promoter-proximal termination in metazoans, the recent finding that transcription frequently initiates in both directions at human promoters (Core et al., 2008; Preker et al., 2008; Seila et al., 2008) begs the question of whether antisense transcription is limited by a novel termination mechanism. In this paper we report that decapping proteins and TTF2 interact with Xrn2 and that these factors localize by ChIP at 5′ ends of genes. Knockdown of decapping and termination factors by shRNA caused a widespread re-positioning of pol II at 5′ ends of genes away from start sites and toward distal positions both downstream and upstream. These results suggest that co-transcriptional decapping and premature termination by a torpedo mechanism is broadly employed to limit transcription of human genes.
To identify candidate termination factors that associate with Xrn2, we performed mass spectrometry of Xrn2 immunoprecipitates (IPs) from RNAse-treated HeLa nuclear extract. TTF2 was among the most strongly enriched proteins in the Xrn2 IP relative to the control anti-GFP IP and was confirmed by Western blotting (Fig. 1A, B). TTF2 interacts with Dcp1a in the yeast two-hybrid assay (Leonard et al., 2003) and consistent with this association, we found that Dcp1a and two other decapping factors Edc3 and Dcp2 co-IP with Xrn2 (Fig. 1A,B). The pol I termination and chromatin re-modeling factors TTF-I and Rsf1 (Längst et al., 1997) were also strongly enriched in the Xrn2 IP (Fig. 1A) suggesting a role for Xrn2 in termination of human rRNA transcription as previously demonstrated in yeast (El Hage et al., 2008; Kawauchi et al., 2008). Additional Xrn2-associated proteins identified by MS include rRNA maturation factors, splicing factors and cleavage-polyadenylation factors (Fig. 1A, Table S1).
We investigated the functional significance of the co-purification of decapping factors and TTF2 with Xrn2. The association of decapping factors with a nuclear enzyme Xrn2 was surprising as decapping is thought to be predominantly cytoplasmic (Franks and Lykke-Andersen, 2008; Song et al., 2010). However, we found that decapping activity is readily detectable in Hela nuclear extract and can be specifically immunoprecipitated by anti-Dcp1a antibody (Fig. S1A). We investigated whether decapping factors might act in the nucleus at sites of transcription by ChIP-Seq analysis of Dcp1a and pol II in Hela cells. Because Dcp1a interacts with TTF2 (Leonard et al., 2003) and Xrn2 (Fig. 1), we further tested whether these three factors co-localize on genes by ChIP-seq with antibodies whose specificity was confirmed by shRNA knockdown of the respective proteins (Fig. S1B, (Jiang et al., 2004)). The Dcp1a ChIP-seq signals were sufficient to identify 745 genes with peaks of enrichment within 500 bases of a TSS (FDR < 0.05, Table S2) detected by the HOMER peak finder (Heinz et al., 2010). Note that while the peak finding analysis of our ChIP-seq results reveals significant enrichment of Dcp1a, Xrn2 and TTF2 near TSSs, it does not provide a complete list of genes where these factors are found at the 5′ end. Furthermore Xrn2 and TTF2 co-localized with Dcp1a near these start sites to a significant extent (Figs. 1C, D, Tables S3, S4). Interestingly peaks of Xrn2 and TTF2 accumulation (FDR <0.05) were more frequent within 500 bases of a TSS than they were in the region 0–3kb downstream of a poly(A) site (Table S2). Importantly, the ChIP signals for Dcp1a, TTF2 and Xrn2 overlap extensively with one another and with pol II at several thousand genes in the region between −1kb and +2kb from the TSS (Figs. 1D, S2E, see also Tables S3, S4). We confirmed the localization of Xrn2 at 5′ ends by ChIP in two human breast cell lines in addition to Hela (Fig. S2A). In addition when transcription elongation was inhibited with DRB, the distribution of Xrn2 on the GAPDH gene shifted toward the 5′ end of the gene in parallel with pol II (Fig. S2B, C). The localization of TTF2 at 5′ ends of genes was further confirmed by ChIP-seq using an independent commercial polyclonal antibody (Fig. S2D). We conclude that these termination and decapping factors commonly localize near the 5′ ends of genes.
Co-localization of a decapping factor with Xrn2 at 5′ ends suggests a possible parallel with cytoplasmic mRNA degradation by decapping and 5′-3′ exonucleolytic degradation by Xrn1 (Coller and Parker, 2004). According to this model, decapping of nascent transcripts coupled to Xrn2-mediated degradation might cause premature termination of transcription by the “torpedo” mechanism. If decapping factors, Xrn2 and TTF2 terminate a population of promoter-proximal polymerases, then their depletion would enhance readthrough transcription downstream and/or upstream of promoters. In this event relative pol II density in promoter-distal regions is expected to increase relative to promoter-proximal regions. To test this prediction, we first knocked down Xrn2 and TTF2 individually and together in stable HEK293 cell lines by infection with lentiviral shRNA expression vectors (Fig. S1B, Table S5). We then measured pol II occupancy along genes by ChIP-Seq in the knockdown lines relative to two controls, the uninfected parent and a line infected with a scrambled shRNA lentivirus. Knockdown of Xrn2 and/or TTF2 had only small effects on the overall enrichment of pol II within genes compared to parent or scrambled shRNA controls, as determined by anti-pol II ChIP-seq reads per kilobase per million in the region from −30 to the poly(A) site (Fig. 2B, S3C). The depletion of Xrn2 and TTF2 achieved in our cell lines therefore does not cause widespread inhibition of transcription initiation. Pol II density profiles normalized to total read-counts (Fig. 2A) revealed that, as predicted by the model, on genes where pol II normally accumulates in the promoter-proximal region, knockdown of Xrn2+TTF2 markedly increased pol II density at promoter-distal positions and decreased it in promoter-proximal positions. Knockdown of Xrn2 or TTF2 individually, caused a similar redistribution of pol II from promoter-proximal to promoter-distal positions, but the effects were smaller than in the double knockdown (Fig. S3A). The complementary changes in pol II density at distal and proximal sites are consistent with a redistribution of polymerase away from the start site and into the body of the gene. In contrast only modest effects on pol II 5′-3′ distribution were evident on genes with lower levels of promoter-proximal pol II accumulation including HIST1H4C, SFN, and ARHGDIA and the non-coding RNA gene HOTAIR (Fig. S4A–D).
On the RBM39 gene where divergent antisense transcription occurs (Fig. S5D) (Core et al., 2008; Preker et al., 2008), depletion of Xrn2+TTF2 modestly increases pol II density upstream of the TSS as well as downstream (Fig. 2A). This observation suggests that Xrn2 and TTF2 may normally limit the amount of divergent antisense transcription. Similar enhancement of antisense transcription in the Xrn2+TTF2 knockdown cells occurred at the ARHGDIA, ID1 and FRAT2 genes (Fig. S4D–F). These observations on individual genes were confirmed in an average of many genes (see Figs 3A, B).
Xrn2 and TTF2 shRNAs did not cause a major inhibition of transcription termination downstream of poly(A) sites (Fig S5E) at the level of knockdown achieved in our cells consistent with a previous report (Banerjee et al., 2009). At some genes like ACTB however termination was delayed (Fig. 2A). In summary the results in Figure 2A show that depletion of two termination factors that localize at 5′ ends of genes caused a shift in the distribution of pol II from promoter-proximal to promoter-distal positions both upstream and downstream of the TSS.
To determine how generally Xrn2/TTF2 depletion affects transcription within human genes, we assessed pol II distribution on a cohort of 5507 genes in each knockdown cell line (Table S2). The genes analyzed were selected on the basis that they are >2kb long and >2kb from neighboring genes and are significantly enriched (FDR<0.05) for pol II in the region within 500 bases of a TSS. First we plotted the relative frequency of pol II ChIP-seq reads on these genes in the region from −1kb to +2kb from the TSS for the Xrn2+TTF2 double knockdown and scrambled shRNA control lines (Fig. 3A). This analysis shows that the relative increase in pol II read frequency at positions both upstream and downstream of the TSS in the double knockdown line is detectable even when averaged over more than 5000 genes. To quantify the effects of Xrn2 and TTF2 depletion on a broad scale, we plotted the ratios of the relative frequencies of pol II ChIP-seq reads (see Experimental Procedures) in each knockdown cell line compared to the scrambled shRNA control (Fig. 3B). As a control we compared the uninfected parent line to the scrambled shRNA (grey line Fig. 3B) and as expected there was little difference. In contrast, each of the knockdown lines demonstrated a specific increase in the relative frequency of pol II at promoter-distal positions compared to the scrambled control with the greatest effect in the Xrn2:TTF2 double knockdown (Fig. 3B). The apparent additive effect of depleting TTF2 and Xrn2 is significant because the only known function in common between these proteins is in transcription termination. Notably depletion of termination factors increased relative pol II frequency at positions both downstream and upstream of the TSS and these effects are widespread as evidenced by the fact that they are detectable in plots averaged over the group of 5507 genes.
To investigate further how termination factors influence pol II distribution across genes, we calculated the escape index (EI), defined as log2 promoter-distal (+301 to the polyA site) pol II density: promoter-proximal (−100 to +300) pol II density (Fig. 3C) in control and knockdown HEK293 cell lines for 5507 genes. This analysis, revealed significant (FDR< 0.01) increases in EI for many genes in the Xrn2+TTF2 double knockdown line, but not the uninfected parent cells, relative to the scrambled shRNA control (Fig. 3D). Importantly the greatest effects of Xrn2/TTF2 depletion on EI were evident for those genes with low EI values that have the greatest accumulation of promoter-proximal pol II density. In contrast genes in the minority class of human genes with high EI values and low levels of promoter-proximal pol II accumulation, were less affected by depletion of these termination factors. This point is particularly clear for the Xrn2:TTF2 double knockdown (Fig. 3D, red) and is further supported by inspection of individual genes with low levels of promoter-proximal pausing and high EI values (Fig. S4A–D). In summary, the results in Figure 2 and and33 show that at many genes the termination factors Xrn2 and TTF2 limit pol II elongation away from the TSS in both directions, consistent with widespread premature termination in promoter-proximal regions.
If Xrn2 and TTF2 cooperate to facilitate termination at promoter-proximal regions by a “torpedo” mechanism, then decapping or RNA cleavage would be required to provide a suitable 5′ phosphorylated substrate for the exonuclease. To test whether decapping factors affect how pol II is distributed between promoter-proximal and distal regions, we stably knocked down Edc3, Dcp1a or Dcp2 in HEK293 cells (Fig. S1C) and analyzed pol II distributions by ChIP-Seq. Similar to the termination factors, depletion of Dcp2, Edc3, or Dcp1a did not cause major changes in overall levels of pol II enrichment within genes when compared to the scrambled shRNA control (Fig. 4B, S3D). Depletion of all these decapping factors, especially Dcp2, specifically increased pol II densities at promoter-distal locations and decreased it at the TSS (Figs. 4A, S3B). As we observed for Xrn2/TTF2, depletion of decapping factors enhanced relative pol II accumulation at the 3′ pause that precedes termination at some genes like ACTB probably because more polymerases reach the 3′ end (Fig. 4A). Knockdown of decapping factors increased pol II relative frequency at promoter-distal positions both upstream and downstream of the TSS (Fig. 5A, B). In addition, plots of escape index (EI, Fig. 3C) for the group of 5507 genes used in Figure 3 revealed that the promoter-proximal to -distal shift in pol II distribution caused by depleting decapping factors is widespread (Fig. 5C). In summary, these results reveal a surprising effect of depleting decapping factors on pol II transcription that strongly resembles the effect of depleting the termination factors Xrn2 and TTF2. We interpret these results to suggest that decapping, coupled to polymerase displacement by the Xrn2 “torpedo” and the ATPase TTF2, facilitates premature termination of bidirectional transcription from human promoters.
The current model for control of pol II transcriptional elongation on a large fraction of metazoan genes is by regulated release from a promoter-proximal pause that is established by NELF and DSIF (Spt4/5) (Chiba et al., 2010; Core and Lis, 2008; Levine, 2011; Nechaev and Adelman, 2011; Peterlin and Price, 2006). Previously the effects of depleting NelfA and Spt5 on transcription were defined by pol II ChIP-Seq in mouse embryonic stem (ES) cells (Rahl et al., 2010) and we compared these results with the effects of depleting decapping and termination factors. In agreement with Rahl et al (2010), pol II relative frequencies and EI on over 5000 mouse genes showed that knockdown of Spt5 increased pol II density at downstream positions relative to the TSS (Fig. 5D, E) just as we saw for knockdown of termination and decapping factors (Figs. 3A,B, 5A,B). The magnitude of the increase in relative pol II density within genes caused by Spt5 knockdown was similar to that caused by Edc3, Xrn2, or TTF2 knockdown and less than Dcp1a, Dcp2 or Xrn2:TTF2 double knockdown. In contrast with the latter factors, Spt5 knockdown appeared to have little effect on divergent transcription as judged by pol II densities upstream of the TSS while the NelfA knockdown had a much more modest effect on average pol II distribution among these genes (Fig. 5E) as previously reported (Rahl et al., 2010). Note that Spt5 knockdown resulted in a step like increase in pol II density immediately downstream of start sites whereas termination and decapping factor knockdown causes a gradual increase in relative pol II density over a 2kb region downstream of the start site. This effect is consistent with premature termination by the “torpedo” occurring over hundreds of bases, similar to termination downstream of genes.
Regulated premature termination could function to control transcription by modulating the flux of polymerases through the body of a gene. We investigated this idea by examining genes that are regulated at the elongation level. At PIM1, ENO1, CCNB1 and UBA52 that are activated by Myc and HSP90AA1 and HSPA8, that are activated by HSF, depletion of Dcp2 or Xrn2+TTF2 increased pol II density within the gene body relative to the TSS (Figs. 6A–D, S6). Consistent with the results in Figures 5D–F, the effects of TTF2/Xrn2 and Dcp2 knockdown on pol II distribution at individual genes are at least as great as the effects of depleting NelfA or Spt5 (Figs. 6A–D, S6). How the transcriptome is affected by global changes in pol II elongation caused by depletion of pausing regulators (Rahl et al., 2010) or decapping/termination factors remains to be determined, but it is likely to be influenced by homeostatic mechanisms that control mRNA stability. Knockdown of Xrn2+TTF2 increased the steady-state level of HSP90AA1 mRNA relative to a mitochondrial mRNA control by 1.6 fold as determined by Q RT-PCR but had no significant effect on abundance of the Myc target mRNA UBA52 (Fig. S7A). Dcp2 and Xrn2+TTF2 depletion by shRNA, but not a scrambled shRNA control, significantly elevated EI on most genes within a group of 173 Myc (Li et al., 2003) and HSF targets (Table S2) thought to be activated by release of promoter-proximal pausing (Fig. 6E). We conclude that depletion of decapping and termination factors can mimic the effects of activators that stimulate transcriptional elongation.
We report a previously unidentified function for mRNA decapping enzymes in the nucleus that is distinct from their well-known role in cytoplasmic mRNA turnover. We propose that in the nucleus, decapping of nascent transcripts near sites of promoter-proximal pol II pausing facilitates premature termination of transcription by providing an entry point for the 5′-3′ RNA exonuclease “torpedo”, Xrn2, (Connelly and Manley, 1988) previously shown to function in termination by pol I and pol II at 3′ ends of genes (El Hage et al., 2008; Kawauchi et al., 2008; Kim et al., 2004; West et al., 2004) (Fig. 7). Consistent with a general role in termination, we found that Xrn2 also co-purified with the pol I termination factors TTF-I and Rsf1(Längst et al., 1997) (Fig. 1A). We further suggest that premature termination is aided by the ATPase, TTF2, that interacts with the decapping factor Dcp1a (Leonard et al., 2003) and displaces pol II from template DNA in vitro (Liu et al., 1998). In support of this model we have presented two lines of evidence:
While we cannot exclude the possibility that Edc3, Dcp1a, Dcp2, TTF2, and Xrn2 might also influence pol II pausing or elongation rate, the most straightforward interpretation of our results is that they affect pol II distribution through their well-established functions in decapping and transcription termination because these two functions can account for all our observations. Hence according to the two-step model in Figure 7, co-transcriptional decapping first acts to expose a 5′ monophosphate end on the nascent RNA and this provides a substrate for Xrn2 to begin degrading the transcript. Knockdown of Xrn2 or TTF2 caused relatively modest effects but the double knockdown induced a larger pol II redistribution into gene bodies (Figs. 3B, D, S3A). The only known function in common between these two proteins is transcription termination. Therefore, the similarity and additive nature of these defects are consistent with both proteins participating in promoter-proximal termination. It remains possible that TTF2 or decapping factors could also influence the processivity of the Xrn2 exonuclease. A re-distribution of pol II from the TSS into the gene body rather than a selective slow-down of elongation within the gene is consistent with the fact that elevated pol II density within gene bodies was largely offset by reduced density around start sites (Figs. 2A, ,4A).4A). As a result, overall pol II occupancy on genes was little affected by depletion of termination or decapping factors in our cell lines (Figs. 2B, ,4B,4B, S3C, D). In future it will be important to test the co-transcriptional decapping model by developing methods for determination of the cap status of nascent transcripts at genes with and without evidence of premature termination.
We note that frequent premature termination by a “torpedo” mechanism in promoter-proximal regions could affect the interpretation of nuclear runon experiments because runon transcripts made by polymerases that have engaged the Xrn2 nuclease may be degraded. This might explain why the excess of promoter-proximal pol II detected by ChIP is not always matched by an equivalent excess of runon signal detected by GRO-seq (Core et al., 2008)(Fig S5A–D).
Regulated premature termination is a well-characterized mechanism of controlling gene expression in prokaryotes (Nudler and Gottesman, 2002; Yanofsky, 2000). In eukaryotes, premature termination by pol II has been detected on viral transcription units (Hay et al., 1982; Kao et al., 1987; Toohey and Jones, 1989) and at some yeast genes (Gudipati et al., 2008; Steinmetz et al., 2001; Vasiljeva et al., 2008) but there is no previous evidence to suggest that it has widespread significance in metazoans. The results presented here suggest that in fact premature termination by a “torpedo” mechanism prevents productive elongation by a substantial fraction of the promoter-proximal pol II transcription complexes at thousands of human genes. Premature termination by this mechanism occurs in yeast mutants defective in mRNA capping (Jiao et al., 2010; Jimeno-Gonzalez et al., 2010) where it prevents production of full-length uncapped transcripts. The extent of the pol II re-distribution effected by termination and decapping factors is not easily reconciled with a function limited to quality control through elimination of the small fraction of transcription complexes with incompletely capped nascent transcripts. Instead our results imply that premature termination by decapping coupled with pol II displacement by Xrn2/TTF2 is a quite general mechanism for limiting productive transcriptional elongation. Depletion of termination and decapping factors enhanced relative pol II occupancy at positions upstream of many promoters while decreasing it near the start sites (Figs. 3A,B, 5A,B). This result suggests that co-transcriptional decapping and polymerase eviction facilitated by Xrn2/TTF2 accounts for termination of at least some of the divergent transcription that commonly occurs at human promoters (Core et al., 2008; Preker et al., 2008; Seila et al., 2008). In addition to its role as a termination factor, Xrn2 has been implicated in formation of the short sense and antisense transcription start site-associated (TSS-a) RNAs that lack cap structures (Seila et al., 2008; Valen et al., 2011). The mechanism of TSS-a RNA formation is not known but it could involve the decapping and exonucleolytic degradation of nascent transcripts.
In summary, our results suggest that co-transcriptional decapping and termination aided by Xrn2/TTF2 is an important constraint on the flux of pol II through thousands of genes. Indeed premature termination appears to exert an effect on transcription in vivo that is as general and as profound as DSIF and NELF mediated promoter-proximal pausing (Figs. 5, ,6,6, S6). In future it will be of interest to investigate how co-transcriptional decapping and subsequent termination are controlled and how they may be affected by activators and promoter-proximal pausing. We speculate that the promoter-proximal pause found at most human genes may serve as a decision point for regulated decapping (Fig. 7). Paused elongation complexes that undergo decapping will be prematurely terminated whereas complexes that escape decapping will retain the potential to resume productive elongation subject to regulation by TFIIS, DSIF, NELF, and PTEFb (Chiba et al., 2010; Nechaev et al., 2010; Peterlin and Price, 2006). The possibility that decapping and coupled premature termination could be subject to regulation is consistent with our observation (Figs. 6, S6) that the effects of depleting decapping and termination factors closely resemble the effects of activators like c-Myc and HSF that stimulate transcriptional elongation (Brown et al., 1996; Rahl et al., 2010). Whether decapping and termination factors are recruited to 5′ ends of genes together in a complex or sequentially remains to be resolved however our preliminary results indicate that Xrn2 is recruited independently of decapping factors because knockdown of Dcp2 did not reduce Xrn2 localization at the 5′ end of GAPDH (Fig. S7). The presence of Dcp1a, TTF2 and Xrn2 at the 5′ end of a gene is not necessarily correlated with strong premature termination, perhaps because their activity is suppressed at some genes. For example decapping and termination factors are found at the 5′ ends of histone genes (data not shown) where there is little evidence of premature termination, and transcription is unaffected by their depletion (Fig. S4A). It will also be of interest to determine whether the recruitment or the enzymatic activity of Xrn2, TTF2, or Dcp2 is regulated by their interactions with one another and other proteins at various promoters. In addition to decapping factors, the deadenylase Ccr4 and the exosome have been implicated in aspects of transcription (Andrulis et al., 2002; Kruk et al., 2011; Qiu et al., 2004**************) suggesting that a broad connection may exist between transcription and RNA turnover.
HeLa and HEK293-Flp-in T-REX-glob cells were grown in DMEM medium supplemented with 10% fetal bovine serum, 1% pen/strep at 37°C and 5% CO2. HEK293-Flp-in T-REX-glob cells contain a stably integrated, hygromycin-resistance gene and CMV β-globin reporter that is not relevant to these studies.
Rabbit anti-pan pol II CTD was described previously (Schroeder et al., 2000). Anti-Xrn2 was raised in rabbits against GST-Xrn2 (a.a. 402–537) and affinity purified. Affinity purified rabbit anti-GFP (Fong et al., 2009), rabbit anti-Dcp1a, and anti-Edc3 have been described (Fenger-Gron et al., 2005; Lykke-Andersen and Wagner, 2005). Anti-Dcp2 and Anti-Aly (11G5, THOC4) were gifts of M. Kiledjian and G. Dreyfuss. Rabbit anti-TTF2 was from Proteintech and affinity purified sheep anti-TTF2 was a gift of D. Price.
Rabbit anti-Xrn2 and anti-GFP control antibodies were affinity purified and coupled to Amino-Link resin (Pierce) at 1 mg/ml and equal amounts were used for precipitation from HeLa Nuclear extract in the presence of RNAseA (20μg/ml) in buffer D (see Extended Experimental Procedures). IP’ed material was fractionated by 4–15% SDS-PAGE and each lane was cut into 14 bands. Proteins were reduced, alkylated and trypsin digested in the gel, extracted and analyzed by LCMS/MS on two analytical platforms: ThermoFisher LTQ XL and LTQ-FT Ultra. Results for the proteins most enriched in the Xrn2 IP relative to the GFP control by number of assigned spectra are listed in Table S1. Total assigned spectra were 22346 and 29540 for the anti-GFP, and 16776 and 23442 for the anti-Xrn2 LTQ and FT analyses respectively.
pLKO.1-puro shRNA lentiviruses (Open Biosystems, Table S5) were used to infect HEK293-Flp-in T-REX-glob cells. For Xrn2:TTF2 double knockdown, a neomycin resistance marker was cloned in place of the puromycin resistance marker in pLKO.1-puro TRCN0000049900 targeting Xrn2 and knockdowns were verified by western blot (Fig. S1B).
ChIP and Illumina library preparation have been described (Kim et al., 2011). Libraries were sequenced on the Illumina Genome Analyzer IIx and Hi-Seq platforms. Single-end 34 base reads (after removing barcodes) were mapped to the hg18 UCSC human genome (Mar. 2006) with Bowtie version 0.12.5 (Langmead et al., 2009)(Table S6). We generated bed and wig profiles using 50bp bins and 200bp windows assuming a 180bp fragment size shifting effect. Results were viewed with the UCSC genome browser, integrated genome browser (IGB), or the R statistical software package.
We determined the central position of each ChIPed DNA fragment and calculated its position relative the TSS. Relative frequencies are defined as read counts per 50 bp bin fixed divided by the total number of aligned reads in all bins. The y-axis in Figs. 3A, 5A, 5D represents the proportion of counts contained in each bin. In Figs. 3B, 5B, 5E log2 values of the shScramble control were subtracted from log2 values of the relative frequencies in each bin of the experimental samples. Peaks of enrichment of Xrn2, Dcp1a and TTF2 in Hela cells were mapped relative to an input DNA background using the HOMER peakfinder (v3.2) (Heinz et al., 2010) with a default option except that fragment length was set to 200bp (-fragLength 200 -inputFragLength 200). Heat map plots of pol II, Dcp1a, Xrn2 and TTF2 in Fig. 1D were determined in the target range 1kb up and 2kb downstream from the TSS of RefSeq genes. Reads per bin per million bases (RPBM, binsize = 50bp) were determined by normalizing each sample to total aligned reads (in millions). Genes were sorted by pol II density.
We defined the Escape Index (EI) to measure the flux of Pol II from the promoter region into the body of a gene. EI corresponds to
where body_density = # tags per base in a range between +301 base from TSS and poly(A) site and promoter_density = # tags per base in a range between −30 and +300 base from TSS. EI changes (ΔEI) in knockdown cells relative to the shScrambled control were compared by one-sample T-test for each gene assuming as a null hypothesis a normal distribution fitted by ΔEI of untreated parent to the shScramble control. We obtained a cut-off threshold for the false discovery rates (FDR) by correcting for multiple p values (Benjamini and Hochberg, 1995).
This work was supported by NIH grant GM063873 to D.B. K.B. and K.G-C. were supported by Ruth L. Kirschstein NRSA awards F31GM095249 and F31GM72099. K.B. was supported by NIHT32-GM08730. S.K. was supported by the American Cancer Society and the Clark family (award PF-07-297-01-GMC). H.K. was supported by ARRA award 3R01GM063873-06S1, R.E.D. was supported by NIH grant AI49558; K.H was supported by NIH grant P30 CA046934-17 and NCRR grant S10RR023015. We thank U. Colorado Functional Genomics Core for shRNA constructs, M. Kiledjian (Rutgers U.), G. Dreyfuss (U. Penn) and D. Price (U. Iowa) for generous gifts of antibodies, T. Blumenthal, S. Johnson, T. Evans, J. Kim and R. Perales for helpful suggestions and J. Dover, D. Farrell, J. Castoe, B. Gao and K. Diener for Illumina sequencing.
Accession Numbers ChIP-seq data sets are deposited at GEO accession GSE36185.
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