1. Chromosomes and Gene Expression

Reporter-ChIP-nexus reveals strong contribution of the Drosophila initiator sequence to RNA polymerase pausing

  1. Wanqing Shao
  2. Sergio G-M Alcantara
  3. Julia Zeitlinger  Is a corresponding author
  1. Stowers Institute for Medical Research, United States
  2. University of Kansas Medical Center, United States
https://doi.org/10.7554/eLife.41461
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Cite this article
  1. Wanqing Shao
  2. Sergio G-M Alcantara
  3. Julia Zeitlinger
(2019)
Reporter-ChIP-nexus reveals strong contribution of the Drosophila initiator sequence to RNA polymerase pausing
eLife 8:e41461.
https://doi.org/10.7554/eLife.41461
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Abstract

RNA polymerase II (Pol II) pausing is a general regulatory step in transcription, yet the stability of paused Pol II varies widely between genes. Although paused Pol II stability correlates with core promoter elements, the contribution of individual sequences remains unclear, in part because no rapid assay is available for measuring the changes in Pol II pausing as a result of altered promoter sequences. Here, we overcome this hurdle by showing that ChIP-nexus captures the endogenous Pol II pausing on transfected plasmids. Using this reporter-ChIP-nexus assay in Drosophila cells, we show that the pausing stability is influenced by downstream promoter sequences, but that the strongest contribution to Pol II pausing comes from the initiator sequence, in which a single nucleotide, a G at the +2 position, is critical for stable Pol II pausing. These results establish reporter-ChIP-nexus as a valuable tool to analyze Pol II pausing.

https://doi.org/10.7554/eLife.41461.001

Introduction

RNA polymerase II (Pol II) pausing is a key regulatory step in metazoan gene regulation. After initiating transcription at the transcription start site (TSS), Pol II often pauses 30–50 bp downstream of the TSS, before being released by p-TEFb into productive elongation (Adelman and Lis, 2012; Gaertner and Zeitlinger, 2014; Yamaguchi et al., 2013). Genome-wide profiling of Pol II and nascent transcripts suggest that pausing is widespread across the genome (Core et al., 2008; Muse et al., 2007; Nechaev et al., 2010; Zeitlinger et al., 2007), and inhibition of p-TEFb blocks the productive elongation of the majority of active genes (Chao and Price, 2001; Jonkers et al., 2014; Ni et al., 2008). Together, these results suggest that pause release is a general and obligatory step in transcription.

Despite evidence that pausing is a common feature of transcription, the stability of paused Pol II varies widely between genes. While the degree of pausing may be influenced by transcriptional activation (Buckley et al., 2014; Danko et al., 2013; Min et al., 2011; Rahl et al., 2010), differences in Pol II pausing across the genome are, to a large extent, a property of the promoter. In the early Drosophila embryo, Pol II pausing is most frequently found at developmental control genes, and a large portion of these genes remain highly paused throughout embryonic development independently of their expression (Gaertner et al., 2012; Zeitlinger et al., 2007). The promoter sequences of such highly paused genes are notably enriched for specific core promoter elements (Hendrix et al., 2008), and promoter swapping experiments demonstrate that Pol II pausing depends on the promoter sequence (Lagha et al., 2013). However, despite the evidence that Pol II pausing depends on the promoter sequence, how core promoter elements regulate Pol II pausing remains elusive.

Drosophila is ideally suited to study the relationship between promoter sequences and Pol II pausing since a large fraction of promoters in the Drosophila genome have clearly defined core promoter sequences that are associated with strong Pol II pausing (Chen et al., 2013; Gilchrist et al., 2010; Hendrix et al., 2008; Juven-Gershon and Kadonaga, 2010; Kwak et al., 2013; Lee et al., 1992; Vo Ngoc et al., 2017b). Moreover, these highly paused promoters typically undergo focused initiation, during which Pol II begins transcription at a defined genomic position within a window of a few nucleotides (Hoskins et al., 2011; Kwak et al., 2013; Ni et al., 2010). These promoters are easier to study than those with dispersed initiation, where Pol II initiation occurs throughout a genomic region of ~100 bp. Finally, these highly paused promoters do not typically show strong +1 nucleosomes near the pause site, making Pol II pausing less likely to be dependent on chromatin context (Benjamin and Gilmour, 1998; Bondarenko et al., 2006; Brown et al., 1996; Gaertner et al., 2012; Gilchrist et al., 2010; Izban and Luse, 1991; Jimeno-González et al., 2015; Kwak et al., 2013; Rach et al., 2011).

These convenient and easy-to-study highly paused promoters in Drosophila have features that are similar to those of mammalian promoters. Their core promoter elements are recognized by the basal transcription factor TFIID, a highly conserved multi-subunit complex whose promoter binding specificity appears to be conserved across metazoans. For example, recent cryo-EM structures of TFIID show human TFIID bound to all core promoter elements of the synthetic Drosophila super core promoter, including the sequences enriched among highly paused promoters (Louder et al., 2016; Patel et al., 2018). This suggests that the highly paused promoters in Drosophila have more elaborate sequence elements, yet rely on conserved metazoan transcription machinery for their function.

TFIID interacts with sequence elements at three regions along the core promoter, which are spaced in a fixed distance from each other (Juven-Gershon and Kadonaga, 2010; Louder et al., 2016; Patel et al., 2018; Vo Ngoc et al., 2017b). The first contact occurs at the TATA box, a well-studied core promoter element that is located ~30 bp upstream of the transcription start site and is bound by TATA-binding protein (TBP), a component of TFIID (Chasman et al., 1993; Kim et al., 1993a; Kim et al., 1993b). The second contact occurs at the initiator (Inr) sequence, which overlaps the nucleotide where Pol II initiates transcription (referred to as the +1) (Chalkley and Verrijzer, 1999; Emami et al., 1997). The Inr is the most common core promoter element in metazoans and is often the only identifiable core promoter element in mammalian promoters (Hoskins et al., 2011; Ohler et al., 2002; Smale and Kadonaga, 2003; Vo Ngoc et al., 2017a). The third DNA region contacted by TFIID lies ~30 bp downstream of the transcription start site. It contains short sequence motifs such as the downstream promoter element (DPE), motif ten element (MTE), and the pause button (PB) (Burke and Kadonaga, 1996; Hendrix et al., 2008; Kutach and Kadonaga, 2000; Lim et al., 2004; Purnell et al., 1994). Despite extensive studies on TFIID-promoter interactions, how core promoter elements affect Pol II pausing is not clear.

Measurements of paused Pol II across the genome, including with ChIP-seq, normally do not assess the duration of pausing directly, but rather represent steady-state Pol II occupancy across a population of cells, in which paused Pol II turns over at different rates. To measure the stability of paused Pol II more directly, a time-course analysis in response to triptolide treatment can be performed. Triptolide inhibits initiation globally and prevents new Pol II from reaching the pause position (Jonkers et al., 2014; Titov et al., 2011). After triptolide treatment, previously existing paused Pol II is lost over time, either by transitioning into elongation or due to premature transcript termination. The decay rate of paused Pol II under these conditions is then proportional to the stability of paused Pol II (Henriques et al., 2013; Jonkers et al., 2014; Krebs et al., 2017).

We recently performed such time-course measurements of paused Pol II across the Drosophila melanogaster genome using a high-resolution exonuclease-based ChIP-seq protocol (ChIP-nexus). This assay has the advantage of distinguishing Pol II signal at the site of initiation and pausing, and thus the Pol II half-life calculations are based on paused Pol II, rather than total Pol II at the promoter (Shao and Zeitlinger, 2017). These experiments confirmed the large differences in pausing stability across promoters and showed significant correlations with core promoter elements. However, whether these promoter sequences directly regulate Pol II pausing or whether they are statistically over-represented among promoters with high or low Pol II pausing stability is not known.

A better understanding of the relationship between the core promoter sequence and pausing behavior is difficult to obtain with current techniques. On one hand, traditional biochemical or reporter gene assays usually use gene expression and not Pol II occupancy as the readout since Pol II pausing was not yet known to be a general regulatory step in transcription when these assays were developed. As a result, in vitro assays for studying Pol II pausing are very limited, and it is not even clear whether Pol II pausing requires a natural chromatin context (Benjamin and Gilmour, 1998; Bondarenko et al., 2006; Brown et al., 1996; Izban and Luse, 1991). On the other hand, Pol II pausing is readily detected by genomics techniques in vivo, but manipulating endogenous genomic promoter sequences to analyze their links to Pol II pausing is time consuming. Therefore, having an assay that measures Pol II pausing outside the normal genomic context would allow quick promoter mutagenesis and greatly facilitate studying mechanistic features in the regulation of Pol II pausing.

Although traditional reporter assays were not designed to detect Pol II pausing, it is nevertheless possible that Pol II pausing occurs in some of these assays (Benjamin and Gilmour, 1998). We reasoned that if paused Pol II occurred on a plasmid, it would be possible to combine the efficiency of plasmid-based sequence mutagenesis with Pol II ChIP-nexus to rapidly test how alterations in promoter sequences affect Pol II pausing. Unexpectedly, we found that paused Pol II can easily be detected on a plasmid by ChIP-nexus and its footprint closely recapitulates the Pol II footprint at the endogenous locus. Using this novel assay, termed reporter-ChIP-nexus, we analyzed the contribution of various core promoter elements and discovered that the Inr sequence, when containing a G at position +2, plays an unexpectedly large and dominant role in stabilizing paused Pol II. Taken together, we show that reporter-ChIP-nexus combines the ease of plasmid-based sequence mutagenesis with high-resolution ChIP profiling, thereby serving as a valuable tool for dissecting the role of specific sequences in Pol II pausing.

Results

Promoter-specific Pol II pausing properties are recapitulated on the reporter

To test whether Pol II pausing is recapitulated on a plasmid, we used a GFP reporter, which has moderate expression when transfected into D. melanogaster Kc167 cells and allows rapid insertion of any type of promoter sequence (Figure 1A and Figure 1—figure supplement 1). After transient transfection of the reporter construct, the entire cell extracts were used for ChIP-nexus since there is no straightforward method for isolating plasmids from fixed cells before chromatin immunoprecipitation. Due to the high number of plasmids per cell, we found that only moderate sequence coverage (~5 to 10 million unique reads per sample) was sufficient to obtain high ChIP signal from the plasmid. Sequencing the endogenous genome in addition to the plasmid also provided a convenient internal control for the ChIP quality and for normalizing samples with each other.

Figure 1 with 2 supplements see all
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Reporter-ChIP-nexus captures paused Pol II.

(A) Reporter-ChIP-nexus is performed by cloning Drosophila pseudoobscura promoters or synthetic promoters into a simple GFP reporter and transfecting into D. melanogaster Kc167 cells. The whole cell lysate from cross-linked cells is used to perform Pol II ChIP-nexus. Exonuclease stop bases are then mapped and shown on the positive strand in red above the line, while reads from the negative end are shown in blue below the line. (B) Results of reporter-ChIP-nexus reveal strong Pol II pausing at the synthetic super core promoter (SCP), which contains the core promoter elements TATA, Inr, MTE, DPE and PB (top). The position of transcriptional initiation is mapped by sequencing the 5’ end of the produced RNA (bottom). The results for the SCP promoter show that the vast majority of RNAs start at the expected site of initiation.

https://doi.org/10.7554/eLife.41461.002

To explore whether Pol II pausing can be detected on this plasmid, we first cloned in the super core promoter (SCP) (Figure 1B and Table S1). This synthetic promoter contains correctly positioned TATA, Inr, MTE, DPE and PB core promoter elements such that they are recognized by TFIID and efficiently direct Pol II transcription (Juven-Gershon et al., 2006; Louder et al., 2016). Since it is a synthetic sequence, it can be unambiguously distinguished from the endogenous promoters in the genome. After transfection of the plasmid, we used ChIP-nexus to map Pol II on this promoter. Interestingly, we observed a strong accumulation of paused Pol II signal at the expected pausing position, suggesting that Pol II can pause on the plasmid (Figure 1B).

To validate that transcription on this promoter indeed begins at the initiator sequence and thus the accumulation of Pol II occurs downstream of transcription start site, we developed a gene-specific 5’ RNA sequencing protocol (Figure 2—figure supplement 2). This assay mapped the vast majority of the 5’ ends of the transcribed RNA to the initiator sequence (Figure 1B), confirming that the SCP functions as expected.

To test whether the Pol II pausing profile on the plasmids recapitulates the pattern of endogenous promoters, we then cloned promoter sequences from Drosophila pseudoobscura into the reporter. D. pseudoobscura promoters are sufficiently different from those in the melanogaster genome (due to ~25 millions of years of divergence), thereby facilitating the unambiguous mapping of Pol II signal on the plasmid.

Since the promoter sequences might have diverged in function between D. melanogaster and D. pseudoobscura, we first performed Pol II ChIP-nexus analysis on a D. pseudoobscura cell line (ML83-63) and analyzed the stability of paused Pol II after 1 hr triptolide treatment. After 1 hr triptolide treatment, promoters with a Pol II half-life of ~5–10 min show strongly reduced Pol II pausing at the pause position and often show increased levels of Pol II at the site of initiation as previously observed (Erickson et al., 2018; Krebs et al., 2017; Shao and Zeitlinger, 2017). In contrast, stably paused promoters maintain high levels of Pol II at the pausing position with no noticeable increase of Pol II at the site of initiation. This allowed us to identify promoters with strong differences in Pol II pausing for further analysis (Figure 2—figure supplement 1).

In total, we selected eight pseudoobscura promoters of ~250 bp in length that had a variety of known core promoter elements bound by TFIID (Table S1). We transfected each plasmid into D. melanogaster Kc167 cells and mapped the initiation start sites using gene-specific 5’ RNA sequencing (Figure 1—figure supplement 2). On all plasmids, the start sites mapped within a narrow window of a few base pairs to the predicted Inr sequence (Figure 2A and Figure 2—figure supplement 2).

Figure 2 with 4 supplements see all
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Reporter-ChIP-nexus recapitulates the endogenous Pol II pausing profile.

(A) To obtain the endogenous Pol II pattern of D. pseudoobscura promoters in the genome, ChIP-nexus was performed in a D. pseudoobscura cell line. Results are shown for the pepck, comm2 and pk promoters. The same promoters were then examined using reporter-ChIP-nexus in D. melanogaster Kc167 cells, which yielded patterns very similar to the endogenous Pol II profiles. The transcription initiation sites were confirmed by 5’ RNA sequencing. (B) To determine the stability of paused Pol II on the plasmid, transfected cells were treated with either DMSO (Control) or triptolide (TRI) for 1 hr. The results show a relative reduction in Pol II similar to that of the endogenous loci after treating the D. pseudoobscura cell line with TRI (see Figure 2—figure supplement 4). Therefore, reporter-ChIP-nexus reveals gene-specific Pol II pausing stability on a plasmid.

https://doi.org/10.7554/eLife.41461.005

We then performed Pol II ChIP-nexus on the transfected cells and compared the Pol II profile on each plasmid to the endogenous profile in D. pseudoobscura cells. Again, we detected strong Pol II signal on all plasmids, precisely at the canonical pausing position. Moreover, the Pol II profile was in most cases indistinguishable from the endogenous Pol II profile (Figure 2A and Figure 2—figure supplement 2), with Pearson correlations similar to or slightly below those of replicate experiments (Table S2 and S3). This indicates that lifting a promoter with TFIID-dependent core promoter elements out of its genomic chromatin context does not abolish Pol II pausing and confirms that the pausing profile at those promoters is highly dependent on the promoter sequence.

To test the versatility of our assay, we also tested a D. pseudoobscura promoter that is more likely dependent on the chromatin context. The promoter of the ribosomal gene RpL13A belongs to the group of promoters that uses TCT as initiator element (Parry et al., 2010). These promoters undergo focused initiation, but unlike other focused promoters, have a strong +1 nucleosome with high levels of H3K4me3 (Figure 2—figure supplement 3). We found that Pol II pausing can be recapitulated on the plasmid for RpL13A when a larger 2 kb region surrounding the core promoter was cloned into the plasmid, but not with a smaller 300 bp region (Figure 2—figure supplement 3). The plasmid with the larger region showed high levels of H3K4me3, similar to the endogenous Rpl13A promoter, while the construct with the smaller insertion did not (Figure 2—figure supplement 3). These results indicate that further technical optimization may be required for examining Pol II pausing on plasmids at promoters where genomic context is essential. This would be consistent with previous studies indicating that plasmids may or may not be correctly chromatinized when transfected into cells (Jeong and Stein, 1994; Reeves et al., 1985).

Focusing on the set of eight selected promoters, we next tested whether the stability of Pol II after triptolide treatment is recapitulated on the plasmid. To directly compare the Pol II ChIP-nexus profile on the plasmid between triptolide-treated and untreated cells, we used the same pool of plasmid-transfected cells for both samples and normalized them to each other using the genome reads. The results show that paused Pol II is indeed lost after treatment with triptolide and that the degree of loss is proportional to the stability of Pol II on the endogenous promoter. For example, some promoters (e.g. comm2) showed a strong reduction of paused Pol II after 1 hr triptolide treatment, while others (e.g pk) displayed minimal loss (Figure 2B and Figure 2—figure supplement 4). Taken together, these results suggest that Pol II not only pauses on the plasmid, but that the promoter-specific stability of paused Pol II can, to some extent, be measured on a plasmid.

The Pol II pausing stability can be altered by changing the downstream promoter sequence

Strong Pol II pausing correlates with the presence of downstream elements such as the PB, MTE and DPE (Chen et al., 2013; Gaertner et al., 2012; Gilchrist et al., 2010; Hendrix et al., 2008; Kwak et al., 2013; Shao and Zeitlinger, 2017). These sequences may be directly implicated in Pol II pausing since they are located near the site of paused Pol II, and changing the position of the DPE relative to the Inr alters Pol II pausing (Kwak et al., 2013). However, further functional evidence for their role in Pol II pausing is lacking, and a detailed mechanistic dissection of these downstream sequences using reporter-ChIP-nexus could be challenging. For example, sequences near the location of paused Pol II would have to be altered, which could indirectly affect the ChIP-nexus profile of Pol II, for example through altered protein-DNA crosslinking efficiency.

We therefore decided to broadly test the role of downstream sequences in Pol II pausing in our initial study. We took two promoters with a short Pol II pausing half-life (Act5C and pepck) and replaced the entire downstream promoter sequence with that of a stably paused promoter (pk or dve) (Figure 3A and Figure 3—figure supplement 1). We then performed Pol II ChIP-nexus on these hybrid promoters with or without triptolide treatment (we treated for 5 min since the wildtype promoters have a short Pol II half-live). While it was difficult to judge whether the wildtype and hybrid promoters had a different Pol II profile under control conditions, we found that a higher fraction of paused Pol II was reproducibly maintained on the hybrid promoters after triptolide treatment compared to the wild-type promoter (Figure 3B). Since the loss of Pol II at the pausing position after triptolide treatment reflects the pausing stability, these data suggest that the replacement of the downstream promoter sequence made Pol II pausing more stable.

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Changes in downstream promoter sequences alter paused Pol II stability.

(A) The pepck and Act5C downstream sequences were replaced with that from the stably paused promoter pk or dve (fusion site: 8 bp after the TSS). (B) Pol II ChIP-nexus data on the plasmids after transfection into Kc167 cells with or without treatment with triptolide (TRI) for 5 min. The wild-type pepck promoter shows a strong reduction of paused Pol II. At the pepck-pk-down and pepck-dve-down fusion promoters, the same TRI treatment did not reduce paused Pol II to the same extent as at the wild-type promoter, suggesting an increase in the paused Pol II stability as a result of changing the downstream promoter sequence. (C) To quantify the difference in paused Pol II stability between different constructs, the ratio of paused Pol II before and after TRI treatment is calculated using two biological replicates. The ratio for the wild-type promoter is then normalized to 1 (light green), and the relative change in this ratio for the fusion promoter is shown on the right (dark green). Error bars refer to the standard error of the mean (TRI treatment: 5 min for all promoters).

https://doi.org/10.7554/eLife.41461.010

To quantify the difference in Pol II pausing stability, we first calculated the relative loss of paused Pol II between control and triptolide-treated samples for each construct. Since the same batch of transfected cells are used for this comparison, and the total read counts are normalized to the genomic DNA, the relative loss of paused Pol II is constant and independent of transfection efficiency. Indeed, we found that these measurements were highly reproducible across biological replicates, although the reproducibility was somewhat lower with shorter treatment times of triptolide, presumably because small experimental fluctuations have a larger effect. To compare two constructs, we then determined the difference of triptolide-induced Pol II loss between the mutant construct and the wild-type counterpart. These calculations show that replacing the downstream region in our promoters increased the half-life of paused Pol II around 2-fold (Figure 3C).

Taken together, these results confirm that downstream promoter sequences indeed influence the stability of Pol II pausing and we will now refer to MTE, DPE and PB collectively as pausing elements. We next focused on the role of the other TFIID-bound regions in Pol II pausing.

An upstream region with a TATA box may reduce paused Pol II stability

The TATA box is often enriched among promoters with the lowest amount of Pol II pausing (Chen et al., 2013; Day et al., 2016; Gaertner et al., 2012; Shao and Zeitlinger, 2017), and there is evidence in mammalian cells that it promotes the release of paused Pol II (Amir-Zilberstein et al., 2007; Montanuy et al., 2008). However, some promoters, including the hsp70 promoter, contain a TATA box and yet show a strong pausing profile (Buckley et al., 2014; Gilchrist et al., 2010; Kwak et al., 2013), questioning the simple model of TATA promoting pause release.

To clarify the correlation between the presence of TATA or other core promoter elements and the stability of Pol II pausing, we first performed a genome-wide data analysis (Figure 4A and Figure 4—figure supplement 1). We scored each core promoter element based on the presence of consensus sequence at the expected promoter position, allowing for one mismatch: TATA box (STATAWAWR), Inr (TCAKTY), and pausing elements (CSARCSSA, KCGGTTSK or KCGRWCG). We then analyzed how their presence, alone or in pairwise combination, is correlated with the half-lives of paused Pol II that we measured previously (Shao and Zeitlinger, 2017).

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A TATA insertion may reduce Pol II pausing.

(A) Analysis of paused Pol II half-lives as a function of various core promoter element combinations. Median paused Pol II half-life of promoters with different combinations of core promoter elements (top) and the number of promoters classified (middle) are shown. A Wilcoxon rank sum test was then used to test whether the various promoter combinations have significantly different Pol II half-lives from each other (bottom). Core promoter elements other than those used for classification were allowed to occur in each group (e.g. Inr and pausing elements can occur at the 194 promoters with TATA box). Similar results were obtained when no promoter element other than those used for classification were allowed (mutually exclusive model Figure 4—figure supplement 1). (B) Experimental strategy to functionally test the role of TATA in Pol II pausing: the upstream promoter sequences of the paused promoter pk, comm2 and dve were replaced with that of the TATA promoter Act5C (fusion site: 7 bp before the TSS), or a canonical TATA box sequence (TATAAAA) was inserted at 31 bp upstream of the TSS. (C) The Pol II ChIP-nexus profiles for wild-type pk, the upstream region replacement (Act5C-up-pk) and the TATA insertion (TATA-pk) under control conditions (left) and after 1 hr treatment with triptolide (TRI, right) show that Pol II pausing is reduced after the pk promoter acquired a TATA box. (D) Quantification of the relative changes in paused Pol II stability for all fusion promoters relative to the wild-type promoters (TRI treatment: 1 hr for pk and dve based promoters, 40 min for comm2 based promoters). Note that only the pk promoter shows a clear reduction in Pol II pausing after insertion of a TATA box.

https://doi.org/10.7554/eLife.41461.012

As expected, promoters with a pausing element or an Inr tended to have a long Pol II half-life (median ~25 min) and promoters with both elements displayed an even longer half-life (median ~30 min), consistent with them functioning together (Figure 4A and Figure 4—figure supplement 1). Also consistent with previous findings (Chen et al., 2013; Day et al., 2016; Shao and Zeitlinger, 2017), any combination that contained a predicted TATA box showed a shorter median half-life of paused Pol II (medians 15–19 min). For example, the median half-life of promoters with a pausing element decreased from ~25 min to ~15 min in the presence of TATA (Wilcoxon p<10−4).

To test whether the TATA box robustly destabilized paused Pol II, we then replaced the entire upstream sequence of the TATA-less promoters pk, comm2 and dve with that of the TATA-containing promoter Act5C (Figure 4B). We reasoned that replacing a larger region will make it more likely that the added TATA box is functional within the larger promoter context. When we then analyzed the stability of Pol II pausing after triptolide treatment (1 hr for pk and dve, 40 min for comm2), we observed reduced Pol II in all constructs as compared to the wild-type promoter (Figure 4C and D and Figure 5—figure supplement 1). However, the effect was relatively small for the comm2 and dve promoter and only the pk promoter showed a strong, almost two-fold, reduction in paused Pol II.

To determine whether this effect was due to the TATA box, we then performed a much smaller alteration and replaced the 7 bp sequence located 31 bp upstream of the transcription start site with a canonical TATA-box sequence (TATAAAA) (Figure 4B). We found that in the pk promoter (but not the comm2 and dve promoter), this small change indeed reduced the stability of Pol II pausing, albeit to a lesser extent than when the entire upstream region was replaced (Figure 4C and D and Figure 5—figure supplement 1). This suggests that the TATA box may indeed play a role in destabilizing paused Pol II, but that the overall promoter context plays an important role, too.

The Inr strongly contributes to the stability of Pol II pausing

To better understand how the TATA box may promote pause release in some promoter contexts, we turned our attention to the Inr. The Inr functions synergistically with the TATA box in transcription if the two elements are optimally spaced from each other (Emami et al., 1997; Malecová et al., 2007; O'Shea-Greenfield and Smale, 1992; Xu et al., 2011), but on the other hand, the Inr is overall more highly enriched among highly paused genes (Gilchrist et al., 2010; Hendrix et al., 2008). This raises the question whether the Inr functions together with TATA in pause release or whether it plays a role in Pol II pausing together with the pausing elements.

To test whether an upstream TATA-containing region is dependent on the Inr, we took the dve promoter, which only weakly responded to the TATA-region replacement and swapped both the upstream region and the Inr with the sequences from the TATA-containing Act5C promoter (Figure 5A). Indeed, this replacement resulted in a strong, more than three-fold reduction in the stability of Pol II pausing (Figure 5B and C), suggesting that the type of Inr made a critical difference in this promoter context. However, even when we only replaced the Inr and not the upstream region in the dve promoter, we also observed a strong reduction in Pol II pausing stability (Figure 5B and C). These results suggest that the Inr sequence itself is an important determinant of Pol II pausing, even independently of the upstream TATA box.

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The initiator sequence plays an important role in Pol II pausing.

(A) Experimental series on the dve promoter testing the role of the Inr in conjunction with a TATA-containing region. Either only the upstream region was replaced with that of the Act5C promoter (fusion site 7 bp before the TSS), the upstream region and the Inr were replaced (fusion site 8 bp after the TSS), or only the Inr was replaced (16 bp region around the TSS). (B) Pol II ChIP-nexus profiles under control conditions and after treatment with triptolide (TRI) for 1 hr show a strong reduction of paused Pol II after replacing the Inr sequence. (C) Quantification of the relative changes in paused Pol II stability for all fusion promoters relative to the wild-type promoter (TRI treatment: 1 hr for all the promoters).

https://doi.org/10.7554/eLife.41461.014

Highly paused promoters and TATA promoters contain different Inr variants

The observation that the Inr sequence itself is important for pausing prompted us to perform a genome-wide search for Inr variants that may preferentially promote or destabilize Pol II pausing. For this purpose, we compared a strict set of naturally occurring TATA-containing promoters that have a relatively short paused Pol II half-life (<30 min) with those of stably paused promoters without a TATA box (half-life >= 60 min). The results revealed a significant difference in the Inr sequence between the two promoter types (Figure 6A and B). The Inr sequence surrounding the first transcribed base of stably paused promoters is best described by the motif TYAGTY (Figure 6B left).

Figure 6
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Initiator sequences differ between TATA and stably paused promoters.

(A) The sequence of stably paused promoters (132 randomly selected from the 490 promoters that did not have a TATA box and had a paused Pol II half-life longer than 60 min) are shown on the left as colored letters for a 100 bp window centered on the transcription start site (top). TATA promoters (132 promoters with paused Pol II half-lives shorter than 30 min) are shown on the right as comparison. A higher magnification of the initiator (Inr) sequences below shows clear differences between the two promoter types. (B) The consensus motif for the Inr sequences of stably paused promoters match more closely the canonical Inr consensus sequence (TCAKTY) and preferentially contains a G at the +2 position in most cases, whereas the Inr sequences of TATA promoters are more degenerate without enrichment of a G at the +2 position.

https://doi.org/10.7554/eLife.41461.016

In contrast, Inr sequences of TATA-containing promoters are more degenerate with frequent mismatches to the Inr consensus sequences (Figure 6B right). These mismatches may even occur at the A, which is the first transcribed base (the +1). However, the most striking difference is that the consensus Inr of stably paused promoters contains a G at the next position (the +2). Stably paused promoters contain a G in 90% of cases, while TATA-containing promoters have a G at this position in only 26% (p<10−46). This raises the possibility that the G at the +2 position of the Inr, which we refer to as Inr-G, plays an important role in stabilizing Pol II pausing.

The Inr-G variant plays a dominant role in stabilizing Pol II pausing

The discovery of the Inr-G variant prompted us to revisit our earlier analysis on how different combinations of promoter elements correlate with the half-life of paused Pol II. Strikingly, when we distinguished between Inr-G and Inr-nonG variants, the presence of the Inr-G variant correlated by far the strongest with Pol II pausing (Figure 7A and B and Figure 7—figure supplement 1). Promoters with an Inr-G variant had a median Pol II half-life of ~44 min, compared to ~14 min for promoters with the Inr-nonG variant (Wilcoxon p<10−48) or ~18 min for promoters with a pausing element (Wilcoxon p<10−24). When the Inr-G variant was found in combination with a pausing element, the median Pol II half-life was almost 60 min (compared to ~14 min for the Inr-nonG variant, Wilcoxon p<10−62). This suggests that while the pausing elements contribute to Pol II pausing, they strongly depend on the Inr-G variant, which overall has the strongest effect on Pol II pausing.

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The G at Inr + 2 position is critical for stable Pol II pausing.

(A) Analysis of paused Pol II half-lives as a function of core promoter element combinations after separating the Inr sequences into those that contain a G at position +2 (Inr-G) versus those that do not contain a G at this position (Inr-nonG). Median paused Pol II half-life (left), promoter numbers (right) and boxplot of their distribution (bottom, median in red) are shown for promoters with different combinations of core promoter elements. The promoters only contain the indicated promoter element, excluding any other promoter element (e.g. the 491 promoters with pausing elements and Inr-G variant do not contain TATA box). Similar results were obtained using a non-mutually exclusive model (Figure 7—figure supplement 1). Note that strong Pol II pausing was observed in all combinations that contain the Inr-G variant. Results from testing between combinations using the Wilcoxon rank test are shown in Table S4. (B) Experimental strategy to test the effect of mutating the G at Inr + 2 position of the Inr to T or A at the stably paused promoters dve, pk and the synthetic Super Core Promoter (SCP). (C) Pol II ChIP-nexus profiles after treatment with triptolide (TRI) for 1 hr show a strong reduction in Pol II pausing after the G was mutated at the dve promoter. (D) Quantification of the relative changes in paused Pol II stability for all fusion promoters relative to the corresponding wild-type promoter. Error bars are from replicate experiments. The duration of TRI treatment was 1 hr for pk and dve derived promoters, 30 min for SCP derived promoters.

https://doi.org/10.7554/eLife.41461.017

Interestingly, the presence of TATA was still correlated with shorter Pol II half-lives (median ~13 min), but the variant of Inr that was present made a strong difference (Figure 7A and Figure 7—figure supplement 1). While promoters with a combination of TATA and Inr-nonG had slightly shorter half-lives (median ~11 min), TATA in combination with the Inr-G variant showed very stable Pol II pausing (median ~59 min, Wilcoxon p<10−7). This suggests that the Inr acts dominantly over TATA in its effect on Pol II pausing, which explains our earlier observations that introducing a TATA upstream region alone only had a small effect on stably paused promoters.

These results strongly suggest that the G at the +2 position of the Inr is critical for stable Pol II pausing. To validate this experimentally, we specifically mutated the G into A or T at three stably paused promoters (dve, pk and the synthetic promoter SCP) and performed Pol II ChIP-nexus under control and triptolide treated conditions (1 hr treatment for dve and pk, 30-min treatment for SCP) (Figure 7B–D and Figure 7—figure supplement 2). Importantly, our mutations did not reduce the overall transcription levels (Figure 7—figure supplement 3), consistent with in vitro studies suggesting that the G is not important for transcription efficiency (Lo and Smale, 1996).

Strikingly, we observed a dramatic reduction in the stability of paused Pol II in the mutated constructs relative to the wild-type construct (Figure 7C and D). The dve promoter showed an almost five-fold decrease when mutated to either T or A. Even the pk and SCP promoters, which have more downstream pausing elements (Table S1) showed a ~ 2 fold reduction. These results show that the G in the Inr is indeed critical for stable Pol II pausing in Drosophila. More generally, this discovery demonstrates that reporter ChIP-nexus can be used to analyze the role of promoter sequences in Pol II pausing and uncover previously unknown roles of promoter elements.

Discussion

The Inr plays an important role in stable Pol II pausing

Genome-wide correlations have suggested a role for promoter sequences in Pol II pausing (Chen et al., 2013; Gaertner et al., 2012; Gilchrist et al., 2010; Hendrix et al., 2008; Nechaev et al., 2010; Shao and Zeitlinger, 2017), but functional data supporting a causal role for these sequences had been largely lacking. Here, we found that Pol II pausing is remarkably well recapitulated on a plasmid and that its stability can be measured on the plasmid by treatment with triptolide. By taking advantage of this assay, which we call reporter-ChIP-nexus, we were able to test the contribution of individual core promoter elements to Pol II pausing. We found that replacement of core promoter regions bound by TFIID tended to change the stability of paused Pol II in the expected direction, but the overall sequence context of the promoter was still important.

Most notably, we found that the Inr sequence played a strong role in the stability of Pol II pausing. This is surprising because the Inr sequence is thought to function synergistically with both the TATA box and the pausing elements (Juven-Gershon and Kadonaga, 2010), which have opposite effects on the stability of Pol II pausing. Identifying the preferred Inr sequences in promoters with a TATA box and with pausing elements resolved this conundrum. Promoters with stable Pol II pausing are highly enriched for Inr sequences resembling the consensus TYAGTY, which closely matches the Inr derived from functional studies (TCAKTY) (Hultmark et al., 1986; Purnell et al., 1994) and from computational analyses (TCAGTY and TCAGTT) (FitzGerald et al., 2006; Ohler et al., 2002; Stark et al., 2007). This in turn suggests that a strong consensus Inr sequence promotes Pol II pausing.

Promoters with a TATA box, in contrast, tend to have more degenerate Inr sequences. This explains why previous genomic sequence analyses have found these two elements to co-occur relatively infrequently in both Drosophila and human promoters (Chen et al., 2013; FitzGerald et al., 2006; Jin et al., 2006; Ohler et al., 2002; Vo Ngoc et al., 2017a), although biochemical experiments clearly show that they function synergistically in transcription (Emami et al., 1997; Malecová et al., 2007; O'Shea-Greenfield and Smale, 1992; Xu et al., 2011).

This difference in Inr sequence between promoter types is best reflected in the presence or absence of a G at the +2 position. Although G and T at this position are equally functional in transcription assays in vitro (Lo and Smale, 1996), stably paused promoters predominantly contain the Inr-G variant. Mutating this G to an A or T drastically reduced Pol II pausing in our assay, suggesting that the G is indeed important for stable Pol II pausing.

It is therefore possible that the degenerate Inr-nonG sequences in TATA-containing promoters represent a tradeoff. On one hand, these Inr sequences probably have sufficient sequence information to influence start site selection and promote TATA-dependent transcription. On the other hand, they may be weakened in their ability to stabilize Pol II pausing and thereby allow a mode of transcription with reduced Pol II pausing. Alternatively, TATA-containing promoters may function equally well with or without Pol II pausing, and therefore, the Inr sequences in these promoters may not have been under strong purifying selection during evolution.

Interestingly, an Inr-G variant (TCAGTT) was identified in comparative Drosophila genomics analyses as the most conserved Inr sequence (Stark et al., 2007). This suggests that the G variant may be under evolutionarily selection in paused promoters, consistent with Pol II pausing playing a critical role in keeping the promoter open (Gilchrist et al., 2008) and achieving coordinated gene expression during development (Lagha et al., 2013).

It is important to point out that while the G is conserved in Drosophila, human Inr sequences are often more degenerate and do not show a significant enrichment of G at the +2 (Carninci et al., 2006; Frith et al., 2008; Lo and Smale, 1996; Vo Ngoc et al., 2017a). However, there are also Inr variants with a much longer consensus sequence (Hendy et al., 2017; Yarden et al., 2009). It is therefore possible that human Inr variants also differentially affect Pol II pausing.

Promoter elements could be linked to Pol II pausing through TFIID

The promoter elements we tested here for Pol II pausing have traditionally been studied for their role in promoter recognition and transcription initiation by TFIID (Burley and Roeder, 1996; Lee and Young, 2000). TFIID is a large, flexible multi-subunit complex that can simultaneously bind to the TATA box, the Inr, and the pausing elements (Louder et al., 2016; Patel et al., 2018), thereby promoting the assembly of the transcription initiation complex. More recent evidence, however, suggests that TFIID has a function beyond initiation. For example, there is accumulating evidence that TFIID promotes Pol II re-initiation (Joo et al., 2017; Oelgeschläger et al., 1998; Yudkovsky et al., 2000; Zhang et al., 2015). Furthermore, genome-wide ChIP-nexus data show that TFIID binding extends beyond the pausing position in vivo (Shao and Zeitlinger, 2017), raising the intriguing possibility that TFIID plays a role in the coordination between consecutive rounds of Pol II initiation, pausing, and pause release.

If TFIID remains bound after initiation, it is possible that it influences the stability of Pol II pausing in a promoter-specific fashion. While TFIID can make contacts with all promoter elements on the super core promoter (Louder et al., 2016; Patel et al., 2018), natural promoters contain fewer promoter element consensus sequences, such that the strength and position of TFIID-promoter contacts may vary between promoters. We therefore speculate that the core promoter elements recognized by TFIID contribute to the dynamics of Pol II transcription by influencing how Pol II enters and remains in the pausing position (Figure 8). If TFIID binds tightly to the Inr and downstream promoter elements, Pol II may initiate and traverse the early transcribed region more slowly, and hence may be more likely to become stably paused (Figure 8A). In contrast, if TFIID is more tightly bound to the promoter upstream through the TATA box, then Pol II initiation and early transcription may occur in a less constrained fashion, which may promote pause release (Figure 8B). These tight DNA contacts in the presence of the TATA box is supported by biochemical experiments (Lee et al., 1991; Starr and Hawley, 1991), as well as more recent single-molecule footprinting studies (Krebs et al., 2017). In summary, although all core promoter elements may help recruit TFIID, they may differentially affect the passage of Pol II and its release into productive elongation, dependent on their location at the promoter.

Figure 8
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Model of how promoter sequences may influence Pol II pausing through TFIID Since the core promoter elements that we analyzed are bound by TFIID, we hypothesize that TFIID affects Pol II pausing dependent on the position it binds to the promoter.

(A)At stably paused promoters, pausing elements and consensus Inr sequences with a G at +2 position promote TFIID-downstream DNA interactions, which prolong Pol II pausing. (B)At TATA-containing promoters, the TATA box and a degenerate Inr sequence favor TFIID interactions with upstream DNA, which promote Pol II pause release.

https://doi.org/10.7554/eLife.41461.021

Future perspective

We found that reporter-ChIP-nexus is a useful method for studying the relationship between promoter sequence and Pol II pausing. This plasmid-based assay recapitulates the endogenous Pol II dynamics at the promoters tested and has sufficient sensitivity to identify changes in Pol II pausing upon promoter sequence alterations. The simplicity of the assay will no doubt be helpful for further studies on the roles of individual promoter sequences in Pol II pausing, including those of promoter types not studied here.

In order to perform a more high-throughput analysis on promoter sequences, some improvements in the assay will be beneficial. First, we currently sequence Pol II ChIP-nexus reads genome-wide in order to capture the reads from the plasmid. Any prior enrichment for plasmid sequences will therefore lower the cost of sequencing. Second, if genomic reads can no longer be reliably used as internal control, a control plasmid that monitors transfection efficiency would be useful. Finally, the assay will greatly benefit from automation. This would not only enable more high-throughput analyses but also improve the scope of the assay. Currently, the accuracy and reproducibility of the assay is higher for promoters with longer Pol II half-lives since longer triptolide treatment times reduce experimental fluctuations in handling and drug penetration. If automated, the assay would allow more tightly spaced time-course experiment in multiple replicates, which could more accurately distinguish smaller differences in Pol II half-lives.

Taken together, reporter-ChIP-nexus promises to open new possibilities for measuring the stability of Pol II pausing at individual promoter sequences in the future. Together with complementary approaches such as live imaging analyses, such studies will contribute to a better understanding of the relationship between promoter sequence, Pol II pausing, and the dynamic production of transcripts over time.

Materials and methods

Key resources table
Reagent type
(species) or
resource		DesignationSource or referenceIdentifiersAdditional
information
AntibodyRabbit polyclonal anti-Rpb3Julia Zeitlinger LabZeitlinger Lab #163185–5010 ug
AntibodyRabbit polyclonal anti-H3K4me3Cell Signaling#972710 ug
Strain, strain background (E. coli)NEB 5-alpha Competent E. coli (High Efficiency)New England Biolabs#C2987H
Strain, strain background (E. coli)One Shot ccdB Survival 2 T1R Competent CellsThermoFisher Scientific#A10460
ChemicalChloroformSigma-Aldrich#C2432
Commercial assay or kitCircLigase ssDNA LigaseIllumina (Epicentre)#CL4115K
Chemical compoundDMSOSigma-Aldrich#276855
Chemical compoundDynabeads Protein ALife Technologies#100-08D
Chemical compoundDynabeads Protein GLife Technologies#100-04D
Commercial assay or kitFastDigest BamHIThermoFisher Scientific#FERFD0054
Chemical compoundFuGENE HD reagentPromega#E2311
Chemical compoundHyClone SFX-Insect Cell Culture MediaThermoFisher Scientific#SH3027802PM
Commercial assay or kitKlenow Fragment (3’ to 5’ exo-)New England Biolabs#M0212S
Commercial assay or kitLambda ExonucleaseNew England Biolabs#M0262L
Chemical compoundOpti-MEM Reduced Serum MediumThermoFisher Scientific#31985062
Chemical compoundPhenol: Chloroform: iso-Amyl alcoholVWR#97064–824
Chemical compoundProtease Inhibitor Cocktail tablets EDTA-freeRoche Diagnostics Corporation#5056489001
Commercial assay or kitProteinase KLife Technologies#25530–049
Commercial assay or kitrecJNew England Biolabs#M0264L
Commercial
assay or kit		Restriction emzyme AfeINew England Biolabs#R0652S
Commercial assay or kitRestriction emzyme EcoRV-HFNew England Biolabs#R3195S
Commercial assay or kitRestriction emzyme SacI-HFNew England Biolabs#R3156S
Commercial assay or kitRNase AThermoFisher Scientific#EN0531
Commercial assay or kitRNase HNew England Biolabs#M0297S
Commercial assay or kitSuperScript II Reverse TranscriptaseThermoFisher Scientific#18064014
Commercial assay or kitT4 DNA PolymeraseNew England Biolabs#M0203S
Chemical compoundTriptolideTOCRIS Bioscience#3253
Chemical compoundTRIzol ReagentThermoFisher Scientific#15596026
Commercial assay or kitDirect-zol RNA MiniPrep kitGenesee#11–330
Commercial assay or kitFast SYBR Green Master mixLife Technologies#4385612
Commercial assay or kitGibson Assembly Master MixNew England Biolabs#E2611S
Commercial assay or kitHigh-Capacity RNA-to-cDNA KitThermoFisher Scientific#4387406
Commercial assay or kitIBI High Speed Plasmid Mini KitMIDSCI#IB47101
Commercial assay or kitNEBNext dA-tailing moduleNew England Biolabs#E6053L
Commercial assay or kitNEBNext end-repair moduleNew England Biolabs#E6050L
Commercial
assay or kit		NEBNext Multiplex Oligos for IlluminaNew England Biolabs#E7335S
Commercial assay or kitQ5High-Fidelity 2X Master MixNew England Biolabs#M0492L
Commercial assay or kitQ5 Site-Directed Mutagenesis KitNew England Biolabs#E0554S
Commercial assay or kitQuick Ligation KitNew England Biolabs#M2200L
Cell line (D. melanogaster)Kc167 cellsDGRC#1
Cell line (D. pseudoobscura)ML83-63 cellsDGRC#33
Sequence-based reagentChIP-nexus oligos(He et al., 2015)See Table S5
Sequence-based reagentGene-specific 5’ RNA sequencing oligosThis paperSee Table S6
Recombinant DNA reagentReporters used in this studyThis paperSee Table S7 and S8
Software, algorithmAnalysis codeGitHubhttps://github.com/zeitlingerlab/Shao_eLife_2019

Methods

Reporter construction

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The pAWG GFP reporter plasmid from the Drosophila Gateway cloning collection was used as the backbone for the reporter plasmid. The Act5C core promoter (−41 to 103 bp around the transcription start site, not including the upstream Act5C regulatory sequences, which is −2470 to −42 bp from the transcription start site) and the downstream Gateway cloning cassette (sequence between attR1 and attR2) were removed by digesting with the SacI and AfeI restriction enzymes. A 6 x UAS sequence from pUASp (as found in DGRC #1189) was inserted between the SacI and Afel restriction sites using Gibson Assembly Master Mix. The resulting plasmid pAWG-UAS has the Act5c upstream regulatory region, 6 x UAS, a GFP coding sequence and an EcoRV restriction site between the UAS sequences and the GFP coding region. Promoter sequences of interests were inserted into pAWG-UAS at the EcoRV cutting site with Gibson Assembly Master Mix. Subsequent mutations of core promoter sequences were performed with the Q5 site-directed site mutagenesis kit. For plasmid amplification and purification, the DNA was transformed into NEB 5-alpha Competent E. coli (High Efficiency) or One Shot ccdB Survival 2 T1R Competent Cells and purified with IBI High Speed Plasmid Mini Kit. Clones were validated with Sanger sequencing using the sequencing primer GCACCGTGACCATCACAGCATA. See Table S6 and S7 for detailed constructs descriptions and sequences.

Cell culture and transcription inhibitor treatment

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D. melanogaster Kc167 cells (DGRC #1) were grown in SFX media at 25°C. D. pseudoobscura ML83-63 cells (DGRC #33) were grown in M3 +BPYE + 10% FCS media at 25°C. Transcription inhibitors were added directly into culture media. Cells were treated with 500 μM Triptolide (TOCRIS Bioscience Cat. No. 3253 dissolved in DMSO) as done previously (Shao and Zeitlinger, 2017). Equivalent amounts of DMSO treatment (2% v/v) were used as control. To best capture the stability of paused Pol II, cells were treated with Triptolide for different time after transfecting with different reporter constructs. Refer to the main text and figure legends for the treatment duration.

ChIP-nexus

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For each ChIP-nexus experiment, 107 Kc167 cells or ML83-63 cells were fixed with 1% formaldehyde in culturing media at room temperature for 10 min with rotation. Fixed cells were washed with cold PBS, incubated with Orlando and Paro’s Buffer (0.25% triton X-100, 10 mM EDTA, 0.5 mM EGTA, 10 mM Tris-HCl pH 8.0, with freshly added Protease Inhibitor) for 10 min at room temperature with rotation, and then centrifuged and re-suspended in ChIP Buffer (10 mM Tris-HCl, pH 8.0; 140 mM NaCl; 0.1% SDS; 0.1% sodium deoxycholate; 0.5% sarkosyl; 1% Triton X-100, with freshly added Protease Inhibitor). Sonication was performed with a Bioruptor Pico for five rounds of 30 s on and 30 s off. Chromatin extracts were then centrifuged at 16000 g for 5 min at 4°C, and supernatants were used for ChIP.

To couple Dynabeads with antibodies, 50 µl Protein A and 50 µl Protein G Dynabeads were used for each ChIP-nexus experiment and washed twice with ChIP Buffer. After removing all the liquid, Dynabeads were resuspended in 400 µl ChIP Buffer. 10 µg antibodies were added, and tubes were incubated at 4°C for 2 hr with rotation. After the incubation, antibody-bound beads were washed twice with ChIP Buffer.

For chromatin immunoprecipitation, chromatin extracts were added to the antibody-bound beads and incubated at 4°C overnight with rotation and then washed with Nexus washing buffer A to D (wash buffer A: 10 mM Tris-EDTA, 0.1% Triton X-100, wash buffer B: 150 mM NaCl, 20 mM Tris-HCl, pH 8.0, 5 mM EDTA, 5.2% sucrose, 1.0% Triton X-100, 0.2% SDS, wash buffer C: 250 mM NaCl, 5 mM Tris-HCl, pH 8.0, 25 mM HEPES, 0.5% Triton X-100, 0.05% sodium deoxycholate, 0.5 mM EDTA, wash buffer D: 250 mM LiCl, 0.5% IGEPAL CA-630, 10 mM Tris-HCl, pH 8.0, 0.5% sodium deoxycholate, 10 mM EDTA). End repair and dA-tailing were performed using the NEBNext End Repair Module and the NEBNext dA-Tailing Module. ChIP-nexus adaptors with mixed fixed barcodes (CTGA, TGAC, GACT, ACTG) were ligated with Quick T4 DNA ligase and converted to blunt ends with Klenow fragment and T4 DNA polymerase. The samples were treated with lambda exonuclease and RecJf exonuclease for generating Pol II footprints at high resolution. After each enzymatic reaction, the chromatin was washed with the Nexus washing buffers A to D and Tris buffer (10 mM Tris, pH 7.5, 8.0, or 9.5, depending on the next enzymatic step).

After RecJf exonuclease digestion, the chromatin was eluted and subjected to reverse crosslinking and ethanol precipitation. Purified single-stranded DNA was then circularized with CircLigase, annealed with oligonucleotides complementary to the BamHI restriction site and linearized by BamHI digestion. The linearized single-stranded DNA was purified by ethanol precipitation and subjected to PCR amplification with NEBNext High-Fidelity 2X PCR Master Mix and ChIP-nexus primers. The ChIP-nexus libraries were then gel-purified before sequencing with Illumina NextSeq 500.

Reporter-ChIP-nexus

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To transform cells with the reporter plasmid, 1.5 × 107 Kc167 cells were diluted in 10 ml SFX media and seeded into a 10 cm cell culture dish. A mixture of 2.5 µg reporter plasmids, 20 µl FuGENE HD reagent and 500 µl Opti-MEM Reduced Serum Medium was then added to the cell culture. Transfected cells were cultured for 48 hr at 25°C to allow proper expression of the reporter. ChIP-nexus was performed on the whole cell extracts from transfected cells following standard ChIP-nexus procedure as described above. After sequencing, reporter-ChIP-nexus samples were aligned to the combined genome of dm3 and the specific reporter construct. Only reads that uniquely aligned to the reporter sequence were used for analysis and PCR duplicates with the same ChIP-nexus barcode were removed.

Reporter expression quantification

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Expression of the reporter was quantified by calculating the expression of GFP RNA from the reporter over the expression of the endogenous RNA from the RpL30 locus, after normalizing for the difference in copy number between the reporter and the endogenous genome.

For cDNA preparations, 5 × 105 transfected cells were transferred into 1.5 ml tubes and washed with cold PBS. After removing all liquid, cells were lysed and incubated with 500 µl TRIzol Reagent at room temperature for 5 min. Total RNA was then extracted with 100 µl Chloroform and purified with a Direct-zol RNA MiniPrep kit. cDNA was generated with a High-Capacity RNA-to-cDNA Kit.

For DNA preparations, 5 × 105 transfected Drosophila Kc167 cells were transferred to 1.5 ml tubes and washed with cold PBS. After removing all liquid, cells were lysed with 50 µl ChIP buffer. Cell lysates were incubated with 150 Elution buffer (50 mM Tris pH 8.0, 10 mM EDTA, 1% SDS), 100 µl TE buffer (50 mM Tris pH 8.0, 1 mM EDTA) and 4 µl RNAse A for 30 min at 37°C. Then, 2 µl Protease K was added to the mixture and incubated at 65°C for 2 hr. The DNA was then purified with an ethanol precipitation. qPCR on both the cDNA and DNA was performed using Fast SYBR Green Master Mix and primers against GFP and RpL30. The relative expression of the reporter was calculated using the following equations.

GFPexpression=2Ct(GFPDNA)−Ct(GFPcDNA)
RpL30expresssion=2Ct(RpL30DNA)−Ct(RpL30cDNA)
GFPrelativeexpression=(GFPexpression)/RpL30expression

For this analysis, cDNA was prepared from two to three biological replicates (cells were transfected and processed on different dates) using 5 × 105 transfected cells each. Two technical replicates (the same biological sample were split into two and processed side by side) were performed during qPCR.

Gene-specific 5’ RNA sequencing

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This method is similar to 5’ RNA amplification of cDNA ends (RACE) but uses reagents and steps from ChIP-nexus. Briefly, total RNA was isolated from 5 × 105 transfected cells as above, and cDNA was generated from the GFP RNA using SuperScript II Reverse Transcriptase and a primer against GFP that has partial TruSeq P5 sequences at the 5’ end (GFP_reversetx_primer). The cDNA was purified by treatment with RNAse A and RNAse H at room temperature for 30 min, followed by ethanol precipitation. The cDNA circularization, cutting and PCR amplification was then performed as in the ChIP-nexus protocol (thus without rolling circle amplifications), except that a new BamHI cutting oligo (reversetx_BamHI_cutting) was annealed before BamHI cutting, and a barcoded P7-GFP fusion primer (reversetx_barcode_primer_#1) was used in the PCR. After sequencing with TruSeq primers, the reads were aligned to the dm3-reporter combined genome, and the first bases of the P5 sequenced reads were recorded as the stop bases of the reverse transcriptase and correspond to the 5’ ends of the GFP RNA from the reporter plasmid.

Quantification and statistical analysis

Normalization of Pol II signal on the plasmid

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Due to variations in the transfection efficiency, absolute Pol II signal on different plasmids is not comparable. As a result, paused Pol II stability needs to be accessed by comparing Pol II signal under control and triptolide-treated conditions. Because control and triptolide-treated samples come from the same pool of transfected cells, they have the same transfection efficiency and the same plasmid/Kc167 genome ratio. Therefore, Pol II signal on the plasmids under control and triptolide-treated conditions can be directly compared after normalizing for read counts.

The changes in Pol II signal after triptolide treatment is calculated in the following way:

  1. Each sample is first normalized to reads/millions to account for differences in read coverage.

  2. For each sample, the total Pol II signal is calculated in a 301 bp window around the promoter region on the plasmid (termed Total_Pol_sig).

  3. The change in Pol II signal on the plasmid in response to triptolide treatment is then calculated as the ratio between Total_Pol_sig from the triptolide-treated sample over Total_Pol_sig from the corresponding DMSO-treated control sample.

It is likely that this method underestimates the changes of Pol II signal between control and triptolide-treated condition since the total occupancy of Pol II is slightly reduced after triptolide treatment (Shao and Zeitlinger, 2017), yet we normalize both samples to equal read counts. However, since the relative loss of Pol II after triptolide treatment is similar across all samples in the Kc167 genome, the relative differences in the above calculated ratio between different plasmids is preserved. For this analysis, two biological replicates (cells were transfected and processed on different dates) were performed using 107 transfected cells each.

Correlation between paused Pol II half-lives and core promoter elements

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In Figure 4A and Figure 4—figure supplement 1, promoters with previously measured paused Pol II half-life (Shao and Zeitlinger, 2017) were separated into groups with either one or two of the following core promoter elements: TATA box (STATAWAWR), Inr (TCAKTY) and pausing elements (CSARCSSA, KCGGTTSK or KCGRWCG). Only motif matches at the expected canonical position and with up to one mismatch were allowed (Table S1). In a mutually exclusive model (Figure 4—figure supplement 1), promoters with TATA-box and Inr cannot contain pausing elements, while in a non-mutually exclusive model (Figure 4A), pausing elements are allowed.

The same method was applied to Figure 7A,B (using a mutually exclusive model) and Figure 7—figure supplement 1 (using a non-mutually exclusive model), except that promoters were further separated into containing Inr-G or Inr-nonG variants based on whether a G is present at the +2 position.

Sequence analysis of TATA promoters and stably paused promoters

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TATA promoters and stably paused promoters were defined using the length of paused Pol II half-life (Shao and Zeitlinger, 2017) and the presence or absence of the TATA box sequence (STATAWAWR with up to one mismatch) at 40 to 20 bp upstream of the transcription start site. TATA promoters have the TATA box sequence and paused Pol II half-lives shorter than 30 min, whereas stably paused promoters lack a detectable TATA box sequence and have paused Pol II half-lives longer than 60 min. In Drosophila Kc167 cell, 132 promoters were defined as TATA promoters and 490 promoters were defined as stably paused promoters.

In Figure 5A, DNA sequences at TATA promoters (n = 132) and stably paused promoters (n = 132, randomly selected from the original 490 promoters) were obtained from the dm3 genome and represented as heatmap. The consensus motif for the Inr sequence in Figure 5B was generated using the R package seqLogo.

Data and software availability

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Raw and processed data associated with this manuscript are deposited in GEO under the accession number GSE116244 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE116244).

All data analysis performed in this paper, including raw data, processed data, software tools, and analysis scripts, have been reproduced in a publicly accessible Linux virtual machine. Instructions for accessing the virtual machine can be found at http://research.stowers.org/zeitlingerlab/data.html. The analysis code is available on GitHub at https://github.com/zeitlingerlab/Shao_eLife_2019 (Shao, 2019 ; copy archived at https://github.com/elifesciences-publications/Shao_eLife_2019).

Data availability

Raw and processed data associated with this manuscript are deposited in GEO under session number GSE116244. All data analysis performed in this paper, including raw data, processed data, software tools, and analysis scripts, have been reproduced in a publicly accessible Linux virtual machine. Instructions for accessing the virtual machine can be found at http://research.stowers.org/zeitlingerlab/data.html. The analysis code is available on GitHub at https://github.com/zeitlingerlab/Shao_eLife_2019 (copy archived at https://github.com/elifesciences-publications/Shao_eLife_2019).

The following data sets were generated
    1. Shao W
    2. Alcantara SG-M
    3. Zeitlinger J
    (2019) NCBI Gene Expression Omnibus
    ID GSE116244. Reporter-ChIP-nexus reveals strong contribution of the Drosophila initiator sequence to RNA polymerase pausing.
    https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE116244
The following previously published data sets were used
    1. Shao W
    2. Zeitlinger J
    (2017) NCBI Gene Expression Omnibus
    ID GSE85741. Paused RNA polymerase II inhibits new transcriptional initiation.
    https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE85741

References

    1. O'Shea-Greenfield A
    2. Smale ST
    (1992)
    Roles of TATA and initiator elements in determining the start site location and direction of RNA polymerase II transcription
    The Journal of Biological Chemistry 267:6450.

Decision letter

  1. Karen Adelman
    Reviewing Editor; Harvard Medical School, United States
  2. James L Manley
    Senior Editor; Columbia University, United States

In the interests of transparency, eLife includes the editorial decision letter and accompanying author responses. A lightly edited version of the letter sent to the authors after peer review is shown, indicating the most substantive concerns; minor comments are not usually included.

Thank you for sending your article entitled "Reporter-ChIP-nexus reveals contribution of initiator sequence to RNA Polymerase II pausing" for peer review at eLife. Your article has been evaluated by three peer reviewers, one of whom is a member of our Board of Reviewing Editors, and the evaluation has been overseen by Aviv Regev as the Senior Editor.

Summary:

In this work, the Zeitlinger lab uses ChIP-nexus (a high-resolution form of ChIP-seq developed in the Zeitlinger lab) combined with a reporter system to study the contribution of promoter DNA sequence to the stability of paused polymerase. Using D. pseudoobscura promoters in the reporter plasmid in a D. melanogaster genomic background, they show that initiation, pausing, and the stability of pausing are recapitulated on the plasmids. This is important validation of the usefulness of the system. Promoter swapping and targeted mutagenesis of reporter promoters then yielded three main results: 1) the downstream region of promoters was important for pause stability, 2) a TATA-box is associated with pause instability, and 3) a strong initiator consensus with a G at +2 was critical for pause stability.

The reviewers all found the ChIP-nexus technique to be powerful, with the potential for high throughput screens with this strategy. However, as presented here, the assay is very low throughput. Incredibly, most of the authors conclusions are drawn from a single perturbation of single test promoter sequence. The work herein is thus considered preliminary, and requires additional validation prior to publication.

In short, the very limited, highly context-dependent data presented in this manuscript reduces enthusiasm for both the assay and the strength of the authors' conclusions. If concerns can be addressed with additional experiments, however, the reviewers are happy to consider a revised manuscript.

Essential revisions:

1) The authors must test each of the promoter mutations in several promoter contexts to validate findings. We recommend that every mutation be tested in at least 3 different promoter backgrounds to allow for reasonable confidence in conclusions.

2) The drug triptolide has been used by a number of groups to measure the stability of paused RNAPII and all data thus far converges towards a median half-life of paused RNAPII on the order of around 5-10 min. Genes that have polymerase remaining at 1 h of triptolide treatment are thus extreme outliers. As such, it is a shame that the authors have selected this type of hyper-stable promoter for use in their assay, as it makes it very difficult- if not impossible- to extrapolate from this extreme case to normal promoter behavior. In the absence of experiments on a more typical promoter, the reviewers are concerned that the results presented here will not be broadly applicable. Thus, when additional promoters are tested, we strongly recommend that the authors select promoters with decay rates more in line with the average gene, and use a more relevant time point for triptolide treatment, on the order of 5-20 min.

3) To better support the authors claims, they should perform a genomic analysis with their existing data on the stability of pausing at promoters with various combinations of the TATA and Inr elements. They already have the melanogaster data from Kc cells +/- triptolide and should have plenty of sequencing depth given the number of plasmids analyzed. Thus, in theory, this would simply require analysis of the genomic reads without additional experimentation. This would serve to corroborate the results and generalize the model to more promoters.

4) The claim that the reporter assay recapitulates the genomic assay could also use strengthening. This point is supported by 8 genes in the current manuscript, which is probably not enough for statistically rigorous conclusions. Figure 2—figure supplement 2 shows the 8 examples, and there is quite some variability in them. The question is how much variability, how to quantify it, and whether or not conclusions can be made given this variability. Given that a lot of conclusions are drawn from comparing the shapes of these promoter distributions, I would think the extent to which they can be reliably made is important. dve, the gene selected for the majority of mutational tests, is an example that shows differences between genomic and reporter contexts.

One idea is to bin the promoters into, say 10-20bp bins, and compare the normalized read counts across the bins for the reporter and genomic assay. Then the mean squared error or the correlation coefficient can be computed comparing the reporter and the genomic assay, and the degree of variability can be measured across more genes. This would at least give us some context for how much variability can be expected, and if the differences observed due to the mutations are significantly more than that baseline difference. This analysis or something similar is considered critical.

5) The novelty of finding a strong initiator at highly paused genes is severely over-exaggerated and previous literature is inadequately cited. The connection between Initiator consensus elements, highly focused transcription initiation and high levels of paused RNAPII has appeared multiple times in the literature previously, at least as far back as 2008 (Hendrix et al., 2008), and also in 2010 (Gilchrist et al., 2010).

Further, pausing has been recapitulated on DNA sequences in vitro, thus we already know that DNA sequence is a major contributor to pausing. See many in vitro studies from the Gilmour lab (some cited) and also Adelman and Lis (2005).

Finally, the authors mention in the Discussion that functional data supporting a role for promoter sequences and pausing are missing. While they are correct that there is a dearth of these important functional studies, the Lis lab did a functional analysis pause elements in the Drosophila Hsp70 promoter, by inserting sequences that changed the position of these elements. See Kwak et al., 2013. Given that there are so few studies like this and the level of detail covering other aspects of the literature, I think this experiment is worth discussing in this context.

It doesn't take anything away from the authors' accomplishments to adequately cite previous research in a scholarly way. Appropriate citations must be added.

6) Also related to the ambiguity in the number of promoters involved in each analysis: In the section "The Inr strongly contributes to the degree of Pol II pausing", I think it would be enhanced if the text listed the number of promoters examined for each group. For example, the text says "We first analyzed the naturally occurring Inr sequences from TATA-containing promoters versus those of stably paused promoters". I would like to know when reading this how many promoters are in each set leading to the observations e.g. that TATA-containing promoters have fewer Gs at the +2 position. In particular, when reading the caption for Figure 5A, it says n=132, and this implied that the other panels also used 132 promoters. However I expect that the other panels were based on a larger set (at least panel B). Ideally, more than 132 promoters would have been used for this analysis. Overall, added clarity on the number of promoter for Figure 5B and elsewhere would be helpful.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Thank you for submitting your article "Reporter-ChIP-nexus reveals strong contribution of initiator sequence to RNA Polymerase II pausing" for consideration by eLife. This evaluation has been overseen by a Reviewing Editor and James Manley as the Senior Editor.

The reviewers have discussed the reviews with one another and the Reviewing Editor has drafted this decision to help you prepare a revised submission.

Summary:

This revised version of the manuscript "Reporter-ChIP-nexus reveals strong contribution of initiator sequence to RNA Polymerase II pausing" is improved over the previous version as concerns the number of promoters and mutations analyzed. However, the way in which the data is discussed is still lacking in clarity, the caveats to interpretation are still not made clear, and the citation of prior literature is still lacking.

Thus, the manuscript is not determined to be acceptable in its current format. However, at this point the main criticisms of the work can all be addressed with text changes. Thus, we invite a resubmission if such changes are comprehensively made.

Essential revisions:

1) As the authors acknowledge in the Introduction, pausing takes place at all (or nearly all) genes. The duration and stability of pausing varies, but pausing appears to be an obligate part of the transcription cycle. Thus, the statement in the Abstract that "a G at the +2 position, is critical for Pol II pausing", is not accurate. The authors should change this to clarify that they mean 'stable pausing' or something of this sort. But a G and +2 is certainly not critical for the fundamental process of pausing itself.

2) The authors should note early in the Results section that 1h Triptolide treatment is a very long treatment, and that secondary effects are likely to occur during such extended intervals of transcription inhibition. Most labs use 2-20 minutes of Triptolide treatment to avoid such potential artifacts. We appreciate that the authors must rely on very long treatments to overcome the considerable noise in their assay, but this should be acknowledged as a potential source of artifact, and as a potential limitation to this study. This idea is briefly mentioned in the Discussion, but it must be noted in the Results section to make readers aware of this caveat.

3) The Bentley lab has recently questioned the interpretation of Triptolide data (in particular that using ChIP-based strategies such as in Shao et al. 17), noting that at some genes the Triptolide-mediated block to initiation traps Pol II at the TSS. This effect is evident at the Pino and pepck genes in this study. The authors should cite the recent work from the Bentley lab, and explain whether their ChIP-nexus assay gives them the spatial resolution needed to separate paused from un-initiated Pol II. This will help place the current work in a proper context, and will be clarifying for the field.

4) Based on the limited number of experiments aimed at testing the role of downstream promoter sequences, and the limitations in interpreting such experiments, the authors should be more circumspect in the conclusions drawn from these studies. We find it to be an over-reach that the authors conclude that downstream sequences affect pausing less than expected. Given the lack of data to this point, it is safest to note that this wasn't rigorously evaluated, and leave it at that.

5) We understand that the authors went into this study expecting the TATA box to elicit a strong effect on pausing, and that they were surprised that this is not the case. However, the large amount of inconclusive data presented on this topic, and the discussion thereof, is confusing. If the main conclusion is that TATA doesn't seem to have a strong effect on pause stability, and that any effect is highly context dependent and easily over-ridden by other sequences, then there is a much more straightforward way to present this. We strongly suggest that the authors simplify the presentation of the TATA data to focus on the main conclusion, rather than enumerating all the inconclusive or contradictory findings they obtained.

6) Given that computational assays have routinely defined the Inr consensus to have a G at +2, should this really be called the Inr-G variant? Instead, we recommend that this be considered the consensus. It should be clarified that this is the 'norm' and not something that deviates from the norm.

7) The discussion of pausing in its genomic context versus on a plasmid at the beginning of the Discussion is difficult to understand in light of the literature.

First, there have been a number of studies demonstrating that stable pausing does not involve downstream nucleosomes. The clearest of these demonstrations was Li and Gilmour, EMBO J, 2103). This paper should be cited and the authors should edit the first paragraph to reflect this work, and the fact that it is not at all surprising that pausing could take place on a plasmid or outside of the chromatin context.

8) It was demonstrated many years ago that pausing is rarely inhibitory in its endogenous context, because of the positive effect of the paused polymerase on maintaining open chromatin (Gilchrist G and D 2008). In this study, it was specifically demonstrated that the positive effect of pausing was clear in the endogenous context, but did not occur on a plasmid. Thus, it has been previously established that the effect of pausing on gene output is different on a plasmid than in the genome, and that any minor inhibitory effect seen on a plasmid is NOT borne out in the endogenous locus. Thus, it is perplexing that the authors claim that pausing is inhibitory based on their reporter assay- since this is known to be an inaccurate read-out of endogenous activity. Thus, the comments in the Discussion suggesting that the +2 G version of Inr would be inhibitory to transcription because it promotes pausing are unfounded and should be removed.

https://doi.org/10.7554/eLife.41461.029

Author response

Essential revisions:

1) The authors must test each of the promoter mutations in several promoter contexts to validate findings. We recommend that every mutation be tested in at least 3 different promoter backgrounds to allow for reasonable confidence in conclusions.

We agree and have now tested 9 additional promoters (2 for downstream pausing elements, 4 for upstream TATA regions, 2 for Inr mutations, 1 ribosomal gene promoter). In addition to these experiments, we have dissected the role of various promoter combinations computationally in more detail and found that these data strengthen and refine our conclusions.

As expected, we found that the downstream pausing elements stabilize pausing and that addition of a TATA box may reduce pausing. Interestingly though, the effect of TATA is not as strong and consistent as expected, and this can be explained by the strong effect of the Inr sequence (with a G at position +2). Mutating the G at the Inr on the other hand strongly reduces Pol II pausing. Consistent with such strong role in pausing, the type of Inr variant has the strongest correlation with the half-life of paused Pol II among all analyzed elements, an analysis we now added. Therefore, we have not only added additional experiments to support our conclusions, but we have strengthened our finding that the Inr plays an important role in Pol II pausing.

2) The drug triptolide has been used by a number of groups to measure the stability of paused RNAPII and all data thus far converges towards a median half-life of paused RNAPII on the order of around 5-10 min. Genes that have polymerase remaining at 1 h of triptolide treatment are thus extreme outliers. As such, it is a shame that the authors have selected this type of hyper-stable promoter for use in their assay, as it makes it very difficult- if not impossible- to extrapolate from this extreme case to normal promoter behavior. In the absence of experiments on a more typical promoter, the reviewers are concerned that the results presented here will not be broadly applicable. Thus, when additional promoters are tested, we strongly recommend that the authors select promoters with decay rates more in line with the average gene, and use a more relevant time point for triptolide treatment, on the order of 5-20 min.

Apart from the stably paused promoters, we have tested the Pol II profile on plasmids using promoters with a half-life in D. mel. of 5 (Act5C), 10 (pino) and 10 (pepck) min, respectively, which are in line with “average” promoters. However, in order to detect significant differences in half-lives between hybrid promoters, we have to use the more stable paused promoters. This is because the accuracy and reproducibility of the assay is higher for promoters with longer half-lives. For promoters with shorter half-lives, we have to use shorter drug treatment times. As a result, small fluctuations in drug treatment time and drug penetration can produce considerable variability in the results. To accurately measure a half-life in the order of 5-20 min, we would need to perform tightly spaced time-course experiment in many replicates. In the future, this might be possible as we plan to automate the assay and reduce sequencing cost by isolating plasmid DNA. However, with the current low-throughput method, using sequences from stably paused promoters is a more feasible approach for obtaining clear and interpretable results.

In fact, this is one of the reasons we pioneered this assay in Drosophila. Drosophila has a much larger fraction of promoters with clearly defined core promoter sequences that are associated with stable Pol II pausing. While pausing may not be as stable in mammalian genomes, the role of core promoter elements and the interaction between core promoter elements and TFIID is nevertheless conserved. For example, the excellent Cryo-EM structures of TFIID derived by Eva Nogales (Louder et al., 2016, Patel et al., 2018) is human TFIID on the Drosophila super core promoter (which includes “pausing elements” such as DPE and MTE). For this reason, we believe that the insights obtained from stably paused promoters will be broadly relevant.

We have now extensively rewritten the Introduction to better explain our reasoning behind the choice of promoters. In short, they are easy to study and due to known interactions with TFIID, are relevant for mammalian promoters. Finally, we point out current limitations of the assay in our Discussion at the end.

3) To better support the authors claims, they should perform a genomic analysis with their existing data on the stability of pausing at promoters with various combinations of the TATA and Inr elements. They already have the melanogaster data from Kc cells +/- triptolide and should have plenty of sequencing depth given the number of plasmids analyzed. Thus, in theory, this would simply require analysis of the genomic reads without additional experimentation. This would serve to corroborate the results and generalize the model to more promoters.

We liked the idea of performing a more combinatorial analysis of the promoter elements with regard to the paused Pol II half-lives, and this approach turned out to be very fruitful. While we had previously analyzed the genome-wide relationship between core promoter elements and paused Pol II half-lives (see Shao et al., 2017), we had not specifically looked at how various combinations correlate with Pol II pausing. This new analysis now revealed that the Inr sequence with a G at the +2 position most strongly correlates with pausing, independently of which other core promoter elements are found in the promoter.

We have now added the results from this new analysis to Figure 4A and Figure 7A with additional information found in Figure 4—figure supplement 1, Figure 7—figure supplement 1 and Table S6 in Supplementary file 1.

4) The claim that the reporter assay recapitulates the genomic assay could also use strengthening. This point is supported by 8 genes in the current manuscript, which is probably not enough for statistically rigorous conclusions. Figure 2—figure supplement 2 shows the 8 examples, and there is quite some variability in them. The question is how much variability, how to quantify it, and whether or not conclusions can be made given this variability. Given that a lot of conclusions are drawn from comparing the shapes of these promoter distributions, I would think the extent to which they can be reliably made is important. dve, the gene selected for the majority of mutational tests, is an example that shows differences between genomic and reporter contexts.

One idea is to bin the promoters into, say 10-20bp bins, and compare the normalized read counts across the bins for the reporter and genomic assay. Then the mean squared error or the correlation coefficient can be computed comparing the reporter and the genomic assay, and the degree of variability can be measured across more genes. This would at least give us some context for how much variability can be expected, and if the differences observed due to the mutations are significantly more than that baseline difference. This analysis or something similar is considered critical.

We had a supplementary table (Table S3 in Supplementary file 1) that showed the correlation coefficient between the bp of the Pol II profiles from the plasmid versus those of the genomic region. They range from 0.77 to 0.96 (mean 0.92), which is very high given that we are not even binning the data into 10-20 bp bins (binning would presumably smooth the data and give higher correlations).

To strengthen this point even further, we have now in the revised manuscript compared these correlation coefficients to those obtained between biological replicate experiments on the plasmids (Table S2 and S3 in Supplementary file 1). Overall, we found the correlations between plasmid and genomic context to be only slightly lower than those between replicates (on average 0.86 versus 0.92), suggesting that the plasmid recapitulates the genomic Pol II pausing profile remarkably well despite possible differences in chromatin or regulatory context. We have added this result to the main text (subsection “Promoter-specific Pol II pausing properties are recapitulated on the reporter”, fourth paragraph), as well.

5) The novelty of finding a strong initiator at highly paused genes is severely over-exaggerated and previous literature is inadequately cited. The connection between Initiator consensus elements, highly focused transcription initiation and high levels of paused RNAPII has appeared multiple times in the literature previously, at least as far back as 2008 (Hendrix et al., 2008), and also in 2010 (Gilchrist et al., 2010).

Further, pausing has been recapitulated on DNA sequences in vitro, thus we already know that DNA sequence is a major contributor to pausing. See many in vitro studies from the Gilmour lab (some cited) and also Adelman and Lis (2005).

Finally, the authors mention in the Discussion that functional data supporting a role for promoter sequences and pausing are missing. While they are correct that there is a dearth of these important functional studies, the Lis lab did a functional analysis pause elements in the Drosophila Hsp70 promoter, by inserting sequences that changed the position of these elements. See Kwak et al., 2013. Given that there are so few studies like this and the level of detail covering other aspects of the literature, I think this experiment is worth discussing in this context.

It doesn't take anything away from the authors' accomplishments to adequately cite previous research in a scholarly way. Appropriate citations must be added.

The statistical association between the Inr and Pol II pausing has been described before, by ourselves in collaboration with Mike Levine’s lab (Hendrix 2008) and in Gilchrist et al., 2010 (shown in Figure 2—figure supplement 2, not described in main text). However, this association could have been indirect since another pausing element, the DPE, requires the Inr to function properly (see review by Kadonaga, Dev Biol 2012). Due to the possible indirect effect, while the association between DPE (or PB) and pausing is more evident, the role of the Inr has been much less clear. In addition, the role of the Inr in Pol II pausing is further complicated by the observation that the Inr functions synergistically with the TATA box, which is associated with reduced Pol II pausing.

Regardless of how one interprets these statistical associations, a causal role in Pol II pausing had not been adequately shown for any core promoter element. We believe our results are novel and significant as we provide strong evidence that core promoter elements impact Pol II pausing individually. To the best of our knowledge, such direct functional examinations have not been done before (the spacing experiments by the Lis lab are interesting but do not address which promoter element mediates the observed effects). Furthermore, in the revised manuscript, we not only provide evidence for a direct role of the Inr in Pol II pausing, but we also show that statistically speaking, the G in the Inr plays a dominant role in Pol II pausing, which is quite surprising.

To provide more clarity for the reader, we have now rewritten the manuscript to better describe what was (or was not) known about the Inr, while citing the requested papers (Introduction, subsection “The Inr strongly contributes to the degree of Pol II pausing”, and subsection “The Inr plays an important role in Pol II pausing”). This should help the reader better understand the significance of our finding.

6) Also related to the ambiguity in the number of promoters involved in each analysis: In the section "The Inr strongly contributes to the degree of Pol II pausing", I think it would be enhanced if the text listed the number of promoters examined for each group. For example, the text says "We first analyzed the naturally occurring Inr sequences from TATA-containing promoters versus those of stably paused promoters". I would like to know when reading this how many promoters are in each set leading to the observations e.g. that TATA-containing promoters have fewer Gs at the +2 position. In particular, when reading the caption for Figure 5A, it says n=132, and this implied that the other panels also used 132 promoters. However I expect that the other panels were based on a larger set (at least panel B). Ideally, more than 132 promoters would have been used for this analysis. Overall, added clarity on the number of promoter for Figure 5B and elsewhere would be helpful.

We had previously described this promoter selection in detail in the Materials and methods under subsection“Sequence analysis of TATA promoters and stably paused promoters”. To make it easier for the reader, we now describe the selection of the promoters briefly in the main text (subsection “The Inr strongly contributes to the degree of Pol II pausing”) and the Figure 5 legend. Furthermore, we perform a more general analysis of the combinatorial relationship between core promoter elements and pausing half-life (Figure 4A and 7A), which places the above-mentioned figure into context.

[Editors' note: further revisions were requested prior to acceptance, as described below.]

Essential revisions:

1) As the authors acknowledge in the Introduction, pausing takes place at all (or nearly all) genes. The duration and stability of pausing varies, but pausing appears to be an obligate part of the transcription cycle. Thus, the statement in the Abstract that "a G at the +2 position, is critical for Pol II pausing", is not accurate. The authors should change this to clarify that they mean 'stable pausing' or something of this sort. But a G and +2 is certainly not critical for the fundamental process of pausing itself.

We agree: in fact, in two out of four sentences we had said “critical for stable Pol II pausing”. We have now changed the remaining two sentences also to “critical for stable pausing”.

2) The authors should note early in the Results section that 1h Triptolide treatment is a very long treatment, and that secondary effects are likely to occur during such extended intervals of transcription inhibition. Most labs use 2-20 minutes of Triptolide treatment to avoid such potential artifacts. We appreciate that the authors must rely on very long treatments to overcome the considerable noise in their assay, but this should be acknowledged as a potential source of artifact, and as a potential limitation to this study. This idea is briefly mentioned in the Discussion, but it must be noted in the Results section to make readers aware of this caveat.

We appreciate the concern about side effects, which we share when using drug treatments. However, we have not observed concerning side effects of treating our cells with triptolide for 60 min (see Shao and Zeitlinger, 2017) and note that many investigators have used such extended treatments:

- The Bentley lab paper cited below used 10-60 min;

- The Shilatifard lab paper (Chen et al.) used a time-course of up to 120 min and performed most of their further analysis on the 60 min timepoint. Like ours, their control gels on polymerase abundance and phosphorylation show no detectable side effects;

- The John Lis lab paper (Jonkers et al.) used 12.5 min, 25 min and 50 min timepoints.

We did not mean to convey the impression in our last response letter that our assay has “considerable noise”. As can be seen from the small error bars in the figures, biological replicates are very reproducible, even when treating for 5 min (Figure 3). Shorter triptolide treatment times may have more noise than longer treatment times, but this could be addressed by performing more replicates. In other words, using 60 min triptolide treatment was not born out of necessity, but a strategic choice, since it allowed us to choose promoters with maximal differences in Pol II pausing stability (and we had not encountered variability or other concerning side effects with 60 min treatment times).

To be sure, we now mention the better accuracy for longer Pol II half-lives in the Results section: “… we found these measurements to highly reproducible across biological replicates, although the reproducibility is somewhat lower with shorter treatments times of triptolide, presumably because small experimental fluctuations have a larger effect.”

3) The Bentley lab has recently questioned the interpretation of Triptolide data (in particular that using ChIP-based strategies such as in Shao et al. 17), noting that at some genes the Triptolide-mediated block to initiation traps Pol II at the TSS. This effect is evident at the Pino and pepck genes in this study. The authors should cite the recent work from the Bentley lab, and explain whether their ChIP-nexus assay gives them the spatial resolution needed to separate paused from un-initiated Pol II. This will help place the current work in a proper context, and will be clarifying for the field.

We agree that it is curious that this paper seems to suggest that the long Pol II half-lives we observe are due to us measuring Pol II at the initiation site. As we clearly describe in our paper, the high resolution of ChIP-nexus allows us to distinguish paused Pol II from Pol II at the initiation site (Shao and Zeitlinger, 2017). While we observe increased Pol II at the site of initiation after triptolide treatment (this is not a novel finding in the Bentley lab paper), we are specifically measuring the exponential decay of paused Pol II at the site of Pol II pausing to determine the half-life of paused Pol II (the position of pausing is determined based on Pol II pausing after flavopiridol treatment).

To make this clear in the present paper, we have made the following changes:

- Introduction: “We recently performed such time-course measurements of paused Pol II across the Drosophila melanogaster genome using a high-resolution exonuclease-based ChIP-seq protocol (ChIP-nexus). This assay has the advantage that it distinguishes between Pol II at the site of initiation and pausing, and thus the Pol II half-life calculations are based on paused Pol II, rather than total Pol II at the promoter (Shao and Zeitlinger, 2017).”

- When we first describe the Pol II profiles after triptolide treatment, we now point out the increased Pol II at the initiation site and cite our paper and the Bentley lab paper: “After 1 hour triptolide treatment, promoters with a Pol II half-life of ~5-10 min show strongly reduced Pol II pausing at the pause position and often show increased levels of Pol II at the site of initiation as previously observed (Shao and Zeitlinger, Erickson et al., 2018). In contrast, stably paused promoters maintain high levels of Pol II at the pausing position with no noticeable increase of Pol II at the site of initiation.”

4) Based on the limited number of experiments aimed at testing the role of downstream promoter sequences, and the limitations in interpreting such experiments, the authors should be more circumspect in the conclusions drawn from these studies. We find it to be an over-reach that the authors conclude that downstream sequences affect pausing less than expected. Given the lack of data to this point, it is safest to note that this wasn't rigorously evaluated, and leave it at that.

We have now removed this section from the Discussion.

5) We understand that the authors went into this study expecting the TATA box to elicit a strong effect on pausing, and that they were surprised that this is not the case. However, the large amount of inconclusive data presented on this topic, and the discussion thereof, is confusing. If the main conclusion is that TATA doesn't seem to have a strong effect on pause stability, and that any effect is highly context dependent and easily over-ridden by other sequences, then there is a much more straightforward way to present this. We strongly suggest that the authors simplify the presentation of the TATA data to focus on the main conclusion, rather than enumerating all the inconclusive or contradictory findings they obtained.

We have now extensively revised the text for clarity and have taken out any unnecessary commentary or discussion of the effect of TATA (we also removed TATA from the Abstract). Instead, we focused on the role of the Inr.

6) Given that computational assays have routinely defined the Inr consensus to have a G at +2, should this really be called the Inr-G variant? Instead, we recommend that this be considered the consensus. It should be clarified that this is the 'norm' and not something that deviates from the norm.

That is a good question, but based on biochemical studies, the Drosophila Inr consensus motif is TCAKTY, where K stands for T or G, and both the T and G at +2 are equally functional. Furthermore, the G is not over-presented among mammalian promoters, suggesting that the G is not the better nucleotide for Pol II initiation per se. The reason computational studies have predominantly found the G variant is presumably because the Inr-G variant is less degenerate, easier to identify and shows increased conservation. Therefore, we believe it is important to point out and emphasize the presence of the G at the +2 position and feel that it conveys the information more precisely.

7) The discussion of pausing in its genomic context versus on a plasmid at the beginning of the Discussion is difficult to understand in light of the literature.

First, there have been a number of studies demonstrating that stable pausing does not involve downstream nucleosomes. The clearest of these demonstrations was Li and Gilmour, EMBO J, 2103). This paper should be cited and the authors should edit the first paragraph to reflect this work, and the fact that it is not at all surprising that pausing could take place on a plasmid or outside of the chromatin context.

Actually, there are many papers supporting a role for nucleosomes in Pol II pausing, but we agree that this is not a resolved issue and the field is quite split. Since this is not an important point in the paper, we therefore decided to take out this entire section from the Discussion.

8) It was demonstrated many years ago that pausing is rarely inhibitory in its endogenous context, because of the positive effect of the paused polymerase on maintaining open chromatin (Gilchrist G and D 2008). In this study, it was specifically demonstrated that the positive effect of pausing was clear in the endogenous context, but did not occur on a plasmid. Thus, it has been previously established that the effect of pausing on gene output is different on a plasmid than in the genome, and that any minor inhibitory effect seen on a plasmid is NOT borne out in the endogenous locus. Thus, it is perplexing that the authors claim that pausing is inhibitory based on their reporter assay- since this is known to be an inaccurate read-out of endogenous activity. Thus, the comments in the Discussion suggesting that the +2 G version of Inr would be inhibitory to transcription because it promotes pausing are unfounded and should be removed.

We have removed this part of the Discussion as well.

https://doi.org/10.7554/eLife.41461.030

Article and author information

Author details

  1. Wanqing Shao

    Stowers Institute for Medical Research, Kansas City, United States
    Present address
    Department of Genetics, The Edison Family Center for Genome Sciences & Systems Biology, Washington University School of Medicine, St Louis, United States
    Contribution
    Conceptualization, Software, Formal analysis, Validation, Investigation, Visualization, Methodology, Writing—original draft, Writing—review and editing.
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-4234-2095

  2. Sergio G-M Alcantara

    Stowers Institute for Medical Research, Kansas City, United States
    Contribution
    Conceptualization, Validation, Investigation, Methodology, Writing—review and editing.
    Competing interests
    No competing interests declared
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0003-3695-4677

  3. Julia Zeitlinger

    1. Stowers Institute for Medical Research, Kansas City, United States
    2. Department of Pathology and Laboratory Medicine, University of Kansas Medical Center, Kansas City, United States
    Contribution
    Conceptualization, Data curation, Formal analysis, Supervision, Funding acquisition, Validation, Investigation, Visualization, Methodology, Writing—original draft, Project administration, Writing—review and editing.
    For correspondence
    jbz@stowers.org
    Competing interests
    J.Z. owns a patent on ChIP-nexus. EP3234199A1
    ORCID icon "This ORCID iD identifies the author of this article:" 0000-0002-5172-3335

Funding

Stowers Institute for Medical Research

  • Julia Zeitlinger

The funders had no role in study design, data collection and interpretation, or the decision to submit the work for publication.

Acknowledgements

We thank our colleagues Joan Conaway, Robb Krumlauf and Kaelan Brennan at the Stowers Institute for Medical Research, Mounia Lagha (Institut de Génétique Moléculaire de Montpellier) and James Kadonaga (UC San Diego) for comments on the manuscript. The work was funded by the Stowers Institute for Medical Research and has been done to fulfill, in part, requirements for Wanqing Shao’s PhD thesis research as a student of the Graduate School of Stowers Institute for Medical Research and for Sergio Garcia-Moreno’s PhD thesis research as a student registered with the Open University.

Senior Editor

  1. James L Manley, Columbia University, United States

Reviewing Editor

  1. Karen Adelman, Harvard Medical School, United States

Publication history

  1. Received: August 28, 2018
  2. Accepted: April 4, 2019
  3. Version of Record published: April 25, 2019 (version 1)

Copyright

© 2019, Shao et al.

This article is distributed under the terms of the Creative Commons Attribution License, which permits unrestricted use and redistribution provided that the original author and source are credited.

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  1. Wanqing Shao
  2. Sergio G-M Alcantara
  3. Julia Zeitlinger
(2019)
Reporter-ChIP-nexus reveals strong contribution of the Drosophila initiator sequence to RNA polymerase pausing
eLife 8:e41461.
https://doi.org/10.7554/eLife.41461

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Further reading

    1. Chromosomes and Gene Expression
    2. Genetics and Genomics

    Decapping factor Dcp2 controls mRNA abundance and translation to adjust metabolism and filamentation to nutrient availability

    Anil Kumar Vijjamarri, Xiao Niu ... Alan G Hinnebusch
    Research Article

    Degradation of most yeast mRNAs involves decapping by Dcp1/Dcp2. DEAD-box protein Dhh1 has been implicated as an activator of decapping, in coupling codon non-optimality to enhanced degradation, and as a translational repressor, but its functions in cells are incompletely understood. RNA-Seq analyses coupled with CAGE sequencing of all capped mRNAs revealed increased abundance of hundreds of mRNAs in dcp2Δ cells that appears to result directly from impaired decapping rather than elevated transcription. Interestingly, only a subset of mRNAs requires Dhh1 for targeting by Dcp2, and also generally requires the other decapping activators Pat1, Edc3 or Scd6; whereas most of the remaining transcripts utilize NMD factors for Dcp2-mediated turnover. Neither inefficient translation initiation nor stalled elongation appears to be a major driver of Dhh1-enhanced mRNA degradation. Surprisingly, ribosome profiling revealed that dcp2Δ confers widespread changes in relative translational efficiencies (TEs) that generally favor well-translated mRNAs. Because ribosome biogenesis is reduced while capped mRNA abundance is increased by dcp2&Delta, we propose that an increased ratio of mRNA to ribosomes increases competition among mRNAs for limiting ribosomes to favor efficiently translated mRNAs in dcp2Δ cells. Interestingly, genes involved in respiration or utilization of alternative carbon or nitrogen sources are up-regulated, and both mitochondrial function and cell filamentation are elevated in dcp2Δ cells, suggesting that decapping sculpts gene expression post-transcriptionally to fine-tune metabolic pathways and morphological transitions according to nutrient availability.

    1. Chromosomes and Gene Expression
    2. Genetics and Genomics

    Mouse B2 SINE elements function as IFN-inducible enhancers

    Isabella Horton, Conor J Kelly ... Edward B Chuong
    Research Article Updated

    Regulatory networks underlying innate immunity continually face selective pressures to adapt to new and evolving pathogens. Transposable elements (TEs) can affect immune gene expression as a source of inducible regulatory elements, but the significance of these elements in facilitating evolutionary diversification of innate immunity remains largely unexplored. Here, we investigated the mouse epigenomic response to type II interferon (IFN) signaling and discovered that elements from a subfamily of B2 SINE (B2_Mm2) contain STAT1 binding sites and function as IFN-inducible enhancers. CRISPR deletion experiments in mouse cells demonstrated that a B2_Mm2 element has been co-opted as an enhancer driving IFN-inducible expression of Dicer1. The rodent-specific B2 SINE family is highly abundant in the mouse genome and elements have been previously characterized to exhibit promoter, insulator, and non-coding RNA activity. Our work establishes a new role for B2 elements as inducible enhancer elements that influence mouse immunity, and exemplifies how lineage-specific TEs can facilitate evolutionary turnover and divergence of innate immune regulatory networks.

    1. Biochemistry and Chemical Biology
    2. Chromosomes and Gene Expression

    An acetylation-mediated chromatin switch governs H3K4 methylation read-write capability

    Kanishk Jain, Matthew R Marunde ... Brian D Strahl
    Short Report Updated

    In nucleosomes, histone N-terminal tails exist in dynamic equilibrium between free/accessible and collapsed/DNA-bound states. The latter state is expected to impact histone N-termini availability to the epigenetic machinery. Notably, H3 tail acetylation (e.g. K9ac, K14ac, K18ac) is linked to increased H3K4me3 engagement by the BPTF PHD finger, but it is unknown if this mechanism has a broader extension. Here, we show that H3 tail acetylation promotes nucleosomal accessibility to other H3K4 methyl readers, and importantly, extends to H3K4 writers, notably methyltransferase MLL1. This regulation is not observed on peptide substrates yet occurs on the cis H3 tail, as determined with fully-defined heterotypic nucleosomes. In vivo, H3 tail acetylation is directly and dynamically coupled with cis H3K4 methylation levels. Together, these observations reveal an acetylation ‘chromatin switch’ on the H3 tail that modulates read-write accessibility in nucleosomes and resolves the long-standing question of why H3K4me3 levels are coupled with H3 acetylation.