When a gene is expressed, the DNA is first transcribed to produce an intermediate molecule called a messenger RNA (mRNA), which is then translated to produce a protein. RNA Polymerase II is an enzyme that makes mRNA molecules in organisms as diverse as plants, animals and yeast.

RNA Polymerase II is a complex made of a number of proteins. The largest protein in this complex includes a ‘carboxy-terminal domain’ that has multiple repeats of seven amino acids one after the other. The first amino acid in each repeat, a tyrosine, is referred to as tyrosine-1. Adding various chemical tags to the amino acids in these repeats co-ordinates the steps involved in the transcription of genes. In yeast, for example, adding a phosphate groups to tyrosine-1 seems to help the polymerase to proceed to make long mRNA molecules. However, it is not known what these chemical tags do in humans or other animals.

Now Hsin et al. (and independently Descostes, Heidemann et al.) have shown that the same phosphate groups on tyrosine-1 perform functions in vertebrates (animals with backbones) that are different to those performed in yeast. These functions include protecting the carboxy-terminal domain from being broken down inside cells, and transcribing the DNA that is upstream of genes.

Hsin et al. replaced tyrosine-1 in RNA Polymerase II from chicken cells with a related amino acid that cannot have phosphate groups added to it. This mutant RNA Polymerase II was unstable and degraded by the molecular machinery in cells that breaks down damaged or unneeded proteins back into amino acids. Hsin et al. also compared the mRNA molecules that are made by the wild-type RNA Polymerase II with those produced by a related mutant. This comparison revealed an unexpected accumulation of RNA molecules that are transcribed in the opposite direction from mRNAs. These RNA molecules, known as ‘upstream antisense RNAs’, have been described only recently. And while the function of these RNAs remains mysterious, the results of Hsin et al. suggest that tyrosine-1 helps to ensure that these RNA molecules are rapidly broken down.

The results of Hsin et al. raise a number of important questions, and foremost among these questions is: how do these newly discovered properties of tyrosine-1 contribute to the control of gene expression in animals? Further work is needed to answer this question.

RNA polymerase II (RNAP II) is a multisubunit enzyme responsible in eukaryotes for transcription of all mRNAs and many non-coding RNAs. Rpb1, the largest subunit, contains a unique C-terminal domain (CTD), composed of up to 52 heptad repeats with the consensus sequence Tyr¹-Ser²-Pro³-Thr⁴-Ser⁵-Pro⁶-Ser⁷ (YSPTSPS). The CTD performs multiple functions during transcription (Buratowski, 2009; Munoz et al., 2010; Egloff et al., 2012; Hsin and Manley, 2012; Heidemann et al., 2013), most of which are dependent on phosphorylation of specific CTD residues. For example, Ser5 phosphorylation (Ser5-P) promotes recruitment of capping enzyme (Fabrega et al., 2003) and Ser2-P can be important for 3′ mRNA processing (Kim et al., 2004). Ser7-P and Thr4-P also function in 3′ processing, of snRNAs (Egloff et al., 2007) and histone mRNAs (Hsin et al., 2011), respectively, with the later also functioning in transcription elongation (Hintermair et al., 2012). Tyr1 can be phosphorylated in mammals (by c-Abl; Baskaran et al., 1993) and yeast (Mayer et al., 2012), where it may prevent premature recruitment of termination factors. However, the function(s) of Tyr1 and Tyr1-P in metazoans are unknown.

We previously utilized chicken DT40 cells to study properties of the Rpb1 CTD. We showed that an Rpb1 derivative containing a CTD with 26 YSPTSPS repeats (Rpb1-26r) plus the ten C-terminal non-consensus residues, important for stability (Chapman et al., 2005), confers cell viability, while a comparable derivative with all Thr4 residues changed to Val was inviable (Hsin et al., 2011). To investigate the functions of Tyr1, we constructed a plasmid encoding a Flag-tagged Rpb1 derivative, Rpb1-Y1F, identical to Rpb1-26r but with all Tyr1 residues replaced by Phe, and expressed this in Rpb1 conditional knock-out cells (DT40-Rpb1; Hsin et al., 2011). Tyr1 was suggested to be essential for viability in S. cerevisiae (West and Corden, 1995), but not in S. pombe (Schwer and Shuman, 2011). To determine whether Tyr1 is required for growth in vertebrate cells, DT40-Rpb1 cells were transfected with the Rpb1-Y1F vector, and tetracycline (tet) was added to turn off wild-type Rpb1 expression. Rpb1-Y1F was unable to complement Rpb1, whereas Rpb1-26r fully restored viability (Figure 1—figure supplement 1A).

We next established cell lines stably expressing Rpb1-Y1F to analyze how the Y1F mutation affects Rpb1 function. Cells expressing Rpb1-Y1F (Y1F) stopped growing around 24 hr in medium containing tet (Figure 1A). To examine whether the inviability of Y1F cells might result from different Rpb1 levels, we analyzed several independent Y1F cell lines by Western blot (WB) with anti-FLAG antibodies (Abs). Rpb1-Y1F levels were indeed significantly reduced compared to Rpb1-26r (Figure 1B). Importantly, accumulation of a lower molecular weight form (indicated by *) was observed in all Y1F cell lines. This corresponds to a derivative likely precisely lacking the CTD, as it migrated slightly more rapidly than an Rpb1 derivative containing six heptads (Figure 1B).

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To begin to investigate the basis for Rpb1-Y1F instability, we determined how many Tyr1 residues were necessary to restore stability. We first analyzed an Rpb1-Y1F derivative (20F+6Y) in which the F residues in the C-terminal six heptads were reverted to Y, and found that this derivative was completely stable (Figure 1—figure supplement 1B), although cells expressing Rpb1-20F+6Y remained inviable (Figure 1—figure supplement 1A). Next, we analyzed an Rpb1-Y1F derivative in which only a single F, in the C terminal-most heptad, was changed back to Y (Rpb1-25F+Y). Strikingly, this single Tyr residue was sufficient to prevent Rpb1 degradation, as the truncated isoform, which we denote Rpb1-b, was absent, and Rpb1-25F+Y levels were comparable to Rpb1-26r in multiple 25F+Y cell lines (Figure 1C; quantitation of the amount of degraded Rpb1 observed in multiple experiments is shown in Figure 1—figure supplement 1C). However, despite the restoration of Rpb1 stability, 25F+Y cells remained inviable (Figure 1D).

We next set out to determine how Tyr1 residues stabilize Rpb1. A first question was whether Rpb1 is indeed Tyr1-phosphorylated in DT40 cells. To address this, we utilized an anti-phospho-Tyr1 Ab (Mayer et al., 2012) to examine Tyr1 phosphorylation (Tyr1-P) of Rpb1-25F+Y and Rpb1-26r by WB; both proteins were indeed Tyr1-phosphorylated (Figure 2A). We next investigated where in cells the Rpb1-b isoform accumulates. We analyzed cytoplasmic, nuclear and chromatin-bound fractions from 26r and Y1F cells by WB with an N-terminal Rpb1 Ab (N20). Rpb1-b (indicated by *) was detected in all three fractions from Y1F cells, but barely or not at all in the 26r fractions (Figure 2B). The relative (and absolute) Rpb1-b levels were lowest in the cytoplasm, while Rpb1-b was essentially the only form on Y1F chromatin. As anticipated, Rpb1-b was not detected in 25F+Y cell fractions (Figure 2—figure supplement 1A). We next determined whether Tyr1-P could also be detected on Rpb1 in all three fractions, in this case using extracts from wild-type DT40 (Figure 2C) and human HEK293 (Figure 2—figure supplement 1B) cells. Robust Tyr1-P was indeed detected in all three fractions in both cell types. Notably, in both cytoplasm and nucleoplasm, Tyr1-P was observed only on hypophosphorylated Rpb1 (the lower band), while it was found primarily on the hyperphosphorylated isoform on chromatin. This suggests both that CTD phosphorylation is limited to Tyr1 in the cytoplasm and nucleoplasm and that Tyr1-P is present on hyperphosphorylated RNAP II found on active genes. We also examined phosphorylation on Ser 2, 5 and 7 and Thr4 (Figure 2C, Figure 2—figure supplement 1B). All these modifications were nearly undetectable in cytoplasmic and nuclear fractions, present almost exclusively on chromatin-associated, hyperphosphorylated Rpb1. Together, our data show that Tyr1, and only Tyr1, is phosphorylated before RNAP II engages in transcription, and support the idea that Tyr1-P functions in stabilizing the CTD when RNAP II is not transcribing, and perhaps also plays a role during transcription. Consistent with this, Tyr1-P was detected on Rpb1 immunoprecipitated by Abs recognizing Ser5-P and Ser2-P (Figure 2—figure supplement 1C).

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We next wished to determine how the CTD is degraded, and whether Tyr1-P indeed plays a role. Since one Tyr residue in the final heptad confers stability, it is unlikely that endoproteolytic cleavage occurred between the CTD and the Rpb1 body. One possibility is that the CTD is degraded by the proteasome, which has been shown to associate both with RNAP II and with active genes (Gillette et al., 2004). Additionally, certain naturally unstructured proteins can be degraded by the proteasome in a ubiquitin-independent manner (Sheaff et al., 2000; Tofaris et al., 2001). Since the CTD is considered a structure-less domain (Meinhart et al., 2005), we hypothesized that the CTD is a natural proteasome substrate, and that Tyr1-P prevents its proteasomal degradation. To test this directly, we performed in vitro proteasome assays using a GST-CTD substrate (containing full-length wt CTD) and purified 20S proteasomes, and detected products by WB. Using an anti-GST Ab, the amount of full-length GST-CTD was greatly diminished and a ladder-like pattern of low molecular weight bands was detected, indicating that the proteasome degraded the GST-CTD protein from the C-terminus (Figure 3A, lane 1 and 2). Consistent with this, the low molecular weight products were undetectable by WB using an anti-CTD Ab (8WG16; Figure 3B). Notably, the 20S proteasome used was in a latent status with a closed gate and minimal enzymatic activity (Groll et al., 2000; Forster and Hill, 2003). Thus, GST-CTD, like for example the unstructured protein p21 (Forster and Hill, 2003), was capable of activating the 20S proteasome. Low concentrations of SDS render the proteasome gate disordered, leading to proteasome activation (Groll et al., 2000; Forster and Hill, 2003). Indeed, addition of 0.01% SDS to reaction mixtures increased CTD degradation (Figure 3A, lane 3 and 4). In contrast, the proteasome inhibitor MG132 inhibited degradation (Figure 3A, lane 5 and 6).

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We next asked whether phosphorylation of GST-CTD affects its stability in the proteasome assay. For this, we first used a recombinant c-Abl derivative to phosphorylate GST-CTD. This resulted in conversion of a fraction of the GST-CTD to a low-mobility, Tyr1-P isoform, although the majority remained unphosphorylated (Figure 3C, lane 1, Figure 3—figure supplement 1A), consistent with the processive phosphorylation by c-Abl observed previously (Duyster et al., 1995). We then performed the proteasome assay described above using c-Abl-phosphorylated GST-CTD (Figure 3C). Strikingly, the Tyr1 hyperphosphorylated GST-CTD (top panel, upper band, and lower panel) was resistant to degradation (lane 2), while the remaining unphosphorylated GST-CTD (top panel, bottom band) was degraded. Addition of 0.01% SDS again promoted degradation of unphosphorylated GST-CTD, but the Tyr1-P isoform remained resistant (lane 3). Significantly, GST-CTD phosphorylated by the Ser5/Ser7 kinase CDK7, which converted essentially all of the substrate to the hyperphosphorylated form, was not protected from degradation (lanes 4–6), indicating a specific role of Tyr1-P in preventing proteasomal degradation.

We next investigated whether the proteasome functions in Rpb1-Y1F degradation in vivo. We added the proteasomal inhibitors epoxomicin and MG132 to Y1F cells, and measured intact Rpb1 Y1F levels by WB (Figure 3D). Both inhibitors led to approximately twofold elevated levels of Rpb1-Y1F (see Figure 3—figure supplemental 1B for quantification). Although considerable truncated Rpb1-Y1F remained, this likely reflects incomplete proteasomal inhibition and/or accumulation of the truncated form prior to addition of the inhibitors. In any event, our data provide evidence that Tyr1, and specifically Tyr1-P, prevents proteasomal degradation of the CTD in vitro and in vivo.

We next wished to determine the genome-wide impact of the 25F+Y mutation on transcript levels. Using 3′READS (Hoque et al., 2013), a deep sequencing method to quantitate poly(A)+ RNAs, we analyzed 25F+Y and 26r cells, as well as S2A, S5A and T4V cells (all of which, like 25F+Y, are inviable; Hsin et al., 2011; Hsin et al., 2014) for comparison. Cells were treated with tet for 24 hr, and a total of ∼5 million reads mapping to 3′ regions of genes were generated for each cell type (Figure 4—figure supplement 1). Reads were classified into sense RNAs and upstream antisense (ua) RNAs (Figure 4A). uaRNAs were defined as transcripts that did not overlap any known protein-coding genes and used a poly(A) site within 2 kb from the TSS (Figure 4—figure supplement 2). Unexpectedly, the number of genes with upregulated uaRNAs was significantly greater than the number of genes with downregulated uaRNAs, by ∼16-fold (p=10^−21.5), in 25F+Y cells (Figure 4B). S2A and S5A cells showed similar trends but to much lesser extents, fourfold (p=10^−6.7) and 5.6-fold (p=10^−9.0), respectively, while T4V cells in fact showed a trend in the opposite direction (Figure 4B). Using RT-qPCR, we validated several of the uaRNAs (Figure 4C). uaRNAs associated with the ARGLU1, METTL14, SH3BP5 and WEE1 genes were upregulated about twofold in two independent 25F+Y cell lines, consistent with results from RNA-seq (Figure 4—figure supplement 3A). Levels of RPLP1- and CCNB2-associated uaRNAs were indistinguishable in 26r and 25F+Y cells by both methods.

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We next investigated the mechanism underling the enhanced accumulation of uaRNAs in 25F+Y cells. uaRNAs are low-abundant, usually rapidly degraded by the nuclear exosome (Preker et al., 2008; Seila et al., 2009; Wei et al., 2011; Ntini et al., 2013). However, protein levels of four exosome subunits were comparable in 26r and 25F+Y cells (Figure 4—figure supplement 4), suggesting that the increase in uaRNAs was unlikely due to decreased exosome levels. Also, poly(A) sites of uaRNAs were unchanged (Figure 4—figure supplement 3B), indicating that enhanced accumulation did not reflect altered poly(A) site utilization. Another possibility was that transcription of these transcripts was increased. However, ChIP assays indicated that Rpb1 levels were in fact reduced upstream of the ARGLU1, METTL14, SH3BP5 and WEE1 genes in 25F+Y cells (Figure 4D; see also Figure 4—figure supplement 5). Finally, ChIP analyses showed more Tyr1-P on these upstream genes than on the corresponding downstream sense genes (Figure 4—figure supplement 6). Our results point to a role for Tyr1-P in regulating accumulation of uaRNAs by contributing to their rapid turnover.

In this study, we described two important functions for Tyr1 residues: Protecting the CTD from proteolysis and ensuring turnover of uaRNAs. Both these functions are likely conserved throughout vertebrates, as Descostes et al. (Descostes et al., 2014) report remarkably similar results in human cells. While CTD stabilization requires only a single Tyr1 residue, and Tyr1 phosphorylation, more Tyr1 residues are required for other essential functions. As shown previously (Baskaran et al., 1993, 1999), c-Abl can phosphorylate the CTD in vitro, which stabilizes it from proteasomal degradation. But the role of c-Abl in Tyr1 phosphorylation in vivo is unclear, as c-Abl inhibitors had at most modest effects on Try1-P levels in cells (unpublished data). In any event, our results add additional and unexpected complexity to the multiple functions played by the CTD in controlling RNAP II activity.

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Author details

1. Jing-Ping Hsin

   Department of Biological Sciences, Columbia University, New York, United States

   Contribution

   J-PH, Conception and design, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article, Contributed unpublished essential data or reagents

   Competing interests

   No competing interests declared.

2. Wencheng Li

   Department of Biochemistry and Molecular Biology, Rutgers University New Jersey Medical School, Newark, United States

   Contribution

   WL, Acquisition of data, Analysis and interpretation of data, Contributed unpublished essential data or reagents

   Competing interests

   No competing interests declared.

3. Mainul Hoque

   Department of Biochemistry and Molecular Biology, Rutgers University New Jersey Medical School, Newark, United States

   Contribution

   MH, Acquisition of data, Analysis and interpretation of data, Contributed unpublished essential data or reagents

   Competing interests

   No competing interests declared.

4. Bin Tian

   Department of Biochemistry and Molecular Biology, Rutgers University New Jersey Medical School, Newark, United States

   Contribution

   BT, Acquisition of data, Analysis and interpretation of data, Drafting or revising the article, Contributed unpublished essential data or reagents

   Competing interests

   No competing interests declared.

5. James L Manley

   Department of Biological Sciences, Columbia University, New York, United States

   Contribution

   JLM, Conception and design, Analysis and interpretation of data, Drafting or revising the article

   For correspondence

   jlm2@columbia.edu

   Competing interests

   JLM: Senior Editor, eLife.

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