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Thr4 phosphorylation on RNA Pol II occurs at early transcription regulating 3′-end processing
Thr4 phosphorylation’s debut at transcription initiation
https://orcid.org/0000-0001-8192-6005
Moreno Rosamaria Y. Conceptualization Formal analysis Investigation Methodology Resources Validation Visualization Writing - original draft Writing - review & editing
Panina Svetlana B. Data curation Formal analysis Investigation Methodology Software Validation Visualization Writing - original draft Writing - review & editing
Irani Seema Conceptualization Investigation
https://orcid.org/0000-0002-3718-5286
Hardtke Haley A. Investigation Validation
Stephenson Renee Investigation
Floyd Brendan M. Formal analysis Investigation Resources
https://orcid.org/0000-0001-8808-180X
Marcotte Edward M. Formal analysis Funding acquisition Resources Supervision
https://orcid.org/0009-0003-5931-2084
Zhang Qian Methodology Writing - original draft Writing - review & editing
Zhang Y. Jessie Conceptualization Funding acquisition Methodology Project administration Resources Supervision Writing - original draft Writing - review & editing *
Department of Molecular Biosciences, University of Texas, Austin, TX, USA.
* Corresponding author. Email: jzhang@cm.utexas.edu
06 9 2024
06 9 2024
10 36 eadq035023 4 2024
01 8 2024
Copyright © 2024 The Authors, some rights reserved; exclusive licensee American Association for the Advancement of Science. No claim to original U.S. Government Works. Distributed under a Creative Commons Attribution License 4.0 (CC BY).
2024
The Authors
https://creativecommons.org/licenses/by/4.0/ This is an open-access article distributed under the terms of the Creative Commons Attribution license, which permits unrestricted use, distribution, and reproduction in any medium, provided the original work is properly cited.

RNA polymerase II relies on a repetitive sequence domain (YSPTSPS) within its largest subunit to orchestrate transcription. While phosphorylation on serine-2/serine-5 of the carboxyl-terminal heptad repeats is well established, threonine-4’s role remains enigmatic. Paradoxically, threonine-4 phosphorylation was only detected after transcription end sites despite functionally implicated in pausing, elongation, termination, and messenger RNA processing. Our investigation revealed that threonine-4 phosphorylation detection was obstructed by flanking serine-5 phosphorylation at the onset of transcription, which can be removed selectively. Subsequent proteomic analyses identified many proteins recruited to transcription via threonine-4 phosphorylation, which previously were attributed to serine-2. Loss of threonine-4 phosphorylation greatly reduces serine-2 phosphorylation, revealing a cross-talk between the two marks. Last, the function analysis of the threonine-4 phosphorylation highlighted its role in alternative 3′-end processing within pro-proliferative genes. Our findings unveil the true genomic location of this evolutionarily conserved phosphorylation mark and prompt a reassessment of functional assignments of the carboxyl-terminal domain.

Thr4 phosphorylation starts in early transcription, recruits transcription regulators, and is crucial for 3′-end processing.

http://dx.doi.org/10.13039/100000002 National Institutes of Health R01GM104896 http://dx.doi.org/10.13039/100000002 National Institutes of Health R01GM125882 http://dx.doi.org/10.13039/100000002 National Institutes of Health R35GM148356 http://dx.doi.org/10.13039/100000002 National Institutes of Health R35GM122480 http://dx.doi.org/10.13039/100000928 Welch Foundation F-1515 EMM Army Research Office W911NF-12-1-0390 L. Leon Campbell Professorship Fund
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pmcINTRODUCTION

The intricate functions of eukaryotic cells depend on the transcription activities of RNA polymerases I, II, and III. Among these, RNA polymerase II (Pol II) stands out as the primary workhorse, responsible for transcribing all messenger RNAs (mRNAs) for protein expression as well as some small nuclear RNA (snRNA) and small nucleolar RNA (snoRNA) (1–3). To effectively manage the substantial workload, Pol II features a unique C-terminal domain (CTD) in its largest subunit RPB1. This domain is characterized by a conserved repetitive sequence of seven residues YSPTSPS (repeated 26 times for Saccharomyces cerevisiae and 52 for Homo sapiens) and is crucial for coordinating mRNA production and processing (4). While occasional deviations from the consensus sequence occur (mostly in the seventh position and sometimes in the fourth), the overarching presence of the repetitive heptad sequence remains consistent across eukaryotes (5). The CTD does not affect the catalytic activity of RNA Pol II, but its absence or even replacement of specific residues can result in cell death (6, 7).

The distinctive repetitive sequence of the CTD has garnered considerable attention in efforts to comprehend its role in eukaryotic transcription. Central to the CTD’s functionality is its capacity for phosphorylation. Five of the seven residues undergo phosphorylation, and blocking this process results in the cessation of transcription (8). Notably, Ser5 and Ser2 are believed to get phosphorylated at specific stages of transcription—initiation and elongation/termination, respectively (9, 10). These phosphorylation events recruit key transcriptional regulatory proteins to the transcribing Pol II to facilitate transcription with precision (11). Conversely, the roles of the other three phosphorylatable residues of the consensus heptad (Tyr1, Thr4, and Ser7) are less clearly defined with limited knowledge about their cellular function, despite confirmation of their phosphorylation in cells. Thr4, in particular, remains the most mysterious.

Genetic studies using Thr4 variants identified elongation and termination defects when phosphorylation at this position is disrupted as well as implications for Thr4 in mitotic cell cycle regulation (12–15). Despite these putative Thr4 functions, previous chromatin immunoprecipitation sequencing (ChIP-seq) indicates that Thr4 phosphorylation does not occur until late in transcription, reaching its maximum around 500 to 2000 bases after the polyadenylation site (Fig. 1A) (16). Although multiple lines of evidence from both genetic and chemical perturbance studies implicate Thr4 in elongation and coprocessing of mRNA, the absence of Thr4 phosphorylation enrichment in ChIP profiles during the early stages of transcription seems to contradict its proposed functions. Furthermore, a seemingly conservative mutation of Thr4 replaced with alanine in the Pol II CTD, results in cell death in human cells but not in S. cerevisiae or Schizosaccharomyces pombe (16–18). Thus, the role of Thr4 and its phosphorylation in eukaryotic transcription are still very puzzling.

Fig. 1. pThr4 antibody signal is perturbed by neighboring pSer5 and enhanced by Ssu72 treatment.

(A) ChIP profile of pThr4 Pol II dataset from PMID: 22549466. The region between the TSS and TES is scaled to 2000 bp for every gene; −2 kb corresponds to −2 kb from the TSS; +2 kb corresponds to +2 kb from the TES. (B) Dot blot loaded with 5 or 10 μg of CTD peptide #1 (singly phosphorylated pT4) or CTD peptide #2 (doubly phosphorylated pT4pS5) and incubated with pThr4 (6D7) antibody. Structure of Ssu72 bound to a pSer5 CTD for specific targeting of pSer5 dephosphorylation (PDB: 4IMJ) is shown. Comparison was performed using unpaired t test. (C) Dot blot showing serial dilutions of pT4S5 CTD peptide treated with WT Ssu72/symplekin or catalytically deficient Ssu72 C13D D144N/symplekin and incubated with the pThr4 antibody. Comparisons were performed using unpaired t tests. (D) WB showing pThr4 recognition in HEK293 cell lysate treated with Ssu72 or mutant Ssu72 lacking activity. Comparison was performed using unpaired (Ssu72 WT versus untreated) or paired (Ssu72 WT vs Ssu72 Mut) t tests. In all plots, means with SEM are shown and quantification was from three independent biological replicates. *P < 0.05.

ChIP is a powerful tool for unraveling the role of transcriptional regulators, but its efficacy heavily relies on the antibodies used in the experiments. While antibodies targeting the phospho-specific Thr4 of CTD exhibit high specificity, avoiding cross-recognition with other phospho-CTD epitopes, their ability to recognize pThr4 is susceptible to interference from flanking phosphorylation marks on a hyperphosphorylated Pol II (16). Specifically, neighboring Ser5 and Ser2 phosphorylation can impede recognition of the pThr4 epitope when tested on a heptad polypeptide (16). This phenomenon, known as the “masking” effect, introduces the risk of false-negative signals and ambiguity in the genome-wide distribution analysis of pThr4.

Here, we leverage the exceptional specificity of a well-characterized CTD phosphatase to remove the masking effect for pThr4 in situ, which allows us to delineate the genomic location of pThr4 and embark on an exploration of pThr4 function. Notably, because Ser5 phosphorylation occurs promptly upon transcription initiation, the potential masking of pThr4 by pSer5 raises valid concerns of underestimating the level of Thr4 phosphorylation and the challenge of accurately pinpointing its genomic location. Intriguingly, our investigation uncovered a distinct peak immediately following the transcription start site (TSS), accounting for almost 50% of pThr4 peaks without masking events. Intriguingly, protein-coding genes exhibit a different pThr4 profile from noncoding RNA genes. In protein-coding genes, Thr4 phosphorylation level drops significantly after the TSS peak, but it starts to rise to a plateau close to the transcription end site (TES), which is missing in noncoding genes. A proteomic study using a reconstructed phospho-CTD system unveiled a noteworthy overlap of approximately two-thirds of the pThr4 and pSer2 CTD interactomes. Notably, this overlap was particularly pronounced among proteins harboring a CTD binding motif known as the CTD-interacting domain (CID). Subsequent x-ray crystallographic examination of a representative protein, RPRD1B, shed light on the fact that CIDs recognize the CTD via either Ser2 or Thr4 phosphorylation in an analogous manner that is conserved across all CIDs. The ChIP analyses of such proteins are consistent with their recruitment via pThr4 rather than pSer2. Our mechanistic study reveals that Thr4 phosphorylation primes the transcription effect of Ser2 phosphorylation. Functional studies focused on a Thr4 variant incapable of phosphorylation unearthed a pivotal role for Thr4 in productive Pol II elongation and termination at the proper polyadenylation site, critical for the stability of mRNA.

RESULTS

Masking effects of flanking phosphorylation on pThr4 detection in human cells

Previous studies that profiled pThr4 Pol II on the human genome revealed most recruitment occurs after the TES (Fig. 1A) (16). This unexpected genomic location raised concerns over the accuracy of this profile because the same report indicates that Thr4 antibody recognition may be blocked by nearby Ser2 or Ser5 phosphorylation on CTD (16). This concern is particularly significant in human cells as Thr4-Ser5 double phosphorylation accounts for ~20% of double phosphorylation on heptads, as reported in an in-depth analysis of endogenous human CTD phosphorylation mapping using mass spectrometry (MS) (19).

Because ChIP experiments heavily rely on antibody specificity, we first tested if the pThr4 antibody (6D7) cross-recognizes other CTD phosphorylation sites. Using peptides of 18-nucleotide oligomer (about 2.5 heptad repeats) phosphorylated at different sites, we observed no detection of any other phosphorylation sites by the pThr4 antibody, confirming its high specificity (fig. S1A). Next, we evaluated the masking effect of Ser5 phosphorylation on pThr4, a frequent double phosphorylation detected in human cells (Fig. 1B). We used synthetic peptides of 18-nucleotide oligomers with pThr4pSer5 double phosphorylation (Fig. 1B). Strong detection of pThr4 was observed in the singly phosphorylated peptide (peptide #1), but no signal was detected when a pSer5 followed the pThr4 (peptide #2) at the same peptide concentration. Thus, neighboring pSer5 blocks detections of pThr4 by the pThr4 antibody. When transcription occurs in cells, phosphorylation of Ser5 occurs at the beginning of transcription after preinitiation complex (PIC) assembles (9). Thus, if the Thr4 were phosphorylated, then it would likely not be detected by the pThr4 antibody.

Another reported Thr4 antibody masking effect in vitro is observed when Ser2, two residues upstream of pThr4, is phosphorylated (pSer2Pro3pThr4Ser5). Despite the in vitro blocking effect, this potential impediment posed to pThr4 identification is less concerning under physiological conditions as pSer2/pThr4 double phosphorylation is rather rare in cells (19). Subsequent biochemical and cellular experiments have shown that, although Ser2 and Thr4 phosphorylations occur around the same time during transcription cycle, they tend to occur on different heptads because kinases avoid placing a phosphate on the neighboring Ser2 when Thr4 is phosphorylated and vice versa (19). When we used a Ser2 kinase, Dyrk1a, to phosphorylate an 18-nucleotide oligomer synthetic peptide containing phosphorylated Thr4 (fig. S1B), pThr4 detection was not affected, probably due to the Ser2 on different heptads was favored for phosphorylation (fig. S1C).

Another phosphorylation mark co-occurring with Thr4 on the same heptad is Tyr1 (accounts for ~5% of doubly phosphorylated heptads) (19). Previous experiments using synthetic peptides with pTyr1/pThr4 double phosphorylation did not report a masking effect (16). To corroborate this, we evaluated the possibility that Tyr1 phosphorylation affects pThr4 detection by treating pThr4-containing CTD peptide with the Tyr1 kinase, c-Abl (fig. S1B). We found that c-Abl treatment does not affect pThr4 antibody recognition (fig. S1C). Last, the antibody characterization showed that Ser7 on the same heptad repeat does not affect epitope recognition for Thr4 phosphorylation (16). Thus, the pThr4 antibody 6D7 is highly specific for pThr4 but might fail to recognize pThr4 when it co-occurs with Ser5 phosphorylation in cells.

Removing masking effect of Ser5 phosphorylation on pThr4 ChIP

The reported enrichment of pThr4pSer5 double phosphorylation in cells raised concerns for Thr4 phosphorylation detection as the masking effect by adjacent phosphorylation may block pThr4 recognition leading to false-negative results (19). To address this issue, we explored the possibility of eliminating the masking effect on pThr4 antibody recognition by selectively removing the interfering phosphate groups on Ser5 of the CTD. Our prior biochemical and structural analyses have established phosphatase Ssu72 as a highly specific phosphatase for Ser5 of the CTD, distinguishing Ser5 from any other Ser/Thr residues in the CTD heptad (20–22). Our earlier MS and biochemical assays affirm that Ssu72 exerts no effect on the levels of pSer2 or pThr4 while effectively eradicating Ser5 phosphorylation on the CTD (21). The structural element that establishes such high specificity is attributed to the requirement that Pro3 must be in the cis configuration to fit into the Ssu72 active site (Fig. 1B) (21). Phospho-Ser5 extends into the active center of Ssu72 when the heptad forms a tight β turn (Fig. 1B). This β turn is sterically hindered by the Tyr1 residue of the same heptad preventing pSer2 or pThr4 placed into the active site (fig. S1D). In addition, the dephosphorylation of Ser5 by Ssu72 is not blocked by flanking phosphorylation, for example, on pThr4 (Fig. 1B) (20). Biochemically, Ssu72 exhibits weak activity against pSer7, about three magnitudes lower than the activity exhibited toward pSer5 (23). This “star activity,” however, is inconsequential for our purpose of profiling pThr4 in the physiological context because the Ser7 phosphorylation state does not affect Thr4 antibody recognition. Thus, the masking effect in Thr4 ChIP profiling, stemming from the flanking Ser5 phosphorylation, can potentially be removed using Ssu72 as a biochemical tool.

We first tested experimentally if the selective removal of Ser5 phosphorylation by Ssu72 will enhance the detection of pThr4 on a pThr4pSer5 doubly phosphorylated peptide. Ssu72 exhibits maximum stability and activity when it is associated with a scaffolding protein called symplekin (24). Therefore, we purified the Ssu72/symplekin complex and treated the pThr4pSer5 doubly phosphorylated peptide (peptide #2) (Fig. 1C). Our results indicate that the recognition of pThr4 on the doubly phosphorylated peptide by 6D7, which was previously undetectable before phosphatase treatment, exhibited a strong signal upon Ssu72/symplekin treatment (Fig. 1C). To ensure that the observed effect is due to dephosphorylation by Ssu72 rather than other interfering factors, we conducted the same experiment with a catalytically inactive mutant, Ssu72 C13D/D144N. In the absence of the phosphatase activity, 6D7 is unable to recognize pThr4 (Fig. 1C).

We then investigated if the Ssu72/symplekin phosphatase complex could enhance the recognition of pThr4 in the cell lysate by removing pSer5 in the nuclear cell extract. We obtained the nuclear extract from human embryonic kidney (HEK) 293 cells and treated it with no phosphatase, active Ssu72/symplekin [wild-type (WT)], or catalytically inactive Ssu72/symplekin phosphatase complex (C13D/D144N variant of Ssu72). Immunoblotting with the pThr4 antibody 6D7 revealed an approximately threefold increase in signal intensity for the phosphatase-treated sample compared to the untreated and the catalytically dead phosphatase-treated samples (Fig. 1D). This suggests that there is a large percentage of Thr4 being masked. However, this is different for the pSer2 signal in the nuclear cell extract treated with Ssu72 and detected by the pSer2 antibody 3E10, showing no significant difference in recognition compared to untreated and inactive Ssu72 mutant samples (fig. S1E). Thus, the treatment of a hyperphosphorylated Pol II CTD using the CTD phosphatase Ssu72 allows for the detection of pThr4 that was previously blocked by flanking Ser5 phosphorylation.

ChIP-seq analysis of pThr4 of the CTD

To map pThr4 Pol II phospho-marks over the human genome using ChIP-seq, we used phosphatases to selectively remove the masking pSer5 and expose pThr4 for antibody binding. After optimizing the time and amount of phosphatase treatment (fig. S2, A and B), we performed ChIP-seq by adding the Ssu72/symplekin complex after the cell lysis step (Fig. 2A). The nuclear lysate was incubated with the phosphatase solution at 4°C for 30 min before binding to the 6D7 antibody (Fig. 2A). To enhance the rigor of our study, we prepared six biological replicates for the ChIP analysis and assessed the reproducibility of pThr4 ChIP-seq signal. The higher-than-normal sampling was to interrogate how different factors such as cell amount, cell density, and treatment variation affect the outcome of the profile. All the six performed replicates had appropriate immunoprecipitation quality estimated by fraction of reads in peaks (FriP) scores (≥4%), consistent with 1% threshold in the ENCODE guidelines (fig. S2C) (25). As another ChIP-seq quality control (QC), several select genomic loci with high estimated pThr4 enrichment were validated using ChIP–quantitative polymerase chain reaction (qPCR) under Ssu72 treatment versus control (fig. S2D).

Fig. 2. Unbiased mapping of Thr4 phosphorylation on the human genome.

(A) Workflow of pThr4 ChIP experiments with Ssu72 phosphatase treatment. (B) ChIP-seq profile of pThr4 Pol II with Ssu72 treatment along human annotated genes. The region between the TSS and TES is scaled to 2000 bp for every gene; −2 kb corresponds to −2 kb from the TSS; +2 kb corresponds to +2 kb from the TES. (C) Left pie graph shows the number of consensus pThr4 Pol II ChIP-seq peaks between two replicates of Ssu72-treated samples. Right pie graph shows the distribution of genomic locations of consensus peaks between two replicates of pThr4-Ssu72. Promoter region is defined as (−1 kb; +1 kb) from the TSS. Bottom pie graph splits consensus peaks into protein-coding or noncoding gene categories. (D) Box plot of scores’ distribution over the TSS, gene body, and the TES for consensus pThr4 peaks. Promoter region is defined as (−1 kb; +1 kb) from the TSS. Outliers not shown. Groups were compared using Kruskal-Wallis tests with post hoc pairwise tests. ***P < 0.001. (E) Normalized ChIP-seq profile of pThr4 Pol II along protein-coding or noncoding genes. (F) pThr4 signal distribution over snoRNA/miRNA genes or lincRNA (RPKM). Pie graphs (below) denote the localization of pThr4 Pol II within genomic coordinates of snoRNA/miRNA or lincRNA genes. (G) IGV tracks showing the pThr4 signal on several protein-coding or noncoding genes.

The mapping of all the six datasets over the human genome produced the same profile for pThr4—a sharp peak at the TSS followed by a relatively low level of pThr4 until the signal increases to form a plateau steadily at the TES (Fig. 2B). Our data show that the reads per kilobase per million mapped reads (RPKM) values at the TSS peak is consistently higher than maximal RPKM values at the TES plateau (TSS/TES ratios ranged from 1.01 to 1.27). We also performed pThr4 ChIP without Ssu72 treatment and found a significant difference in the TSS/TES ratio wherein the peak at the TSS is markedly lower compared to replicate datasets with Ssu72 treatment (fig. S2E). Previously, it has been noticed that 5% of the genes have a very small peak at the TSS in pThr4 ChIP (16). Our enriched pThr4 binding at the TSS can explain a potential role in transcription elongation where T4A mutation of CTD prevents Pol II from proceeding after the TSS (16). Because Ser5 phosphorylation is mostly enriched at the TSS, it is likely that removing its masking effect reveals a high abundance of pThr4. Thus, the Thr4 phosphorylation profile highlights a sharp peak at the beginning of the transcription. We also notice the plateau near the TES seems to be slightly shifted compared to previous pThr4 ChIP (16), yet the transcriptional implication of such shift requires a follow-up study.

Of the six analyzed ChIP-seq datasets, replicates 1 and 2 had the highest FriP scores (10 and 6%, respectively) (fig. S2C) and the highest numbers of MACS2-called broad pThr4 peaks (q < 0.1; 71,235 and 77,353 peaks, respectively) (table S1). Therefore, we focused on replicates 1 and 2 for more detailed follow-up analysis and derived a consensus pThr4 peakset shared between them (Fig. 2C, left). First, analysis of the genomic distribution of the consensus pThr4 peaks revealed that nearly half of the peaks were located at the promoters, defined as (−1 kb; +1 kb) from the TSS (Fig. 2C, right). The absolute majority (>75%) of pThr4 peaks were mapped to protein-coding genes (Fig. 2C, bottom). Furthermore, pThr4 peaks located in promoters and the TES had significantly higher scores (P < 2.2 × 10−16) than peaks mapped to the gene body (exons/introns) (Fig. 2D), highlighting the biological significance of the pThr4 signal in both transcription start and end. Consistently, the T4A mutant of Pol II was reported to have significantly deregulated, pausing index both at the 5′-initiation site (increased stalling) and 3′-termination site (decreased stalling) compared to WT control (16).

One of the first questions we addressed was if all the genes shared the same pThr4 profile because it was reported that pThr4 has a special function in snoRNA termination (26). In addition, we noticed weak pThr4 peaks in snoRNA, microRNA (miRNA), and long noncoding RNA (lncRNA) genes (Fig. 2C). To answer that, we separated the genes into protein-coding versus noncoding sequences (Fig. 2E). The analysis revealed that the pThr4 profile over protein-coding genes was similar to the overall profile with the sharp peak at the TSS and plateau at the TES region (Fig. 2E). In contrast, noncoding RNAs had a markedly different profile with a wide peak around the TSS region (Fig. 2E). Despite the difference in profile, the score distribution of peaks across snoRNA/miRNA genes followed the same pattern (fig. S2F). To identify how abundant pThr4 binding was in noncoding RNAs, we downloaded genomic regions corresponding to “snoRNA/miRNA” and “lincRNA transcripts” tracks (hg19) from the UCSC Table Browser and overlapped them with genomic coordinates of the consensus pThr4 peakset. A total of 459/2273 unique snoRNA and miRNA sequences (20%) and 1643/21,630 (~8%) long intergenic noncoding RNA (lincRNA) sequences had consensus pThr4 peaks (Fig. 2F) (26). Figure 2G shows examples of pThr4 peaks in protein-coding and noncoding genes. Overall, ChIP-seq results suggest that the distribution of pThr4 binding appears to follow a gene-specific pattern and maps predominantly to TSS and TES regions of protein-coding genes.

Pol II CTD pThr4 interactomes

The latter half of the pThr4 ChIP profile where it plateaus close to the TES is reminiscent of the previous mapping of Ser2 phosphorylation across various cell types and organisms, highlighting a tendency for these two phosphorylation marks to accumulate and plateau near the TES (4). To compare the genomic localization of pThr4 and pSer2, we performed ChIP-seq against pSer2 in the HEK293 cell line (Fig. 3A). The profiles of pThr4 and pSer2 exhibit similarity in shape and transcriptional timing, although pSer2 has only weak signals at the TSS (Fig. 3A). Although the different antibodies used in the pSer2 and pThr4 ChIP do not allow for a direct quantifiable comparison, the profiles do imply that pThr4 and pSer2 occurrence during transcription might be coinciding.

Fig. 3. Recruitment pattern of Thr4 phosphorylation.

(A) Normalized ChIP signal of pThr4 Pol II (samples were treated with Ssu72) and pSer2 Pol II along human annotated genes. The region between the TSS and TES is scaled to 2000 bp for every gene; −2 kb corresponds to −2 kb from the TSS; +2 kb corresponds to +2 kb from the TES. (B) Schematic showing the workflow of T4E pulldown. A 26X CTD peptide with Thr4 mutated to glutamate in every repeat was used as a bait and compared to unphosphorylated WT 26X CTD. Phosphorylated CTD substrates were incubated with the HEK293 nuclear cell extract overnight. CTD substrates were pulled down using glutathione beads and MS/MS analysis was conducted. (C) Volcano plots comparing T4E or pSer2 immunoprecipitation to unphosphorylated CTD immunoprecipitation as a control. Enriched factors were determined using a P value of <0.05. Factors mentioned in text are labeled and shown as red dots. (D) Pie graph showing proteins present in both T4E and pSer2 immunoprecipitation that are either positively or negatively enriched in both pulldowns have differing recruitment compared to control. (E) Schematic showing the protein domain architecture of human CID proteins.

To elucidate the function of Thr4 phosphorylation in eukaryotic transcription, we conducted a proteomic analysis to identify proteins recruited by pThr4 and compared it to proteins recruited in a pSer2 pulldown, considering the highly similar genomic location of the two CTD posttranslational modifications (PTMs) (Fig. 3C). We recombinantly expressed a 26X GST-CTD construct with Thr4 mutated to glutamate at every heptad to mimic the negative charge of a phosphoryl group. Using a label-free proteomic approach, we conducted pulldowns using a GST-26X T4E-CTD as the “bait” protein with an unphosphorylated GST-26X CTD as a control (Fig. 3B). Equal amounts of the nuclear cell lysate containing phosphatase/protease inhibitors were added to individual samples (fig. S3A) and incubated overnight while mixing. After several salt washes, the samples were analyzed by comparing the abundance of pulled-down proteins in phosphorylated samples compared to that in the control.

In parallel, we conducted pSer2 pulldown using a protocol we described before with in vitro reconstruction of pSer2 phosphorylation (27). Comparison of the pSer2 interactome and the T4E pulldown revealed a large proportion of proteins that appeared concurrently in both the T4E and pSer2 samples (Fig. 3C and table S2). Notably, 110 proteins (64% of the total) demonstrated simultaneous recruitment or depletion between the two pulldowns, underscoring the overlapping recruitment profiles of pSer2 and pThr4 (Fig. 3D and fig. S3, B and C).

A close inspection of the proteins pulled down by the pSer2 or pThr4-mimic (T4E) reveals an overrepresentation of proteins containing a protein motif called the CID (Fig. 3E and fig. S3D). This binding motif is highly conserved throughout eukaryotes, and most identified CIDs exhibit selective binding toward pSer2 over pSer5. In our proteomic study, proteins containing this domain were spotted in at least one if not both pulldowns, including RPRD1A, RPRD1B, RPRD2, SCAF4, and U2SURP (Fig. 3E). This parallel result led us to wonder if, generally, CIDs are dually capable of recognizing either pSer2 or pThr4 in vitro. Almost all known CID-containing proteins have been implicated in termination and mRNA processing. A previous detailed characterization of yeast Rtt103 (which contains a CID) has shown that it can bind phosphorylated Thr4 in addition to binding phosphorylated Ser2 (28). SCAF4/SCAF8 are CID proteins that suppress early, alternative polyadenylation (APA) sites and regulates transcriptional termination (29). Similarly, Rtt103, a yeast CID termination factor, forms a complex with Rat1/Rai1 (30) and is important in regulating termination of both protein-coding and a subset of noncoding genes through interactions with pThr4 marks on snoRNA (13). Furthermore, a Cleavage and Polyadenylation (CPA) complex member, PCF11, is highly prevalent at the 3′ end of genes (31) and influences APA site usage, with decreased PCF11 levels leading to isoforms with distal sites chosen (32). The role of CID proteins in termination might be linked to pThr4 phosphorylation rather than pSer2 phosphorylation.

CID motif–containing protein RPRD1B can bind both pSer2 and pThr4

To test if proteins containing a CID motif that were previously characterized as pSer2 binders can also bind to pThr4, we started with a structural and biophysical analysis of RPRD1B, a human transcription regulator that contains a CID that strongly interacts with pSer2. We first used fluorescence anisotropy (FA) to measure the interaction of RPRD1B with a 16-nucleotide oligomer CTD polypeptide phosphorylated at a single position either at Ser2 or at Thr4 located in the middle of the heptad. Purified RPRD1B can bind to both pSer2 and pThr4 CTD peptides, with a little tighter association to the pThr4 peptide [dissociation constant (Kd) of 22.8 ± 11 μM for pSer2 and 5.3 ± 2 μM for pThr4] (Fig. 4A). To investigate the recognition mode of human CID proteins to a pThr4 CTD, we cocrystallized RPRD1B with a 15-nucleotide oligomer, singly phosphorylated Thr4 CTD peptide. We determined the complex structure at 2.5 Å (statistics for data collection and refinement in table S6). The RPRD1B-CID is composed of eight α helices arranged in a right-handed superhelical array, characteristic of the conserved CID fold (33–35). RPRD1B exists as a monomer in our crystal structure, consistent with its oligomerization state in solution as shown in its gel filtration profile (fig. S4A) (35). We observed a stretch of elongated positive density, consistent with the CTD polypeptide based on which we modeled in 12 residues of the 15-nucleotide oligomer CTD peptide (Fig. 4B and fig. S4B). Previously, RPRD1B has been cocrystallized with a CTD peptide containing Ser2 phosphorylation [Protein Data Bank (PDB): 4Q94] (35). Unexpectedly, the mode of binding in the CTD backbone is almost identical in the two structures, with the CTD peptide anchored in place in both structures through several hydrophobic pockets (fig. S4C). The aromatic ring of Tyr1 of the CTD is situated in a hydrophobic pocket formed by Val23 and Tyr61, and Pro3 fits into a hydrophobic core composed of Tyr61, Leu104, Leu107, and Ile110 (fig. S4C). The hydroxyl of Tyr1 forms hydrogen bonds with Asp65 (fig. S4C). Notably, in both structures, the recognition of the phospho-CTD residue depends on Arg106 (Fig. 4C). Arg106 forms salt bridge interactions with the phosphate group when the phosphate group is extended either from Ser2 or Thr4 (Fig. 4C). Mutations in this arginine residue do not alter the stability of RPRD1B (fig. S4D) but do abolish binding to pThr4 or pSer2 (Fig. 4A). The only difference in the CTD peptide recognition of RPRD1B for pSer2 or pThr4 CTD peptide is R114, which can potentially form a salt bridge with the phosphate group of the Thr4 phosphate but is located too far (6.9 Å) from the Ser2 phosphate (Fig. 4C).

Fig. 4. CID proteins recognize both pSer2 and pThr4 CTD.

(A) FA measurements of the CID of RPRD1B with FITC- labeled pS2/pT4 CTD peptides. (B) 2Fo-Fc electron density map (contour to 1.0 s) of RPRD1B’s CID complexed with the pThr4 CTD peptide. (C) Side-by-side view of RPRD1B binding to the pThr4 or pSer2 CTD peptide (PDB: 4Q94). Conserved Arg106 and Arg114 are shown with stick representation. (D) FA measurements of WT or mutant CID of SCAF4, RPRD1A, and RPRD2 with pS2/pT4 CTD peptides. Experimental isotherms were fitted to a total binding model. Binding assays were performed in triplicate. Error bars indicate the SD. (E) Structural modeling of the conserved recognition of CID of SCAF4 (pink), RPRD1A (blue), and RPRD2 (light blue) binding to the pThr4 CTD peptide.

CID is a dual-binding module

Our structure of RPRD1B in complex with pThr4 CTD peptides indicates that Arg106 mediates pSer2/pThr4 recognition through salt bridge formation to the phosphorylated CTD residue (Fig. 4A). This arginine is conserved across all CID proteins except for Nrd1 in S. cerevisiae, which does not exhibit significant binding toward pSer2. We thus speculated that CIDs might be all capable of binding to pThr4 and pSer2 via this conserved arginine. A pSer2 binding protein we just identified, CHERP, can also bind to a pThr4 CTD (Kd = 10.7 ± 2 μM for pSer2 and 3.9 ± 1 μM for pThr4) (27). To verify the potential dual recognition of pSer2 and pThr4 by CID-containing proteins, we isolated the CIDs of RPRD1A, RPRD2, and SCAF4 and quantified their binding to CTD phospho-peptides using FA (fig. S4, E and F). Similar to RPRD1B and CHERP, RPRD1A, RPRD2, and SCAF4 all exhibit strong binding to both pSer2 and pThr4 CTD peptides with comparable affinity (RPRD1A, Kd = 12.3 ± 5 μM for pThr4 and 13.8 ± 1 μM for pSer2; RPRD2, Kd = 6.6 ± 2 μM for pThr4 and 8.7 ± 2 μM for pSer2; and SCAF4, Kd = 8.1 ± 4 μM for pThr4 and 6.7 ± 4 μM) (Fig. 4D). The structures of these CIDs are highly similar to RPRD1B, with all key residues for CTD recognition conserved (Fig. 4E). Structural superimposition reveals that RPRD1A, RPRD2, and SCAF4 share an identical binding groove as RPRD1B. Notably, the position of the conserved Arg that can bind the CTD phosphate group can extend and interact with either Ser2 or Thr4 (Fig. 4E). Thus, transcription regulators containing a CID, previously identified as Ser2 binding proteins, can also bind to pThr4, sometimes with an even stronger affinity than that observed for pSer2.

RNA Pol II recruits CID-containing proteins through pThr4/pSer2

The in vitro association of CID-containing proteins with the phospho-CTD motivates us to evaluate their association with RNA Pol II in cells. Using RPRD1B as an example, we first examined the cellular location through immunofluorescence (IF) staining in HEK293 cells (Fig. 5A). As anticipated, HA-RPRD1B and pSer2 Pol II colocalize in transfected HEK293 cells. IF of HA-RPRD1B and pThr4 Pol II reveals substantial overlap in Pol II clusters (Fig. 5A). However, colocalization of RPRD1B with a hyperphosphorylated Pol II is greatly diminished when RPRD1B lacks a CID (Fig. 5A). The marked drop in colocalization indicates that RPRD1B relies on the CID interaction to associate with a phosphorylated Pol II at either Thr4 or Ser2 during ongoing transcription, echoing our in vitro observations of RPRD1B interacting with phospho-CTD polypeptides.

Fig. 5. CID proteins colocalize with pSer2 and pThr4 Pol II.

(A) Representative confocal fluorescent images of HA-RPRD1B full length or HA-RPRD1B ΔCID (red), pSer2 or pThr4 Pol II (green), and DAPI (blue) in HEK293 cells. Scale bars, 5 μm. Profile intensity plots between red and green channels are shown. (B) Confocal fluorescent images of HA-CHERP full length (red) and pSer2/pThr4 Pol II (green). Scale bars, 5 μm. (C) Confocal fluorescent images of HA-RPRD1B (red) and 52X T4A Pol II or 52X WT Pol II (yellow). Scale bars, 5 μm. Profile intensity plots between red and green or red and yellow channels are shown. All IF experiments were performed three independent times. (D) Anti-YFP coimmunoprecipitation of T4A Pol II or WT Pol II with HA-RPRD1B. Representative blots are shown, and quantification is based on three independent biological replicates. Plot shows means with SD. IB, immunoblot; IP, immunoprecipitation; WCL, whole-cell lysate. (E) Distribution of the input normalized ChIP signal of HA-RPRD1B or HA-CHERP across human annotated genes (27). Average peak signal between a window 2 kb upstream/downstream from the TSS/TES. *P < 0.05.

Recently, through proteomic studies, we identified a transcription splicing factor, CHERP, as a pSer2 binding protein (27). Similar to RPRD1B and other pSer2 binding CID proteins, CHERP also exhibits strong binding to pThr4 CTD in vitro (27). We thus explore if CHERP colocalizes with a Thr4 phosphorylated RNA Pol II in cells. When we conducted the same IF assay, hemagglutinin (HA)–tagged CHERP colocalizes extensively with phosphorylated Ser2 and Thr4 Pol II (Fig. 5B). As we showed previously, the recruitment of CHERP to RNA Pol II relies on its CID (27). Thus, the CIDs found in transcription regulators, which mediate binding to pSer2 and pThr4 in vitro, are responsible for the recruitment of their parent proteins to ongoing transcription.

To understand the function of T4 phosphorylation, we generated a construct of mammalian RPB1 with a CTD in which Thr4 in every heptad repeat in the 52 repeats of RPB1 is replaced by alanine, thus impossible to get phosphorylated at the Thr4 position. To test if the recruitment of RPRD1B relies on Thr4 phosphorylation, we visualized the subcellular localization of HA-RPRD1B in conjunction with either RPB1 T4A-EYFP or WT-EYFP (Fig. 5C). We observed colocalization of RPRD1B with WT RPB1 but not in the T4A variant. We next examined if RPRD1B associates with RPB1 T4A in the cell extract. To this end, we performed coimmunoprecipitation assays with extracts from cells expressing similar protein levels of RPB1 WT-EYFP or T4A-EYFP. Using the enhanced yellow fluorescent protein (EYFP) tag for immunoprecipitation, we found that endogenous RPRD1B coimmunoprecipitates with WT RPB1, but there is a significant reduction in binding toward RPB1 T4A (Fig. 5D).

To identify the genomic location for CID-containing proteins, we conducted ChIP-seq analysis of HA-RPRD1B. We expressed an HA-tagged full-length RPRD1B in HEK293 cells and used double cross-linking for ChIP-seq analysis as RPRD1B binding to chromatin is an indirect interaction through direct Pol II binding (fig. S5A). The ChIP profile of RPRD1B shows a high peak around the TSS with a small flat plateau close to the TES of genes. The profile is consistent between biological replicates with high reproducibility (Fig. 5E and fig. S5, B to D). RPRD1B has been detected at the downstream region of the LEO1 gene (36) as well as showing strong occupancy at the promoter and polyadenylate [poly(A)] cleavage sites of the cyclin D1 gene (37). The profile for genomic binding of RPRD1B is reminiscent of the result in our CHERP study where a similar profile is found with a pronounced peak right at the TSS of the genes (Fig. 5E). Notably, the genomic localization of CHERP was eliminated when the CID was omitted in CHERP (27). In both cases, the genomic location of these CID-containing proteins resembles the pThr4 ChIP profile more than the pSer2 ChIP profile. Together, all our data suggest that there is a strong likelihood that CID-containing transcription regulators are recruited to the transcription apparatus through Thr4 phosphorylation rather than Ser2 phosphorylation.

T4 phosphorylation cross-talks with S2 phosphorylation

With T4 and S2 exhibiting overlapping interactomes, we considered the possibility that Thr4 and Ser2 phosphorylation cross-talk. Because pThr4 seems to occur at an earlier stage of transcription (Figs. 2B and 3A), we inquired if the level of Thr4 phosphorylation affected the phosphorylation of Ser2. To address this issue, we transiently transfected plasmids containing a WT RPB1 or a variant with all Thr4 replaced by Ala. To eliminate the interference of endogenous RPB1, we introduced a mutation into each plasmid, rendering the introduced protein resistant to α-amanitin. We established a concentration of α-amanitin that eliminated endogenous RPB1 and monitored the expression of either EYFP-RPB1 mutants using Western blot (WB) to ensure equal levels of RPB1 (T4A versus WT CTD) between conditions (fig. S6, A to C). With the exogenous WT RPB1 and T4A expressed at similar levels, we treated the cell with α-amanitin to eliminate the endogenous Pol II. After 48 hours of treatment, we collected the cells and analyzed the genomic location of the pSer2 in RPB1. We noticed that the pSer2 distribution varied greatly upon T4A mutation with a greatly reduced level of phosphorylation (Fig. 6A). Unlike the substantial accumulation of Ser2 phosphorylation close to the TES in normal RPB1, the loss of Thr4 phosphorylation in the RPB1 T4A variant suppresses Ser2 phosphorylation, especially at the TES region. Thus, the phosphorylation of Thr4 affects the pattern of Ser2 phosphorylation. The cross-talk between the two marks deserves further investigation and demands reevaluation of previous observation attributed to Ser2 phosphorylation to consider the possibility that it is the secondary effect from Thr4 phosphorylation.

Fig. 6. Function of Thr4 phosphorylation in 3′ processing.

(A) Normalized pSer2 ChIP signal along human genes when T4A or WT Pol II CTD is expressed. (B) Volcano plot showing the log fold change (thresholds of 1.5 and −1.5) of gene expression changes due to 52X T4A CTD expression with an adjusted P value threshold of 0.05. (C) GO analysis of the above significant negatively regulated genes upon 52X T4A-CTD expression. (D) Histogram of comparisons between significant ψ values in cells expressing 52X T4A or 52X WT RNA Pol II CTD (FDR < 0.05). Positive ψ values are shown in green, and negative ψ values are shown in orange. (E) Example of the gene (CCND1) found to prefer distal poly(A) site usage when transcribed. Difference between proximal and distal poly(A) sites estimated by LABRAT is 6234 bp. (F) GO analysis of biological processes for transcripts with significant exon inclusion and exclusion events.

Effects of Thr4 phosphorylation on transcription

Thr4 phosphorylation has been implicated in cell cycle regulation (12) where Thr4 mutations display serious mitotic defects such as multipolar spindles and polyploid cells (12). Furthermore, Thr4 modification might also play a role in transcription-coupled DNA repair (26). To understand how Thr4 phosphorylation affects transcription, we characterized transcriptional alterations in mammalian cells expressing an RPB1 T4A variant after we removed endogenous Pol II through α-amanitin elimination for 48 hours. The reproducibility of biological replicates was high as the 52X T4A and 52X WT duplicates clustered together with a strong correlation coefficient (fig. S6D). In terms of the differentially expressed genes (DEGs), there were 996 down-regulated genes (|fold change| > 1.5 and adj. P value < 0.05) and 257 up-regulated genes in 52X T4A cells (Fig. 6B and table S3). Pathway analysis reveals that down-regulated genes include an abundance of those playing a role in cytoskeleton organization (Fig. 6C). This is consistent with previous results showing that mitotic Pol II at centrosomes exclusively retains pThr4 marks and mutations at this position alter mitotic division (12).

Our mechanistic study indicates that the CID-containing proteins identified to be recruited to Pol II through Thr4/Ser2 phosphorylation and these proteins mostly play a role in a proper 3′ untranslated region (3′UTR) in the transcriptome (30, 38). The sequence content of the 3′UTR region can vary greatly depending on if a more proximal or distal poly(A) site is used to terminate the transcript (39). The 3′UTR maintains the stability of the transcript by housing regulatory regions that determine mRNA localization, binding regions for RNA binding proteins, miRNA recognition sites, and AU-rich elements (AREs) that promote mRNA decay in a controlled manner (40, 41). Misregulation of poly(A) site usage can lead to removal or addition of binding sites, thereby altering the metabolism of the mRNA product (42). To ascertain the global impact of pThr4 on polyadenylated transcripts, we compared APA site usage of cells containing T4A or WT Pol II α-amanitin mutants by the LABRAT method (43). APA analysis showed a significant increase in distal poly(A) usage, denoted by a positive Δ-Ψ value, in 573 genes when 52X T4A RPB1 is expressed compared to 52X WT, whereas only 140 genes showed the opposite trend (Fig. 6D, fig. S6E, and table S4). Gene ontology (GO) analysis for the 573 genes with increased 3′UTR lengths revealed processes that maintain intracellular transport and mitotic cell cycle progression among the top 20 enriched pathways (table S4).

To further dissect the effects of distal poly(A) site usage in the T4A Pol II mutant on its phenotype, we used the ARED-Plus database to predict genes with AREs in their 3′UTRs (44). The analysis showed that one-third of the genes (181/573) favoring distal poly(A) sites contained predicted AREs in 3′UTR. In addition, 14% of this overlap was simultaneously found to be down-regulated by DEG analysis (tables S1 and S3), which supports a notion about the decreased stability of ARE-containing transcripts. We followed up on one of the most significant and large GO categories among distal poly(A) genes, “Cell cycle process” [66 genes, fold change = 1.9, false discovery rate (FDR) = 0.00014] (table S4). Using this gene subset as a query against database (44), we found that 19 of 66 genes were predicted to contain AREs in their 3′UTR (table S4). Among these genes, there were important pro-proliferative mediators such as cyclins CCND1 and CCND3, cyclin-dependent kinase CDK2, mitotic checkpoint protein BUB3, and NEK7 kinase, a regulator of cell division. In addition, two genes in this group—microtubule stabilizer CLASP2 and huntingtin HTT—were also down-regulated based on DEG analysis. By querying the LABRAT output against the transcript annotation from the UCSC Genome Browser (hg38), we confirmed the presence of ARE sequences in the longest transcripts of these genes with distal poly(A) sites and the lack of thereof in the transcripts favoring proximal poly(A) sites (CCND1 as an example is shown in Fig. 6E). Therefore, distal poly(A) site usage could add AU-rich sequences in mRNA transcripts of genes important for cell cycle progression that may eventually lead to AU-mediated decay and proliferation arrest. This is consistent with the reported phenotype that the T4A mutant affects cell cycle regulation (12). Distal poly(A) site usage is reminiscent of termination defects in budding yeast where Pol II is detected transcribing far downstream at snoRNA termination sites when T4 is mutated (13). Likewise, a T4V substitution also displays delayed dissociation of Pol II at termination sites with a downstream shift in positions of poly(A) sites in budding yeast (14).

Although the CTD itself does not directly interact with the core components of spliceosome, the presence of CTD is believed to promote spliceosome assembly (45). To investigate the effects of pThr4 specific roles on the selection of splicing site, we conducted deep sequencing of RNA transcription to examine five major alternative splicing events (ASEs) by the rMATS tool (46). The alternative splicing analysis revealed 6359 unique isoform changes across five different event types (fig. S6F and table S5) with a significance threshold of FDR < 0.05 and inclusion level difference (ILD) ≥ 10% between experimental groups. Although variations are identified in each type of ASE, there was no obvious trend to favor inclusion or exclusion of certain exons or introns. However, GO enrichment analysis of transcripts with ASEs highlighted biological processes such as “mitotic cell cycle” and “chromosome segregation” to be significantly overrepresented (Fig. 6F). Dysregulation of the mitotic cell cycle could emanate from alternative splicing changes in transcripts involved in its maintenance of which phosphorylation at Thr4 controls.

Overall, our data suggest an important functional role of Thr4 phosphorylation in transcripts’ 3′-end processing, specifically in the preferential choice of distal poly(A) sites that potentially leads to addition of binding sites, such as AREs, and decreased stability of the transcripts.

DISCUSSION

The role of Thr4 within the highly conserved sequence of the CTD heptad in RNA Pol II has long confounded researchers. Its significance becomes apparent as its replacement induces severe defects in elongation, termination, and processing, ultimately culminating in mammalian cell death (6, 7). However, despite its crucial role in early transcriptional events, Thr4 phosphorylation marks remain undetected until the conclusion of transcriptional events, typically occurring 500 to 2000 bp downstream of polyadenylation sites (16, 47). Scientists have pondered the possibility that this contradiction may stem from the strong masking effect of neighboring Ser5 phosphorylation on the pThr4 antibodies at the onset of the transcription cycle (47). Although pThr4 antibodies exhibit high specificity, they are susceptible to interference from Ser5 phosphorylation (16), a prevalent event during transcription initiation.

Our use of phosphatase complex, Ssu72/symplekin, to eliminate masking Ser5 phosphorylation revealed a pronounced peak near the TSS in nearly all expressed protein-coding genes across all six datasets. While a previous study detected a minor peak at the TSS in the top 5% of genes, our study identified the pThr4 peak in nearly 10,000 protein-coding genes (16). The emergence of the previously unidentified pThr4 peak near the TSS elucidates the involvement of pThr4 in elongation, consistent with genetic and mutagenesis studies of T4A cells. Previous investigations have demonstrated that T4A substitution within the CTD prompts RNA Pol II to stall at the TSS (16). Our ChIP analysis supports this observation, indicating that significant Thr4 phosphorylation initiates precisely when productive elongation should commence. Consequently, the inability to phosphorylate Thr4, as observed in T4A mutants, hinders Pol II from entering productive elongation, underscoring the critical role of Thr4 phosphorylation in transcriptional dynamics. The transcriptomic analysis also unveils the molecular mechanism behind T4A lethality and cell cycle regulation (12). Upon T4A mutation, the genes involved in mitotic cell cycle regulations are affected in a multilayer manner. The foremost category of down-regulated genes is involved in cytoskeleton organization (Fig. 6C), the cell cycle genes altered to a distal poly(A) site (Fig. 6D), and alternations in splicing events lead to changes of cell cycle genes (Fig. 6F).

Our study uncovered a distinctive property of pThr4 not observed in other Pol II PTMs: Its genomic profile varies among gene types, suggesting a gene-specific CTD code. Analysis of ChIP-seq results reveals a consistent distribution profile of pThr4 among protein-coding genes, whereas noncoding genes such as snoRNAs exhibit a distinct profile characterized by a broad peak around the TSS with no significant association at the TES (see Fig. 2E). These distinct profiles imply that the role of T4 phosphorylation may differ between these gene groups, highlighting a gene-specific CTD coding system. Given the association of pThr4 alteration with environmental cues (17, 18), it may serve as a mark specifically reserved for stress response via snoRNA regulation. The T4A variant in yeast caused significant changes in snoRNA expression (13, 26). Investigating if the distribution of T4 phosphorylation varies across gene categories thus holds substantial interest and could illuminate evolutionary adaptations in transcriptional regulation.

The similarity between the pThr4 phenotype and function lies in its resemblance to pSer2. Their ChIP profiles exhibit notable similarities, and the proteins recruited by this mark to RNA Pol II are nearly identical as found in proteomic studies from our and other labs (14, 27). Our ChIP mechanistic study reveals cross-talk between the two, wherein pThr4 influences the level of Ser2 phosphorylation. The crucial occurrence of pThr4 at the outset of transcription raises the intriguing possibility that many factors previously attributed to Ser2 phosphorylation function may deserve to be credited to pThr4 instead. For instance, a family of proteins containing a motif (CID) for CTD recognition, traditionally known to bind pSer2, demonstrates similar, sometimes even tighter, binding to pThr4 in vitro, with a ChIP recruitment profile more consistent with pThr4. Notably, these factors predominantly play roles in 3′-end processing. Given pThr4’s earlier appearance in the transcription cycle and its potent binding to regulatory proteins, it is prudent to reconsider the previous characterization of Ser2 as one of the two major PTM marks in eukaryotic transcription, potentially overlooking the collaborative relationship of Ser2 and Thr4. Our ChIP-seq studies of T4A Pol II provide evidence that loss of phosphorylation at this site lowers levels of Ser2 phosphorylation. Considering that both marks are phosphorylated by the positive transcription elongation factor b (P-TEFb), which is recruited to Pol II by a mediator and BRD4 close to the TSS, the new ChIP profile of pThr4 necessitates a reassessment of the functions traditionally ascribed to pSer2, suggesting that Thr4 phosphorylation may substantially contribute to processes originally solely attributed to pSer2.

MATERIALS AND METHODS

Cell culture

HEK293 cells were purchased from the American Type Culture Collection (Manassas, VA, United States). Cells were routinely cultured in Dulbecco’s modified Eagle’s media (Sigma-Aldrich, St. Louis, MO, United States, product no. D6429), supplemented with 10% Opti-Gold fetal bovine serum (GenDEPOT, Katy, TX, United States) at 37°C in humidified atmosphere with 5% CO2. HyClone penicillin and streptomycin mix (Cytiva, Marlborough, MA, United States) was added to the media to reach a final concentration of 1%.

Cloning

The 26X T4E-CTD and 52X T4A substrate was ordered as a synthetic gene and cloned into a pET28a (Novogene, Sacramento, CA, United States) derivative vector encoding a 6xHis tag followed by a glutathione S-transferase (GST) tag and a 3C protease site or an N-terminal HA-tagged mammalian expression vector. The RPRD1B-CID (encoding residues 2 to 133), RPRD1A-CID (residues 2 to 133), RPRD2-CID (residues 19 to 149), and SCAF4-CID (residues 1 to 139) and Dyrk1a kinase domain (residues 127 to 485) were ordered as synthetic genes. The full-length RPRD1B cDNA (clone: HG14027-G) encoding residues 1 to 326 was cloned into a mammalian expression vector containing a cytomegalovirus promoter and an N-terminal HA tag. The 52X WT CTD harboring α-amanitin resistant and an EYFP tag was from Addgene (plasmid no. 75284). Drosophila Ssu72 (1 to 195) and symplekin (residues 19 to 351) were cloned into a pET28b vector encoding a 6xHis tag and Small Ubiquitin-like Modifier (SUMO) tag.

Protein expression and purification

For protein expression, BL21 (DE3) cells expressing RPRD1B-CID, RPRD1A-CID, RPRD2-CID, SCAF4-CID, Dyrk1a, or GST-CTD substrates were grown in 1-liter cultures at 37°C in Luria-Bertani (LB) broth (Thermo Fisher Scientific, Waltham, MA, United States) containing kanamycin (50 μg/ml). Once the cultures reached an OD 600 (optical density at 600 nm) value of 0.6 to 0.8, the protein expression was induced with 0.25 mM isopropyl-β-d-thiogalactopyranoside, and the cultures were grown for an additional 16 hours at 18°C. The cells were pelleted and resuspended in a lysis buffer [50 mM tris-HCl (pH 8.0), 500 mM NaCl, 15 mM imidazole, 10% glycerol, 0.1% Triton X-100, and 10 mM 2-mercaptoethanol (BME)] and sonicated at 90 A for 2.5 min of 1-s on/5-s off cycles on ice. The lysate was cleared by centrifugation at 15,000 rpm for 45 min at 4°C. The supernatant was loaded over 3 ml of Ni-NTA beads (Qiagen, Germany) equilibrated in a lysis buffer then washed with a wash buffer containing 50 mM tris-HCl (pH 8.0), 500 mM NaCl, 30 mM imidazole, and 10 mM BME. The recombinant protein was eluted with a buffer containing 50 mM tris-HCl (pH 8.0), 500 mM NaCl, 300 mM imidazole, and 10 mM BME. Protein fractions were pooled and dialyzed overnight at 4°C in a 10.0-kDa dialysis membrane (Thermo Fisher Scientific) against a dialysis buffer [50 mM tris-HCl (pH 7.5), 100 mM NaCl, and 10 mM BME]. The protein was polished using gel filtration chromatography and loaded onto a Superdex 75 or 200 size exclusion column (GE) in a gel filtration buffer. For Ssu72 and symplekin, the individual proteins were concentrated, combined, and dialyzed overnight followed by size exclusion chromatography of the complex. Peak fractions were analyzed by SDS.

Dot blot

Serial dilutions of pThr4 CTD peptide or Dyrk1a/Abl1 kinase-treated pThr4 CTD peptide were spotted on an activated nitrocellulose membrane. The membrane was allowed to dry and blocked by soaking in 5% bovine serum albumin (BSA)/Tris-buffered saline with 0.1% Tween 20 detergent (TBS-T) for 30 min at room temperature. The membrane was then incubated with primary antibody (1:1000) in BSA/TBS-T for 30 min at room temperature. The membrane was washed three times with TBS-T for 5 min each. Then, the membrane was incubated with secondary antibody (1:10,000) for 30 min at room temperature. The membrane was washed three times and visualized on a LI-COR Odyssey CLx image reader.

Western blot

Cells were lysed in a radioimmunoprecipitation assay lysis buffer [50 mM tris-HCl (pH 8.0), 150 mM NaCl, NP-40, 0.5% sodium deoxycholate, and 0.1% SDS] and 1× protease inhibitor cocktail (Roche, Indianapolis, IN, United States). Protein concentrations were quantified with the Bradford protein assay. Briefly, 25 μg of protein extracts was loaded and separated by SDS–polyacrylamide gel electrophoresis gels. Blotting was performed with standard protocols using a PVDF membrane (Bio-Rad, Hercules, CA, United States). Membranes were blocked for 1 hour in a blocking buffer [5% BSA in phosphate-buffered saline with Tween detergent (PBST)] and probed with primary antibodies at 1:1000 dilution at 4°C overnight. After three washes with PBST, the membranes were incubated with diluted goat anti-rabbit or anti-rat secondary IRDye 680RD antibody at 1:10,000 (LI-COR, Lincoln, NE, United States) for 1 hour at room temperature. After washing, membranes were visualized on the LI-COR Odyssey CLx image reader. For WB or dot blot analysis, phospho-specific antibodies, pThr4 (catalog no. 61361; 1:100 to 1:800 dilution for WB and IF) and pSer2 (Sigma-Aldrich, stock keeping unit (SKU): MABE953; 1:1000 dilution for WB and IF), green fluorescent protein (GFP) antibody (catalog no. 50430-2-AP; 1:1000 dilution), and HA antibody (catalog no. C29F4; 1:1000 dilution for WB and 1:800 for IF) were used. The RPRD1B antibody is from Cell Signaling Technology (catalog no. 74693; 1:1000 dilution for WB).

Coimmunoprecipitation

Cellular extracts were prepared by incubating cells with a lysis buffer [50 mM tris-HCl (pH 8.0), 150 mM NaCl, 0.5% NP-40, 1 mM phenylmethylsulfonyl fluoride, and 1× protease inhibitor) for 30 min on ice. The supernatant was collected by centrifugation at 12,000g for 20 min at 4°C. For immunoprecipitation, Dynabeads Protein A (20 μl, Invitrogen) was incubated with 2 μg of antibody overnight at 4°C with rotation. Subsequently, 250 μg of protein was incubated with the antibody-bound beads for an additional 2 hours and washed three times with a lysis buffer. The precipitated proteins were eluted from the beads with a 2× SDS loading buffer and boiled for 5 min, followed by WB analyses. Three independent replicates of each IP experiment were performed.

Immunofluorescence

In brief, HEK293 cells were transfected with HA-RPRD1B and 52X T4A Pol II or 52X WT Pol II using polyethylenimine (PEI) (1:7 plasmid-to-reagent ratio) to overexpress the protein of interest. Cells were washed with phosphate-buffered saline (PBS) and fixed in 1% formaldehyde for 15 min at room temperature. Cells were permeabilized with 0.2% Triton X-100 to allow antibody labeling. Subsequently, the samples were blocked with 2% BSA for 30 min and incubated with primary antibody for 1 hour at room temperature. After washing with PBS, the cells were stained with secondary antibody [goat anti-rabbit immunoglobulin G (IgG) (H+L) cross-adsorbed secondary antibody Alexa Fluor 488 or goat anti-rat IgG (H+L) cross-adsorbed secondary antibody Alexa Fluor 568, Thermo Fisher Scientific] for 1 hour at room temperature. Cells were counterstained with 4′,6-diamidino-2-phenylindole (DAPI) for nuclear visualization, and coverslips were mounted with antifade fluorescent mounting media (Abcam, catalog no. ab104135). Standard fluorescence images were captured using a confocal microscope (Zeiss LSM 710). Confocal images were acquired with the Plan-Apo 63x oil immersion lens and analyzed using the Zen/ImageJ program.

Crystallization

Initial crystallization conditions for RPRD1B-CID with pThr4 CTD peptide were identified using sparse-matrix screening using a Phoenix crystallization robotic system (Art Robbins Instruments). The identified hits for crystallization were optimized systematically using the sitting drop vapor diffusion technique. The complex structure was crystallized in 20 to 32% PEG-3350 (polyethylene glycol, molecular weight 3350), 0.1 M lithium sulfate, and a 1:3 molar ratio of protein to peptide. In all crystallization setups, a protein solution (~20 mg/ml) was mixed with an equal volume of the reservoir solution and equilibrated against 500 μl of the reservoir at room temperature. All crystals were cryoprotected with mother liquor supplemented with 30% glycerol and flash-frozen in liquid nitrogen.

Data collection, processing, structure determination, and refinement

X-ray diffraction data for the RPRD1B-pThr4 structure were collected at the Advanced Photon Source beamline 23-ID-D (Argonne National Laboratories). The datasets were indexed, integrated, and scaled using HKL-2001 (48). The structures were determined by molecular replacement (MR) using Phase-MR2 from the PHENIX Suite of program (49). One monomer of the RPRD1A-CID (PDB: 4JXT) was used as a search model for the initial phases. Structure refinement was performed using phenix.refine along with iterative model building in COOT (50). TLS parameters were included in the refinement of all structures. The final structures were evaluated after refinement using MolProbity (51). The refinement statistics for the structures are summarized in table S5. All figures were prepared with PyMol (The PyMOL Molecular Graphics System, version 1.8, Schrödinger LLC).

Phosphorylation sample preparation

Kinase reactions were performed in a buffer containing 2 mM adenosine 5′-triphosphate, 50 mM Tris (pH 8.0), and 10 mM MgCl2 and supplemented with the CTD substrate (1 mg/ml) for 15 hours. Reactions were initiated by adding 0.6 μM Dyrk1a. The reaction time was optimized so that no further phosphorylation occurred on the substrate. Reactions were quenched with the addition of 10 mM EDTA.

Label-free proteomics sample preparation and CTD affinity purification

Dyrk1a (0.6 μM) was used to phosphorylate the 26x yeast GST-CTD substrate (1 mg/ml) in a 100-μl reaction for 15 hours. Likewise, a 26x yeast GST-T4E CTD substrate was incubated in a similar manner without any kinase treatment. Glutathione Agarose beads were washed thrice in buffer C [20 mM Tris (pH 8.0), 150 mM NaCl, and 10 mM BME], and the treated GST-CTD samples were added to the beads and incubated overnight. A total of 200 million HEK293 cells were grown, collected, and the cell pellet was resuspended in buffer A [10 mM Hepes (pH 7.4), 100 mM NaCl, 300 mM sucrose, 3 mM MgCl2, 0.5% Triton X-100, 1:100 protein, and phosphatase inhibitor]. Cells were then vortexed, incubated on ice for 15 min, and centrifuged at 15,000g for 10 min at 4°C. The supernatant is discarded, and the cell pellet was resuspended in buffer B [10 mM Tris (pH 8.0), 150 mM NaCl, and 1:100 protease and phosphatase inhibitors (PPI)] supplemented with 1:1000 benzonase. This mixture was incubated at room temperature for 1 hour and centrifuged at 15,000g for 10 min. The supernatant was collected as the nuclear fraction. After overnight incubation, the GST-CTD bound beads were washed twice with buffer C and once with buffer B. The nuclear fraction was added to the substrate-bound beads and incubated at 4°C overnight. Then, the beads were centrifuged at 4000g for 2 min at 4°C. The beads were washed twice with low salt buffer [20 mM Tris (pH 8.0), 150 mM NaCl, 10% glycerol, 0.1% Triton X-100, and 1:100 PPI] for 5 min per wash and thrice with high salt buffer [20 mM Tris (pH 8.0), 500 mM NaCl, 10% glycerol, 0.1% Triton X-100, and 1:100 PPI]. To the beads, 100 μl of elution buffer was added and spun at 4°C for 2 hours. Then, the beads were centrifuged at 4000g for 2 min at 4°C, and the supernatant was collected for the pulldown.

Pulldown samples were exchanged into 5 mM tris-HCl using 3-kDa Amicon filters. Samples were then denatured in 2,2,2-trifluoroethanol and 5 mM tris(2-carboxyethyl)phosphine at 55°C for 45 min. Proteins were alkylated in the dark with 5.5 mM iodoacetamide, and the remaining iodoacetamide was quenched with 100 mM dithiothreitol. MS-grade trypsin was then added to the solution at an enzyme:protein ratio of 1:50, and the digestion reaction was incubated at 37°C for 4 hours. Trypsin was quenched by adding 10% formic acid, and the volume was reduced to 500 μl in a vacuum centrifuge. Samples were then filtered using a 10-kDa Amicon filter and desalted using Pierce C18 tips (Thermo Fisher Scientific). The samples were resuspended in 95% water, 5% acetonitrile, and 0.1% formic acid prior to MS.

Proteomics MS and protein identification

Peptides were separated on an Acclaim PepMap100 C-18 column (75 μM × 25 cm; Thermo Fisher Scientific) using a 5 to 50% acetonitrile + 0.1% formic acid gradient for 120 min and analyzed online by nanoelectrospray ionization tandem MS on a Thermo Fisher Scientific Fusion Tribrid Orbitrap mass spectrometer, using a data-dependent acquisition strategy and analyzing two biological replicates per sample. Full precursor ion scans (MS1) were collected at a high resolution (120,000). MS2 scans were acquired in the ion trap in rapid scan mode using the Top Speed acquisition method and fragmenting by collision-induced dissociation. Dynamic exclusion was activated with a 60-s exclusion time for ions selected more than once.

Proteins were identified with Proteome Discoverer 2.3 (Thermo Fisher Scientific), searching against the UniProt human reference proteome. Methionine oxidation [+15.995 Da], N-terminal acetylation [+42.011 Da], N-terminal methionine loss [−131.04 Da], and N-terminal methionine loss with the addition of acetylation [−89.03 Da] were all included as variable modifications. Peptides and proteins were identified using a 1% FDR.

To score changes in protein abundance, a z score was estimated between the unmodified control and the kinase-treated sample for each protein as in (52). To generate volcano plots, datasets from both replicates were log2 transformed, missing values were imputed using fancyimpute version 0.7.0, and data were quantile normalized. Enriched proteins were defined using a P value of 0.05. P values in volcano plot analyses were calculated using a two-tailed, two-sample t test.

Fluorescence polarization

CTD peptides with double repeats were labeled with fluorescein isothiocyanate (FITC) and purchased from Biomatik. Protein and peptide concentrations were determined according to their absorbance at 280 nm. Fluorescence polarization values were collected on a Tecan F200 plate reader in a buffer [50 mM Tris (pH 8.0), 300 mM NaCl, and 10 mM BME] at room temperature. Samples were excited with vertically polarized light at 485 nm and at an emission wavelength of 535 nm. RPRD1B-CID, RPRD1A-CID, RPRD2-CID, and SCAF4-CID protein was titrated into a reaction mixture containing a buffer supplemented with 10 nM FITC-peptide. Measurements were taken in triplicate, and the experimental binding isotherms were analyzed in GraphPad Prism v9 using a total binding mode to obtain Kd values.

Reverse transcription qPCR

Total RNA was harvested from HEK293 or HEK293T cells using the DirectZol RNA Miniprep kit (Zymo Research, Irvine, CA, United States, product no. R2050). cDNA was generated using the AzuraQuant cDNA synthesis kit (Azura Genomics) following the manufacturer’s instructions. qPCR was done using the AzuraQuant Green Fast qPCR Mix Lo-Rox (Azura Genomics) in a ViiA-7 Real Time PCR system (Applied Biosystems). All qPCR experiments were conducted in biological triplicate; error bars represent means ± SEM. Relative gene expression was assessed using the ∆∆Ct method normalized to actin β (ACTB) expression. Student’s t test was used to compare groups. All primers used in this study can be found in table S7.

Differential scanning fluorometry

Purified recombinant RPRD1B-CID at a final concentration of 5 μM was incubated with 10X SYPRO Orange (Molecular Probes) in a 96-well low-profile PCR plate (ABgene, Thermo Fisher Scientific), and fluorescence was captured in a LightCycler 480 (Roche). Protein melting curves were carried out with a temperature acquisition mode using a total of 10 acquisitions per 1°C in each cycle from 20° to 95°C. The melting temperature was derived using the Boltzmann equation.

RNA isolation, library preparation, and RNA sequencing

Total RNA was isolated from HEK293T cells (at least ~106 cells per sample) using the DirectZol RNA Miniprep kit (Zymo Research). RNA integrity was assessed by Novogene Co. using the RNA Nano 6000 assay kit of the Bioanalyzer 2100 system (Agilent Technologies, CA, United States). Libraries were prepared at Novogene Co. according to the manufacturer’s instructions for the NEBNext Ultra RNA Library Kit for Illumina. The resulting libraries tagged with unique dual indices were checked for size and quality using the Agilent Bioanalyzer 2100. Libraries were loaded for sequencing on the NovaSeq 6000 (Illumina, San Diego, CA, United States) instrument (paired-end 2X150).

Analyses of RNA-seq data and APA

Quality of raw reads was assessed using FastQC read quality reports (https://usegalaxy.org) (53). Adapter Illumina sequences were trimmed off by Trimmomatic v.0.38 with default parameters (54). Next, reads were aligned to a human reference genome, GRCh38 version, using HISAT2 fast aligner v.2.2.1 with default parameters and –unstranded (55). The Gencode v38 gtf file was used as annotation gtf. Last, mapped fragments were quantified by featureCounts v.2.0.1 in Galaxy (56). Differential expression was analyzed using edgeR v.3.36.0; genes with FDR < 0.05 were considered as differentially expressed (57). RNA sequencing (RNA-seq) data were deposited in Gene Expression Omnibus (GEO) under the accession number GSE262702. Quantification of differential APA usage was performed using LABRAT (43). Tffasta was used to filter transcripts that have ill-defined 3′ ends, and the last two exons of each transcript were extracted. For --librarytype, RNA-seq was chosen. The 3′ ends were then quantified using Salmon. Calculatepsi was used to calculate the relative usage of these ends, compare across conditions, and Ψ values were calculated for each gene in each sample with an expression level cutoff of 5 Transcripts Per Million (TPM). Enrichment analysis of biological processes was performed with ShinyGO v.0.80 (58). ARE search in sequences of select subset of genes preferring distal poly(A) sites (Ψ52xT4A-Ψ52xWT > 0) was performed using the ARED-Plus database (44).

Analyses of ASEs

rMATS turbo v.4.1.2 was used for detection of five major alternatively spliced events upon 52X T4A vs. 52X WT expression (with parameters, --libType set to unstranded, FDR < 0.05; ILD ≥ 10%) (46). As input files for rMATS, alignment .bam files from HISAT2 mapper and gencode v38 annotation gtf were used.

ChIP and ChIP-seq

To generate 52X T4A and 52X WT CTD for ChIP studies, transient transfection of 12 μg of either plasmid was performed using PEI. Following transfection, α-amanitin (2.5 μg/ml) was added to cells for 48 hours. Briefly, for HA-tagged proteins, HEK293 cells were double cross-linked with 2 mM disuccinimidyl glutarate (DSG) for 15 min followed by secondary fixation with 1% formaldehyde for 10 min at room temperature. Single cross-linking was used for RPB1 ChIP using 1% formaldehyde for 10 min. Cross-linking was quenched with 0.125 M glycine for 5 min. Cells were successively lysed in lysis buffer LB1 [50 mM Hepes-KOH (pH 7.5), 140 mM NaCl, 1 mM EDTA, 10% glycerol, 0.5% NP-40, 0.25% Triton X-100, and 1× PPI), LB2 [10 mM tris-HCl (pH 8.0), 200 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, and 1× PPI], and LB3 [10 mM tris-HCl (pH 8.0), 100 mM NaCl, 1 mM EDTA, 0.5 mM EGTA, 0.1% Na deoxycholate, 0.5% N-lauroylsarcosine, and 1× PI]. Chromatin was sonicated to an average size of ~200 to 500 bp using a UCD-200 Biorupter (30-s on and 30-s off for 30 min). A total of 5 μg of HA antibody (catalog no. C29F4), pThr4 antibody (Active Motif, catalog no. 61361), or pSer2 antibody (Sigma-Aldrich, SKU: MABE953) was premixed in a 50-μl volume of Dynabeads Protein A or Protein G (Invitrogen) and was added to each sonicated chromatin sample and incubated overnight at 4°C. For pThr4 samples, the Ssu72/symplekin complex (55 μM) was added to sonicated chromatin and incubated at 28°C for 30 min before immunoprecipitation. The chromatin-bound beads were washed two times with low salt buffer [0.1% Na deoxycholate, 1% Triton X-100, 1 mM EDTA, 50 mM Hepes (pH 7.5), and 150 mM NaCl], once with high salt wash buffer [0.1% Na deoxycholate, 1% Triton X-100, 1 mM EDTA, 50 mM Hepes (pH 7.5), and 500 mM NaCl], once with LiCl wash buffer [250 mM LiCl, 0.5% NP-40, 0.5% Na deoxycholate, 1 mM EDTA, and 10 mM tris-HCl (pH 8.0)], and twice in Tris-EDTA (TE) buffer. The chromatin was reverse cross-linked overnight at 65°C with shaking at 750 rpm. After DNA extraction using phenol-chloroform, the DNA was resuspended in 10 mM tris-HCl (pH 8.0). The purified DNA was subjected to qPCR to confirm target region enrichment before moving on to deep sequencing library preparation. For sequencing, the extracted DNA was used to construct the ChIP-seq library using the NEBNext Ultra II DNA Library Prep Kit followed by sequencing with an Illumina NovaSeq X Plus system. For pThr4 datasets, libraries were sequenced with an Illumina HiSeq 3000.

Analysis of ChIP-seq data

After the initial assessment of read quality, pThr4 (untreated samples), pSer2, and RPRD1B (HA tag) ChIP-seq data were mapped onto a human reference genome, hg38, with a Bowtie2 v. 2.5.0 aligner for paired-end reads using default parameters (59). pThr4 (treated with Ssu72) single-end ChIP-seq reads were mapped onto a human reference genome, hg19, using BWA v.0.7.17 (60). Coverage tracks in .bigwig format were generated from filtered.bam files (mapq > 20) and visualized in the IGV v.2.4.16 software (61).

After alignment, MACS2 v.2.2.7.1 in Galaxy (parameters: --broad; --broad-cutoff of q < 0.1 for pThr4) was used to call peaks for immunoprecipitation samples against an input (62). Bioconductor R package “chipseeker” v.1.18.0 was used for deriving the consensus pThr4-Ssu72 peakset and peak annotation using gencode hg19 gtf as a reference (63). For peak annotation, promoters were defined as (−1000 bp, +1000 bp from TSS) regions. The Gencode v38 gtf file was used as the annotation gtf for pSer2 and RPRD1B data. TSS/TES profiling was done using plotProfile on matrices generated with 50-bp bins using the computeMatrix function from the deeptools v.2.2.3 (64). Calculation of TSS/TES ratios was performed using values derived with --outFileNameMatrix parameter of computeMatrix function and custom R script. The TSS and TES for ratio estimation were defined as values of the bin with maximal signal in the first (bins 1 to 60) and second (bins 61 to 120) halves of the profile, respectively. Reproducibility of ChIP-seq replicates was assessed by Pearson correlation analysis using the plotCorrelation function and/or binding affinity heatmaps in the DiffBind R package (64, 65). ChIP-seq data were deposited in GEO under the accession number GSE262826.

Single-end ChIP-seq data from PMID: 22549466 (samples GSM920945-pThr4 ChIP, GSM920947-Pol II ChIP in consensus T48_control, GSM920949-Pol II ChIP in T4A mutant, input) were reanalyzed using the following steps. After the initial QC step, raw reads were trimmed using default parameters of -trim-galore (v.0.6.3). BWA v.0.7.17 was used to align reads onto a reference genome (hg19). After filtering out low-quality alignments (mapq < 20), the “MarkDuplicates” tool from Picard v.2.18.2 was applied to reduce duplication levels. TSS/TES profiling was done using matrices generated with 50-bp bins using the computeMatrix function from the deeptools.

Statistical analyses

Statistical analyses were performed using RStudio v4.0.5 and GraphPad Prism v9. Two-tailed, independent sample t test was used for comparing the two groups (if not stated otherwise). P < 0.05 was considered as significant. Correlations were assessed using two-tailed Pearson r coefficients. Protein bands were quantified and compared using the ImageJ software.

Acknowledgments

We thank the UT Austin Center for Biomedical Research Support Biological Mass Spectrometry Facility [Research Resource Identifiers (RRID): SCR_021728] and the Center for Biomedical Research Support Microscopy and Imaging Facility (RRID: SCR_021756) for granting access to their equipment for conducting proteomics and IF experiments, respectively. We also thank the Advanced Photon Source (BL23-ID-D) where crystallographic data collection was performed.

Funding: This work was supported by the National Institutes of Health grant R01GM104896 (Y.J.Z.), National Institutes of Health grant R01GM125882 (Y.J.Z.), National Institutes of Health grant R35GM148356 (Y.J.Z.), National Institutes of Health grant R35GM122480 (E.M.M.), Welch Foundation grant F-1515 (E.M.M.), and Army Research Office grant W911NF-12-1-0390 (E.M.M.). We also thank funding from L. Leon Campbell Professorship fund.

Author contributions: Conceptualization: R.Y.M., S.I., and Y.J.Z. Methodology: R.Y.M., S.B.P., Q.Z., and Y.J.Z. Investigation: R.Y.M., S.B.P., S.I., H.A.H., R.S., and B.F. Visualization: R.Y.M. and S.B.P. Supervision: Y.J.Z. and E.M.M. Writing—original draft: R.Y.M., S.B.P., Q.Z., and Y.J.Z. Writing—review and editing: R.Y.M., S.B.P., Q.Z., and Y.J.Z.

Competing interests: The authors declare that they have no competing interests.

Data and materials availability: The MS proteomics data generated in this study have been deposited in the ProteomeXchange Consortium via the PRIDE partner repository under accession code PXD050595. The RNA-seq data used in this study are available in GEO under accession code GSE262702. The ChIP-seq data used in this study are available in GEO under accession code GSE262826. The x-ray crystal structure was deposited in PDB under accession code 9B9L. Blots were deposited to Mendeley Data under DOI: 10.17632/y2jwsy43hs.1. All data needed to evaluate the conclusions in the paper are present in the paper and/or the Supplementary Materials.

Supplementary Materials

The PDF file includes:

Figs. S1 to S6

Tables S6 and S7

Legends for tables S1 to S5

Other Supplementary Material for this manuscript includes the following:

Tables S1 to S5
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