Regulation of ATR activity by the RNA polymerase II phosphatase PNUTS-PP1

Ataxia telangiectasia mutated and Rad3-related (ATR) kinase is a key factor activated by DNA damage and replication stress. Here, we show that ATR signaling is increased in human cells after depletion of the RNAPII phosphatase PNUTS-PP1, which dephosphorylates RNAPII on Ser 5 of its carboxy-terminal domain (CTD) (pRNAPII S5). Increased ATR signaling was observed in the presence and absence of ionizing radiation or replication stress and even in G1 phase after depletion of PNUTS. Vice versa, ATR signaling was reduced, in a PNUTS dependent manner, after inhibition of the CDK7 kinase mediating pRNAPII S5. Furthermore, CDC73, a well-known RNAPII-CTD binding protein, was required for the high ATR signaling after depletion of PNUTS and co-immunoprecipitated with RNAPII and ATR. These results suggest a novel pathway involving RNAPII, PNUTS-PP1 and CDC73 in ATR signaling and give new insight into the diverse functions of ATR.


Abstract:
Ataxia telangiectasia mutated and Rad3-related (ATR) kinase is a key factor activated by DNA damage and replication stress. Here, we show that ATR signaling is increased in human cells after depletion of the RNAPII phosphatase PNUTS-PP1, which dephosphorylates RNAPII on Ser 5 of its carboxy-terminal domain (CTD) (pRNAPII S5). Increased ATR signaling was observed in the presence and absence of ionizing radiation or replication stress and even in G1 phase after depletion of PNUTS. Vice versa, ATR signaling was reduced, in a PNUTS dependent manner, after inhibition of the CDK7 kinase mediating pRNAPII S5. Furthermore, CDC73, a well-known RNAPII-CTD binding protein, was required for the high ATR signaling after depletion of PNUTS and co-immunoprecipitated with RNAPII and ATR. These results suggest a novel pathway involving RNAPII, PNUTS-PP1 and CDC73 in ATR signaling and give new insight into the diverse functions of ATR. 5 crucial player in the G2 checkpoint, we addressed whether PNUTS-PP1 might suppress ATR signaling. Our results show that ATR signaling increases after PNUTS depletion in a manner not simply correlating with DNA damage or replication stress. The increased ATR signaling rather appears to depend upon CTD phosphorylation, which is counteracted by PNUTS-PP1. Furthermore, the known CTD-binding protein, CDC73, is required for the high ATR signaling and co-immunoprecipitates with ATR.

Results:
In our previous work (Landsverk et al., 2010), we observed increased phosphorylation of CHK1 and RPA32 at late timepoints (2-24 hr) after IR in PNUTS depleted HeLa cells.
As CHK1 and RPA32 are ATR targets (Reaper, Griffiths et al., 2011, Shiotani, Nguyen et al., 2013, we addressed whether ATR signaling was affected specifically. Indeed, depletion of PNUTS with two different siRNA oligos caused increased IR-induced phosphorylation of the ATR substrates CHK1 S317 and RPA S33, but not of the ATM substrate CHK2 T68 ( Fig 1A). Phosphorylation of CHK1 and RPA were increased both at early (5min-1h) and late (6h) timepoints after IR, as well as in the absence of IR (Fig 1A), suggesting a general role for PNUTS in suppressing ATR signaling. In agreement with this notion, pCHK1 S317 and pRPA S33 were higher also during thymidine-induced replication stress in PNUTS depleted cells (Fig 1B). Similar results were found in U2OS cells (Fig S1A), and the effect was clearly ATR-mediated, as the ATR inhibitor VE-821 inhibited the increased CHK1 phosphorylation after IR and thymidine (Fig S1B,C). Inhibition of ATR activity was not a general effect after depletion of a PP1 regulatory subunit because knockdown of another abundant nuclear regulatory subunit, NIPP1 (Jagiello et al., 1995), did not increase CHK1 S317 or RPA S33 phosphorylation ( Fig S1D). Furthermore, the increased ATR signaling was not due to off-target effects of the siRNA oligonucleotides, since expression of mouse pnuts-EGFP to near endogenous levels abrogated the increased CHK1 phosphorylation after depletion of human PNUTS, both in the absence and presence of IR ( Fig 1C).
To address the importance of PP1 for the inhibitory effects of PNUTS on ATR signaling, siRNA-resistant wild type and PP1-binding deficient PNUTS were overexpressed in cells depleted of endogenous PNUTS. Wild type PNUTS, but not PP1-binding deficient PNUTS RAXA (Kreivi et al., 1997), partially abrogated increased CHK1 phosphorylation in the absence of exogenous stress and after IR or thymidine (Fig 1D and S2A), showing that PP1-PNUTS binding is important for the negative effect of PNUTS on ATR signaling.
Higher expression levels of the PNUTS RAXA mutant did not alter these results ( Fig   S2B).
Potentially, PNUTS-PP1 could counteract ATR signaling by generally dephosphorylating ATR substrates, as is the case for Saccharomyces cerevisae PP4 and the ATR homologue Mec1 (Hustedt, Seeber et al., 2015). To address this, we added an ATR inhibitor after induction of ATR signaling by IR. If PNUTS-PP1 directly dephosphorylates CHK1 and RPA, depletion of PNUTS should cause delayed removal of pCHK1 S317/S345 and pRPA S33 after addition of the ATR inhibitor. However, both pCHK1 S317 and pRPA S33 declined at a similar rate in cells transfected with control siRNA and PNUTS siRNA (Fig   2A), showing that phosphatase activity against these substrates is similar under these conditions. Furthermore, overexpression of PNUTS did not decrease pCHK1 S317 or pRPA S33 (Fig 1D and data not shown). This suggests PNUTS-PP1 does not directly dephosphorylate these ATR targets. To further verify this finding, we also examined pCHK1 S317/S345 and pRPA S33 after addition of the ATR inhibitor to thymidine-treated cells transfected with control siRNA and PNUTS siRNA ( Fig S2C). Decline of pCHK1 S317 and pCHK1 S345 occurred similarly also under these conditions, consistent with the notion that CHK1 is not a direct substrate of PNUTS-PP1. On the other hand, pRPA S33 declined less in PNUTS depleted cells in the presence of thymidine ( Fig S2C). As pRPA S33 declined similarly in cells transfected with control and PNUTS siRNA after IR (Fig   2A), this most likely implies that another kinase contributes to pRPA S33 in PNUTS depleted cells after prolonged replication stress (thymidine 16h). ATR-independent phosphorylation of pRPA S33 has e.g. been reported in the presence of hydroxyurea (HU) in combination with ATR inhibitor (Toledo, Murga et al.). Altogether, these results nevertheless suggest that PNUTS-PP1 does not suppress ATR signaling by generally counteracting phosphorylation of its downstream substrates.
As the RNAPII CTD is the only known direct substrate of PNUTS-PP1 (Ciurciu et al., 2013, Lee et al., 2010, and RNAPII has a proposed role in ATR activation (Derheimer et al., 2007, Lindsey-Boltz & Sancar, 2007, we addressed whether dephosphorylation of RNAPII CTD is likely involved in the effects of PNUTS depletion on ATR signaling. We first verified that higher levels of pRNAPII S5 could be observed after depletion of PNUTS in HeLa cells ( Fig 2B). We next added THZ1, a specific inhibitor of CDK7, the kinase mediating phosphorylation of RNAPII S5 (CTD) (Heidemann, Hintermair et al., 2013, Kwiatkowski, Zhang et al., 2014, to control siRNA versus PNUTS siRNA transfected cells during thymidine-induced replication stalling. To allow a robust activation of ATR signaling before inhibition of CDK7, thymidine was added two hours prior to THZ1. Remarkably, both pRNAPII S5 and pCHK1 S317 were reduced upon addition of THZ1 to cells transfected with control siRNA (Fig 2C, lanes 11-13), and both pRNAPII S5 and pCHK1 S317 remained high in PNUTS depleted cells (Fig 2C, lanes 14-16). This strongly indicates that pCHK1 S317 depends on pRNAPII S5. Of note, while the ATR inhibitor VE822 reduced pCHK1 S317 equally in both PNUTS depleted and cells transfected with control siRNA (Fig S2C), the CDK7 inhibitor THZ1 only reduced pCHK1 S317 in the control siRNA transfected cells (Fig 2C), thus ruling out the possibility that THZ1 should directly inhibit ATR kinase.
The finding that pRNAPII S5 levels remained high in PNUTS depleted cells after THZ1 treatment (Fig 2C) is consistent with a major role of PNUTS-PP1 in mediating the dephosphorylation of this residue (Fig 2C, compare lanes 14-16 with lanes 11-13). However, we observed that pRNAPII S7 and pRNAPII S2 also remained higher in PNUTS depleted cells under these conditions, though the effects appeared weaker than observed for pRNAPII S5 (Fig S3A). Therefore, PNUTS-PP1 may also participate in direct dephosphorylation of pRNAPII S2 and/or S7, or, dephosphorylation of S2 and S7 may depend upon S5. Furthermore, this shows that high ATR signaling correlates with RNAPII CTD phosphorylation in general, rather than with pRNAPII S5 specifically under these conditions.
To confirm the correlation between ATR signaling and RNAPII CTD phosphorylation, we added THZ1 to IR-treated cells. Similarly as observed during replication stress, pRNAPII S5 and pCHK1 S317/S345 were reduced after THZ1 in cells transfected with control siRNA (Fig S3B). And again, pRNAPII S5 and pCHK1 S317/S345 remained high in cells depleted for PNUTS ( Fig S3B). An inhibitor of translation, cycloheximide, did not reduce pRNAPII S5 and pCHK1 S317/S345 after IR (Fig S3B), neither in control nor in PNUTS depleted cells, suggesting the effects of THZ1 on ATR signaling are independent of de novo protein production (via transcription and translation). To further explore the correlation between RNAPII CTD phosphorylation and ATR signaling, THZ1 was added prior to IR. As expected, pCHK1 S317 was suppressed by THZ1 in HeLa cells (Fig S3C).
Similar effects were obtained with two other transcription inhibitors which also caused reduced RNAPII CTD phosphorylation, 5,6-Dichloro-1-β-D-ribofuranosylbenzimidazole (DRB) and triptolide, but not by cycloheximide ( Fig S3C). Altogether these results support a link between RNAPII CTD phosphorylation and ATR signaling and indicate PNUTS-PP1 inhibits ATR activity by dephosphorylating pRNAPII CTD.
ATR is commonly known to be activated via replication stress, such as typically induced by hydroxyurea (HU). To compare ATR signaling in PNUTS depleted cells with ATR signaling after HU, we added different amounts of HU to non-transfected HeLa cells.
HeLa cells treated with 60-100 µM HU for 24 hr showed equal or higher levels of replication stalling compared to PNUTS depleted cells 48 hr after siRNA depletion, as measured by lower uptake of the nucleoside analog EdU ( Fig S4A). However, higher pCHK1 S317/S345 could still be observed in the PNUTS depleted cells (48 hr after siRNA transfection- Fig 3A), suggesting that ATR activity induced by depletion of PNUTS cannot simply be explained by replication stress. Notably, at 24 hr after siRNA transfection, there was no detectable difference in EdU uptake between control and siPNUTS transfected cells (Fig S4A), and therefore PNUTS depletion at 48 hours was comparable to 24 hours HU treatment.
We reasoned that phosphorylated RNAPII CTD might permit ATR activation even in the absence of replication e.g. in G1. To address this issue, cells in G1 and S phases of the cell cycle were sorted based on EdU incorporation and DNA content ( Fig 3B). Remarkably, pCHK1 S317 was higher in both G1 and S phase after depletion of PNUTS, with and without IR (Fig 3C). To validate the purity of the G1 population following sorting, thymidine, which specifically targets S phase cells, was added for 30 min after EdU labeling ( Fig   S4B). Induction of pCHK1 S317 and presence of cyclin A could only be detected in the S phase population (Fig S4B), confirming pure populations. These results support that ATR signaling is increased even in G1 phase following PNUTS depletion.
ATR is also well known to be activated by DNA damage, such as occurring after IR (Jazayeri, Falck et al., 2006). We therefore next compared PNUTS depleted cells with IRtreated control siRNA transfected cells to address whether the high ATR activity after PNUTS depletion could correlate with DNA damage. Higher levels of DNA damage markers pATM S1981, pDNAPK S2056, pCHK2 T68 and H2AX, but lower levels of pCHK1 S317, were observed in IR-treated control cells (1 and 6h after 10 Gy) compared to PNUTS depleted cells (Fig 3D,E). Furthermore, the lack of DNA damage signaling in PNUTS depleted cells was not caused by a reduced ability to activate ATM or DNAPK, as this occurred normally after IR ( Fig S4C). The high ATR activity in PNUTS depleted cells is therefore not likely caused by DNA damage.
Moreover, we assessed RPA loading onto chromatin, which can occur both in response to DNA damage and replication stress. To measure RPA loading we used a flow cytometry based assay similar to one previously shown to detect end resection (Ferretti, Himmels et al., 2016). Higher RPA loading, but lower CHK1 S317 phosphorylation, could be observed in control cells after 10 Gy compared to PNUTS depleted cells (Fig 3F compared to 3D). This suggests a lack of correlation between RPA loading and ATR signaling after depletion of PNUTS. To further investigate this, we also codepleted PNUTS with RPA70, an essential component of the RPA complex (Iftode, Daniely et al., 1999). As expected, this resulted in strongly reduced pRPAS33 (Fig S4D,E). However, ATR-dependent CHK1 S345 phosphorylation was not reduced in cells codepleted for PNUTS and RPA70 compared to control cells or cells depleted for RPA70 alone (Fig S4D,E). The increased ATR signaling following depletion of PNUTS is therefore likely not due to increased amounts of ssDNA-RPA.
We also addressed the involvement of other known key upstream ATR activating proteins, namely TOPBP1 and ETAA1. Though pCHK1 S345 was reduced, ATR-dependent pRPA S33 was not reduced in cells co-depleted for TOPBP1 and PNUTS compared to cells depleted for PNUTS alone, in the absence or presence of IR (Fig 4A-C). Notably, enhanced pRPA S33 after IR in cells co-depleted for PNUTS and TOPBP1 was dependent on PNUTS as depletion of TOPBP1 alone did not cause increased pRPA S33 (Fig 4B,C).
Conversely, upon co-depletion of PNUTS with ETAA1, pRPA S33 was reduced, but pCHK1 S345/S317 was not, compared to cells depleted of PNUTS alone (Fig 4D). Again the enhanced pCHK1 S317/S345 was dependent on PNUTS, as pCHKS317/S345 was not enhanced in cells depleted of ETAA1 alone compared to cells transfected with control siRNA ( Fig 4D). Triple depletion of PNUTS, ETAA1 and TOPBP1 suppressed both pCHK1 S317/S345 and pRPA S33 ( Fig 4D) in agreement with recent findings suggesting that TOPBP1 is required for pCHK1 S317/S345 and ETAA1 for pRPA S33 (Bass, Luzwick et al., 2016, Haahr, Hoffmann et al., 2016. Neither TOPBP1 nor ETAA1 therefore appear to be required for PNUTS dependent ATR signaling in general, but rather play essential downstream roles in the phosphorylations of CHK1 and RPA, respectively. To further characterize known ATR regulators following depletion of PNUTS, we compared their levels in cells transfected with PNUTS or control siRNA 24 or 48hr after siRNA transfection. Levels of ATR and ATRIP were not detectibly altered ( Fig S5A).
However, ETAA1, CLASPIN and TOPBP1 levels were increased in PNUTS depleted cells compared to control transfected cells, particularly at 48 hrs after siRNA transfection ( Fig   S5A). The co-depletions of PNUTS with ETAA1 or TOPBP1 nevertheless suggest that the ATR signaling can occur independently of either of these factors, though they are required for downstream phosphorylations (Fig 4). Also, as CLASPIN levels were downregulated, but pCHK1 S317 was higher after IR in PNUTS depleted cells relative to cells transfected with control siRNA (Fig S5B), this suggests CLASPIN is not either essential for enhanced ATR signaling upon PNUTS downregulation.
Our results showing a connection between RNAPII CTD phosphorylation and ATR signaling (Fig 2B,C and S3) suggest the CTD may be acting as a signaling platform for ATR activity. We therefore searched for factors that might participate in signaling from phosphorylated RNAPII CTD towards ATR. In the literature, we identified three proteins, BRCA1, PRP19 and CDC73, that associate with hyperphosphorylated RNAPII and have been linked to ATR (David, Boyne et al., 2011, Krum, Miranda et al., 2003, Marechal, Li et al., 2014, Phatnani, Jones et al., 2004, Poli, Gerhold et al., 2016, Turner, Aprelikova et al., 2004. We found that co-depletion of BRCA1 or PRP19 with PNUTS did not reduce the high ATR signaling (data not shown). However, co-depletion of CDC73 with PNUTS reduced both pCHK1 S317/S345 and pRPA S33, but not pRNAPII S5, in the presence or absence of IR ( Fig 5A). The reduction in pCHK1 S345 phosphorylation after co-depletion was observed with several siRNA oligos against CDC73 (four out of five) ( Fig S5C). Furthermore, expression of siRNA resistant Flag-CDC73 partially rescued the effects on pCHK1 S317/S345 and pRPAS33 downregulation after co-depletion of CDC73 with PNUTS ( Fig 5A), excluding siRNA off-target effects. These results suggest that CDC73 and PNUTS are acting in the same pathway for ATR activation and are consistent with a role for CDC73 in signaling from phosphorylated RNAPII CTD to ATR. CDC73 interacts genetically with the ATR homologue Mec1 in Saccharomyces cerevisiae, and a physical interaction has been proposed but not previously shown (Poli et al., 2016). To address whether CDC73 physically interacts with ATR and RNAPII, we performed co-immunoprecipitation (co-IP) experiments of endogenous proteins in HeLa cells.
Interestingly, PNUTS and PP1 were also detected in the CDC73 co-IPs (Fig 5B and S5D).
We verified that the immunoprecipitations were specific by using lysates from cells depleted of CDC73, which pulled down less ATR and RNAPII ( Fig S5D). Notably, RNAPII is a well known interactor of CDC73 ((Rozenblatt-Rosen, Hughes et al., 2005) and reviewed in (Van Oss, Cucinotta et al., 2017)). Furthermore, the depletion of CDC73 was only partial and significant amounts of CDC73 were present in the co-IPs from cells transfected with CDC73 siRNA (Fig S5D, CDC73-high exposure), which may explain the residual ATR and RNAPII pulled down under these conditions. To address whether ATR and pRNAPII S5 could physically associate, we performed ATR co-IPs. To enrich for active ATR in these experiments, we used pATR T1989 antibodies. This efficiently pulled down ATR and faint bands corresponding to pRNAPII S5 and RNAPII could also be detected, suggesting an interaction in live cells ( Fig 5C). All the co-IPs were performed after treatment with the endonuclease benzonase, strongly suggesting that the interactions were not mediated by DNA. Altogether these results support a role for phosphorylated RNAPII and CDC73 in the high ATR activity after PNUTS depletion.

Discussion:
ATR kinase plays a central role in signaling after DNA damage and replication stress.
Here we show for the first time that the RNAPII phosphatase PNUTS-PP1 suppresses ATR signaling. Our results suggest that ATR signaling is restrained by PNUTS-PP1 mediated dephosphorylation of RNAPII CTD, and thus support a role for RNAPII in ATR signaling. Furthermore, we have found that a well-known RNAPII binding factor, CDC73, is required for the high ATR signaling in PNUTS depleted cells. Moreover, our results support recent findings that TOPBP1 and ETAA1 may direct ATR activity towards different sub-strates. Altogether, based on these results we propose a new model for ATR signaling via RNAPII ( Fig 5D).
Notably, the high ATR signaling in PNUTS depleted cells does not correlate with DNA damage or replication stress and can occur even in G1 phase of the cell cycle (Fig 3).
Signaling to ATR by pRNAPII CTD may therefore be a more general event that simply responds to RNAPII stalling, in line with previous reports showing that perturbation of transcription can induce ATR activation in the absence of DNA damage and prior to detection of replication-stress (Derheimer et al., 2007, Kotsantis, Silva et al., 2016. The Saccharomyces cerevisae ATR homologue Mec1 was shown to promote removal of RNAPII at sites of transcription-replication conflict (Poli et al., 2016). Viewed in light of our results, ATR activity at sites of stalled transcription might thus promote removal of RNAPII, even in the absence of replication. Removal of stalled RNAPII is likely important also outside of S phase, because RNAPII could create an obstacle for further transcription in a region which might e.g. contain an essential-or tumor suppressor gene. In agreement with prolonged RNAPII stalling being detrimental to the cell, it has been shown to be a strong signal for apoptosis (Ljungman & Zhang, 1996).
However, as RNAPII CTD phosphorylation occurs during the normal transcription cycle, where e.g. pRNAPII S5 is required for 5' mRNA capping (Egloff et al., 2012), a strict linear correlation may be unlikely as it would imply ATR activation merely as a consequence of normal transcription. We suggest that the level of pRNAPII CTD at individual sites may determine the degree of ATR activation by RNAPII. Perhaps ATR is activated only when pRNAPII CTD reaches an abnormal threshold level at an unexpected site. Supporting this, RNAPII CTD phosphorylation is often associated with RNAPII stalling, and can occur within the entire transcribed region (Alexander et al., 2010, Boehm et al., 2003, Munoz, Perez Santangelo et al., 2009. Furthermore, other post-translation modifications on the CTD (Harlen & Churchman, 2017) may contribute to fine-tune ATR activity via RNAPII.
Of note, in the alternative splicing response to UV, pRNAPII CTD was proposed to occur downstream of ATR activation, and ATR activation to occur independently of transcription in HaCaT cells (Munoz, Nieto Moreno et al., 2017). These results may appear to be contradictory to ours. However, we did not detect any reduction in pRNAPII S5 after ATR inhibitor during replication stress in HeLa cells (Fig S2C) suggesting ATR is not always upstream of pRNAPII CTD. Furthermore, the differing results may be explained by the existence of several pathways for ATR activation acting in parallel, e.g. via RNAPII, via ssDNA-RPA, and via unknown pathways. The contribution from each pathway is likely to vary between cell types and with different stresses.
Remarkably, following depletion of PNUTS ATR signaling did not correlate with RPA loading and was not reduced by RPA70 co-depletion (Fig 3D,F and S4D,E). These results are in agreement with previous studies showing that RPA loading and ATR activation do not always correlate (Dodson, Shi et al., 2004, Kousholt, Fugger et al., 2012. However, in our co-depletion experiments, we cannot exclude that very small amounts of ssDNA-RPA (less than 5% of RPA70 was remaining after siRNA depletion (Fig S4D)) may contribute to the ATR activity. Indeed, recently, ATR activation was shown to occur at centromeres in mitosis, by a proposed mechanism involving R-loops and ssDNA-RPA which can subsequently lead to small amounts of ssDNA-RPA (Nguyen, Yadav et al., 2017), it is thus tempting to speculate that depletion of PNUTS may cause small amounts of ssDNA-RPA via R-loops, and thereby enhance ATR signaling. On the other hand, there is an intimate connection between stalled RNAPII and R-loops (Santos-Pereira & Aguilera, 2015). Interestingly, it was recently shown that overexpression of RNAse H can also cause release of stalled RNAPII, suggesting that R loops can promote RNAPII stalling (Sridhara, Carvalho et al., 2017). Another speculation might therefore be that R-loops could activate ATR by leading to stalling of RNAPII and subsequent RNAPII CTD phosphorylation.
Our results point to a new role for CDC73 upstream of ATR activation. CDC73 is a component of the PAF1 complex, including PAF1, CTR9, LEO1, RTF1 and WDR61, involved in all stages in RNAPII transcription (Van Oss et al., 2017). However, the PAF1 complex does not appear to be essential for transcription as depletion of e.g. CDC73 was found to both up and down-regulate mRNA expression (Rozenblatt-Rosen, Nagaike et al., 2009). In Saccharomyces cerevisae the PAF1 complex was required for removal of RNAPII during collisions of transcription and replication (Poli et al., 2016). Leo1 is a target of Mec1, and thus Mec1 has been thought to primarily have an upstream role in triggering the removal of RNAPII via phosphorylation of Leo1 and the Ino80 chromatin remodeling complex (Hustedt et al., 2015, Poli et al., 2016. However, as our results point to a role for CDC73 in ATR activation, this may suggest the existence of a feedback loop, whereby stalling of RNAPII in head-on collisions between the transcription and replication machineries leads to activation of ATR via CDC73, and the resultant ATR activity leads to removal of RNAPII. Interestingly, our results fit well with the observation that head-on collisions, which cause disruption of RNA transcription, lead to ATR activity, whereas codirectional collisions, which do not disrupt RNA transcription, do not (Hamperl, Bocek et al., 2017). CDC73 is also a well known tumor suppressor gene. It is currently not clear how CDC73 acts as a tumor suppressor, though roles in Wnt signaling, regulation of P53 and CYCLIN D levels and homologous recombination repair have been suggested (Herr, Lundin et al., 2015, Jo, Chung et al., 2014, Mosimann, Hausmann et al., 2006, Woodard, Lin et al., 2005. ATR activity protects genome integrity by stabilizing stalled forks during replication stress and promoting DNA repair and checkpoint activation (Yazinski & Zou, 2016). However, in addition ATR activity can promote apoptosis in non-cycling cells, which are the majority of cells in humans (Kemp & Sancar, 2016). Therefore, CDC73 could potentially protect against cancer by promoting RNAPII-mediated ATR activity leading to cell death in non-cycling cells with DNA damage. Consistent with this interpretation, PNUTS, which counteracts CDC73 in ATR activation, is a putative proto-oncogene (Kavela, Shinde et al., 2013).

Materials and methods:
Cell culture and treatments: Human epithelial HeLa and human osteosarcoma U2OS cells were grown in Dulbecco's Modified Eagle Medium (DMEM) containing 10% fetal calf serum (Life Technologies). The cell lines were authenticated by short tandem repeat profiling using Powerplex 16 (Promega) and regularly tested for mycoplasma contamination. HeLa BAC cells stably expressing EGFP mouse pnuts were a generous gift from the laboratory of Tony Hyman (GeneTex) and anti-ETAA1 (Haahr et al., 2016). Peroxidase-conjugated secondary antibodies were from Jackson Immunoresearch. Blots were imaged in a Chemidoc MP (BioRad) using chemiluminescence substrates (Supersignal west pico, dura or femto; Thermo Scientific). Quantifications were performed and images processed in Image Lab 4.1 (BioRad) software. Linearity of detection was verified by including a dilution series of one of the samples (see e.g. Figure 1B) and excluding saturated signals. The resulting standard curve allowed accurate quantification. To blot for total protein after detection of a phosphorylated protein, membranes were stripped using ReBlot Plus Mild Antibody Stripping Solution (Millipore).

Statistics:
All experiments, except when otherwise stated, were performed three times or more.