Phosphorylation of Human Rad9 Is Required for Genotoxin-activated Checkpoint Signaling*

Pia Roos-Mattjus a b, Kevin M. Hopkins c, Andrea J. Oestreich d, Benjamin T. Vroman e, Kenneth L. Johnson f, Stephen Naylor a f g h, Howard B. Lieberman c and Larry M. Karnitz d e g i j

From the aDepartment of Biochemistry and Molecular Biology, the gDepartment of Molecular Pharmacology and Experimental Therapeutics, the dProgram in Tumor Biology, the Divisions of eDevelopmental Oncology Research and iRadiation Oncology, and the fBiomedical Mass Spectrometry and Functional Proteomics Facility, Mayo Clinic and Foundation, Rochester, Minnesota 55905 and the cCenter for Radiological Research, Columbia University, New York, New York 10032

Received for publication, February 13, 2003 , and in revised form, March 27, 2003.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Rad9, a key component of genotoxin-activated checkpoint signaling pathways, associates with Hus1 and Rad1 in a heterotrimeric complex (the 9-1-1 complex). Rad9 is inducibly and constitutively phosphorylated. However, the role of Rad9 phosphorylation is unknown. Here we identified nine phosphorylation sites, all of which lie in the carboxyl-terminal 119-amino acid Rad9 tail and examined the role of phosphorylation in genotoxin-triggered checkpoint activation. Rad9 mutants lacking a Ser-272 phosphorylation site, which is phosphorylated in response to genotoxins, had no effect on survival or checkpoint activation in Mrad9–/– mouse ES cells treated with hydroxyurea (HU), ionizing radiation (IR), or ultraviolet radiation (UV). In contrast, additional Rad9 tail phosphorylation sites were essential for Chk1 activation following HU, IR, and UV treatment. Consistent with a role for Chk1 in S-phase arrest, HU- and UV-induced S-phase arrest was abrogated in the Rad9 phosphorylation mutants. In contrast, however, Rad9 did not play a role in IR-induced S-phase arrest. Clonogenic assays revealed that cells expressing a Rad9 mutant lacking phosphorylation sites were as sensitive as Rad9–/– cells to UV and HU. Although Rad9 contributed to survival of IR-treated cells, the identified phosphorylation sites only minimally contributed to survival following IR treatment. Collectively, these results demonstrate that the Rad9 phospho-tail is a key participant in the Chk1 activation pathway and point to additional roles for Rad9 in cellular responses to IR.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Cells respond to genotoxic stress by activating checkpoint signaling pathways that delay cell cycle progression, providing time to repair DNA damage, activate transcriptional responses and, if the damage is too severe, induce apoptotic pathways (16). The phosphatidylinositol 3-kinase-related kinases (PIKK)1 ATM and ATR are central components of the checkpoint signaling pathways (7). ATM and ATR are activated by genotoxins and phosphorylate downstream signaling proteins, including Chk1 and Chk2, two protein kinases that phosphorylate proteins that regulate checkpoint responses (810).

In addition to the PIKKs, Rad1, Hus1, Rad9, and Rad17 (using Schizosaccharomyces pombe nomenclature) are evolutionarily conserved proteins essential for Chk1 activation (1113). Rad9, Hus1, and Rad1 form a stable heterotrimeric complex, called the 9-1-1 complex (1417). Biochemical, biophysical, and molecular modeling studies predict that the 9-1-1 complex resembles PCNA (14, 16, 1921), a homotrimeric clamp that is loaded around DNA at primer-template junctions (22, 23). Loading of PCNA is carried out by the clamp loader replication factor C (p140-RFC), a heteropentameric complex consisting of 1 large subunit (p140) and four small subunits (24). The 9-1-1 complex also interacts with a putative clamp loader, the Rad17-RFC complex (25, 26), which is composed of Rad17, an RFC-like protein, and the four small RFC subunits (15, 19). In vitro, recombinant Rad17-RFC complex loads the 9-1-1 complex onto DNA (27), and in intact cells, multiple genotoxins induce the association of the 9-1-1 clamp with chromatin (28) in a process that requires Rad17 (12). Collectively, these observations support a model in which the Rad17-RFC clamp loader is a DNA damage sensor that loads the 9-1-1 clamp onto sites of DNA damage in the checkpoint signaling pathway (1, 3, 29, 30).

Recent studies demonstrated that ATR and the 9-1-1 complex collaboratively induce Chk1 activation. In both humans and Saccharomyces cerevisiae, the 9-1-1 complex associates with chromatin in a PIKK-independent manner following DNA damage (12, 31, 32). Likewise, ATR forms foci in the absence of Rad17 (12). Taken together, these results suggest that the 9-1-1 complex and ATR associate with DNA lesions independently of one another. Nonetheless, both the 9-1-1 complex and ATR are required for Chk1 activation (1113, 33), suggesting that these chromatin-bound complexes coordinately transduce a Chk1-activating signal in response to genotoxins; it remains unclear, however, how these protein complexes are regulated or how they cooperatively couple with the downstream signaling pathways.

One possible 9-1-1 regulatory mechanism is phosphorylation. Both Rad9 and Rad1 are phosphorylated in response to DNA damage (28, 3438). Because Rad9 binds DNA in the absence of ATR and pharmacologic inhibition of PIKKs does not affect Rad9 chromatin binding, it is unlikely that Rad9 phosphorylation regulates 9-1-1 chromatin binding (12, 39). Instead, phosphorylation of 9-1-1 complex members may regulate downstream signals that are activated by the clamp complex.

In the present study we examined the role of Rad9 phosphorylation in checkpoint signaling. We identified 9 phosphorylation sites in the carboxyl-terminal 119-amino acid Rad9 tail, which extends beyond the amino-terminal PCNA homology domains and is not required for interaction with Hus1 and Rad1 (14, 40). One of the sites we found was the previously identified Rad9 damage-inducible phosphorylation site Ser-272. We assessed the effects of Rad9 phosphorylation by expressing Rad9 mutants in Mrad9–/– embryonic stem (ES). These studies showed that cells expressing a Rad9 mutant in which Ser-272 was converted to Ala were indistinguishable from cells expressing wild-type Rad9. In contrast, the remaining Rad9 phosphorylation sites were essential for some but not all Rad9-dependent cellular responses. Collectively, these results demonstrate that the phospho-Rad9 tail plays a critical role in the transduction of downstream checkpoint signals.


    EXPERIMENTAL PROCEDURES
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Antibodies—Antibodies generated to human Rad9 (monoclonal and polyclonal), Rad1 (polyclonal), Hus1 (monoclonal), and Rad17 (polyclonal) have been described previously (28, 34). Phospho-Chk1 (P-Ser-345) and Chk1 (G-4) antibodies were purchased from Cell Signaling Technology (Beverly, MA) and Santa Cruz Biotechnology (Santa Cruz, CA), respectively, and were used according to the manufacturer's instructions. Anti-Chk2 antibodies (41) were a generous gift from J. Chen, (Mayo Clinic, Rochester, MN) and anti-phospho-Rad9 (P-Ser-272) antibodies (37) were a generous gift from E. Lee (University of Texas, San Antonio Texas). The p53 antibody (CM5) was purchased from Novocastra Laboratories, Ltd. (Newcastle upon Tyne, UK). The JNK1 (C-17) antibody was purchased from Santa Cruz Biotechnology.

Cell Culture and Cell Transfection—Human K562 erythroleukemia cells were cultured in RPMI 1640 (BioWhittaker, Walkersville, MD) containing 10% fetal bovine serum (Biofluids, Rockville, MD). K562 cells (1 x 107) were transiently transfected as previously described (34). Mouse ES cells were maintained on gelatinized tissue culture plates in knock-out DMEM (Invitrogen, Carlsbad, CA) containing 15% ES cell-qualified fetal bovine serum (Cell & Molecular Technologies, Inc., Phillipsburg, NJ), 0.1 mM non-essential amino acids, 2 mM L-glutamine (Invitrogen), 10–4 M 2-mercaptoethanol (Sigma) and 103 units/ml ES-GRO (Chemicon International, Temecula, CA). Cells were grown at 37 °C and 5% CO2. ES cells were transfected with LipofectAMINE 2000 according to the manufacturer's instructions (Invitrogen). Stable clones were selected in medium containing G418 (0.2 mg/ml) and characterized for expression levels by immunoblotting. Two independent clones of each mutant expressing similar levels of protein were used for further studies.

Derivation of Mrad9/ ES Cells—The Mrad9/ ES cells were originally created and characterized by Kevin M. Hopkins, Wojtek Auerbach, Xiang Yuan Wang, M. Prakash Hande, Haiying Hang, Debra J. Wolgemuth, Alexandra L. Joyner, and Howard B. Lieberman. Briefly, mouse ES cells were electroporated with a targeting vector (Mrad9neo/loxP) that, when integrated into the MRad9 locus, inserts loxP sites in the 5'-untranslated region and in the second intron. Following selection in 200 µg/ml G418 and gancyclovir to enrich for homologous targeting events, Southern blots and PCR were used to identify clones in which a single Mrad9 allele was targeted. One Mrad9neo/loxP ES cell line was cultured in 300 µg/ml G418 to select for cells that targeted the second Mrad9 allele, and a cell line with two targeted MRad9 alleles was identified by Southern blotting and PCR analyses. To delete the first and second Mrad9 exons via Cre-mediated recombination (42), the cells were transiently transfected with an HS-cre expression vector (43) and pPur (Clontech, Palo Alto, CA), and selected in puromycin. The clones were then examined by Southern blotting and PCR analyses to identify one in which the first and second exons of both Mrad9 alleles were deleted.

Drug and Radiation Treatments—Hydroxyurea (Sigma) was prepared fresh in phosphate-buffered saline (PBS). Cells were irradiated with a 137Cs source at a dose rate of ~10 Gy/min (Mark I, JL Shepherd and Associates, San Fernando, CA). For UV treatment, cells were washed with PBS to remove medium. Residual PBS was removed prior to irradiation with a UV-C irradiator (Spectrolinker XL-1000, Spectronics Corp., Westbury, NY). Fresh, warmed medium (37 °C) was added after UV irradiation.

Cell Extract Preparation and Analysis of Rad9 Chromatin Binding—To analyze protein expression levels, cells were lysed in 50 mM HEPES, pH 7.4, 1% Triton X-100, 10 mM NaF, 30 mM Na4P207, 150 mM NaCl, 1 mM EDTA, containing freshly added 10 mM {beta}-glycerophosphate, 1 mM Na3VO4, 10 µg/ml pepstatin A, 5 µg/ml aprotinin, 10 µg/ml leupeptin and 20 nM microcystin-LR for 10 min at 4 °C. For protein-protein interaction studies, cells were lysed in 150 mM KCl, 10 mM MgCl2, and 10 mM HEPES, pH 7.4, (supplemented with 100 µg/ml digitonin, 10 µg/ml aprotinin, 5 µg/ml pepstatin, 5 µg/ml leupeptin, 20 nM microcystin-LR, 1 mM Na3VO4, and 10 mM {beta}-glycerophosphate) for 10 min at 4 °C. All lysates were either immunoprecipitated as indicated or boiled with sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) sample buffer. Proteins were separated by SDS-PAGE, transferred to Immobilon P membranes (Millipore, Bedford, MA) and immunoblotted as described (34). Fractionation of the 9-1-1 complex into released and chromatin-bound fractions was done as described previously (28).

Rad9 Expression Vectors and Rad9 Mutagenesis—S-tag-Rad9-pIRES2-EGFP was generated by adding an in-frame S-tagTM (Novagen Inc.) to the amino terminus of human Rad9 using a PCR strategy. The resulting PCR product was cloned into pIRES2-EGFP (BD Sciences Clontech, Palo Alto, CA) using XhoI and BamHI. To generate stable ES cells expressing wild-type and mutant Rad9s, untagged human Rad9 was cloned into pIRES2-EGFP. Human Rad9 phosphodeficient mutants were generated by mutating the mapped phosphorylation sites using either the GeneEditor kit (Promega, Madison, WI) according to the manufacturer's instructions or by sequence-overlap extension (44). All mutants were sequenced to confirm mutations.

Purification of S-tagged Rad9 —A total of 2 x 109 K562 cells were transiently transfected by electroporation with S-tag-Rad9-pIRES2-EGFP (50 pooled transfections). For Edman radiosequencing analysis, cells were recovered by centrifugation 24 h after transfection, washed twice with phosphate-free media (ICN Biomedicals, Inc., Aurora, OH), and resuspended in 80 ml of phosphate-free media containing 5% fetal bovine serum and 5 mCi of [32P]orthophosphate (ICN Biomedicals, Inc.). Cells were cultured an additional 2 h, harvested, washed in ice-cold PBS, and lysed in buffer containing 10 mM HEPES, pH 7.4, 150 mM KCl, and 10 mM MgCl2 (containing freshly added 100 µg/ml digitonin, 10 µg/ml aprotinin, 5 µg/ml pepstatin, 5 µg/ml leupeptin, 20 nM microcystin-LR, 1 mM Na3VO4, and 10 mM {beta}-glycerophosphate) for 10 min at 4 °C. Following centrifugation, the supernatant was added to S-protein agarose beads (Novagen Inc.), and urea was added to a final concentration of 2 M. The mixture was incubated with rotation for2hat 4 °C. The beads were washed with cold lysis buffer (50 mM HEPES, pH 7.6, 1% Triton-X100, 10 mM NaF, 30 mM NaP2O7, 150 mM NaCl, 1 mM EDTA) supplemented with 2 M urea, 1 mM Na3VO4, and 10 mM {beta}-glycerophosphate. Proteins were released from the beads by heating in SDS-PAGE sample buffer and were resolved by SDS-PAGE. The gel was stained with Coomassie Blue, and the slowest migrating (most highly phosphorylated) Rad9 band was excised.

Radiosequencing of Rad9 —To identify the amino acid sequence of the radiolabeled peptides, the Coomassie Blue-stained band corresponding to fully phosphorylated, radiolabeled Rad9 was excised from the gel and cut into pieces no larger than 2 mm. Prior to digestion, the gel pieces were destained until clear, then reduced with dithiothreitol, and alkylated with iodoacetamide. In situ enzymatic digestion was performed overnight at 37 °C with modified sequencing-grade trypsin (Promega). Peptides were extracted from the gel, and a portion of the sample was separated on an Applied Biosystems 173A Microblotter Capillary HPLC System (Applied Biosystems, Foster City, CA) using a 0.5 x 150-mm C18 column with an acetonitrile gradient in the presence of 0.1% trifluoroacetic acid. The eluting peptides were spotted directly onto a strip of polyvinylidinedifluoride membrane. The strip was exposed to film to identify 32P-labeled peptides. Spots containing 32P-labeled peptides were excised, treated with Biobrene in methanol, and applied to either an Applied Biosystems Procise 492 HT or the Procise cLC Protein Sequencing System and run using pulsed liquid chemistry. These data were analyzed with the ABI Model 610A data analysis software (Applied Biosystems). To identify the positions of 32P-labeled amino acids within the radioactive peptides, the remainder of the sample was separated on the capillary HPLC exactly as described above. However, the radiolabeled fractions were instead collected manually and covalently bound to aryl amine-derivatized polyvinylidinedifluoride disks using the Sequelon-AA kit (Applied Biosystems), following the suggested instructions. The disks were chromatographed on an Applied Biosystem's Procise cLC modified to collect the derivatized amino acids released by each cycle directly into a 96-well microtiter plate. The extraction solvent of 90% MeOH/10% water/0.01% H3PO4 was substituted for n-butyl chloride in the S3 bottle on the sequencer. Each sequencer fraction, representing one Edman degradation cycle, was spotted onto a custom-made polystyrene plate and dried. The plate containing the released amino acids was counted either by direct counting using a Packard Matrix 96 counter (Packard Biosciences, Boston, MA) or by exposing the plate to a phosphor screen that was scanned with a Storm Phosphorimager (Molecular Dynamics, Piscataway, NJ).

Nano-scale Liquid Chromatography-Mass Spectrometry (nLC/MS) and Tandem Mass Spectrometry (nLC/MS/MS) of Rad9 —For mass spectrometric analysis, S-tag-Rad9 was purified as described above with the exception that the cells were not labeled with [32P]orthophosphate. After S-protein purification, one portion of the sample was left untreated, and the other portion was treated with {lambda}-protein phosphatase according to the manufacturer's instruction (New England Biolabs, Beverly, MA). The proteins were released from the beads by heating in SDS-PAGE sample buffer and were resolved by SDS-PAGE. The gel was stained with Coomassie Blue. Protein bands corresponding to dephosphorylated and fully phosphorylated Rad9 were excised. Gel slices containing phosphorylated and dephosphorylated Rad9 were destained, reduced with dithiothreitol, and alkylated with iodoacetamide. The gel was dried under vacuum and rehydrated in buffer containing 200 mM ammonium bicarbonate containing 0.2 mg/ml Zwittergent 3–16 and 40 ng/ml porcine trypsin (Promega) for 20 min. The gel slice was rinsed in 50 mM ammonium bicarbonate and incubated overnight at 37 °C in 50 mM ammonium bicarbonate. The next day the sample was sonicated, acidified with 10% trifluoroacetic acid, and sonicated again.

nLC/MS and nLC/MS/MS measurements were performed on aliquots of the tryptic digest using a Micromass Q-Tof-II mass spectrometer (Micromass, Manchester, UK). Reversed-phase nLC separations were performed on a 75-µm i.d. x 5-cm long PicoFit column packed with Aquasil C18 stationary phase (NewObjective, Inc., Cambridge, MA). Mobile phase A consisted of water/acetonitrile/n-propanol (98:1:1) containing 0.2% formic acid. Mobile phase B consisted of acetonitrile/n-propyl alcohol/water (80:10:10) containing 0.2% formic acid. A linear gradient from 0–50% B over 30 min was performed. The liquid chromatography pumping system (Michrom UMA, Michrom BioResources Inc., Auburn, CA) was operated at 50 µl/min and split to a column flow of 0.2 µl/min just prior to the sample introduction valve.

Peptides from both the phosphorylated and dephosphorylated samples were compared using Mass Lynx software (Micromass, Manchester, UK). Novel peptides were considered putative phosphopeptides if the mass of the tryptic peptide corresponded to the mass of Rad9 tryptic peptide plus 80 mass units per phosphate group. Peptides of the appropriate masses were subjected to MS/MS to identify phosphorylation sites. MS/MS experiments were performed using argon as the collision gas, and the collision energy was determined as a function of mass/charge (m/z) and charge (z). Sites of phosphorylation were determined by a combination of Sequest data base searching (ThermoFinnigan, San Jose, CA) and manual spectrum interpretation.

S-phase Checkpoint Assay—One day prior to the experiment, 1–2 x 104 cells were plated onto gelatinized 96-well plates. Cells were either left untreated or treated with IR (30 Gy) or UV-irradiated (20 J/m2), and incubated for 40 min at 37 °C following treatment. [methyl-3H]Thymidine (TRA120, Amersham Biosciences) was then added at 2 µCi/well, and the cells were incubated for an additional 20 min. Cells were released by trypsinization and harvested by transferring to glass filters. The filter-bound cells were lysed with distilled water (Skatron semiautomatic harvester, Skatron As., Lier, Norway). Filter-bound radioactivity was determined by liquid scintillation counting. [3H]Thymidine incorporation was calculated as the ratio of treated to control samples.

Cell Survival and Clonogenic Assays—Varying numbers of ES cells were plated on gelatinized 60-mm dishes in triplicate and 4 h later treated with the indicated doses of ionizing radiation or HU. HU was washed off with PBS 24 h later, and fresh medium was added. Cells were incubated for 14 days, stained with Coomassie Blue, and colonies were counted. Survival was calculated as the percentage of colonies in the untreated dishes compared with the treated dishes, taking into account the plating efficiency, using the equation: colonies counted/(cells plated)(plating efficiency).

For UV survival assays 1–2 x 104 cells were plated onto gelatinized 6-well plates. Plates were washed with PBS 24 h later. After removal of residual PBS, the cells were UV-irradiated (15 J/m2), and fresh media was added. Plates were washed 72 h later and stained with Coomassie Blue.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Mapping of Rad9 Phosphorylation Sites—Rad9 is extensively and constitutively phosphorylated even in cells that have not been exposed to exogenous genotoxins (3436). Rad9 is further inducibly phosphorylated in response to DNA damage (3438). Despite the extensive phosphorylation, the role of this Rad9 modification remains unclear. To identify Rad9 phosphorylation sites, we subjected Rad9 tryptic peptides to both Edman radiosequencing and mass spectrometry. To isolate sufficient quantities of Rad9 for these analyses, we transiently overexpressed S-tag-Rad9 in K562 cells. S-tag-Rad9 interacts with Hus1 and Rad1 and is converted to a chromatin-bound form following treatment with ionizing and UV radiation (39). Moreover, S-tag-Rad9 restores Chk1 activation in Rad9-deficient cells (data not shown). Collectively, these results demonstrate that S-tag-Rad9 has biochemical properties similar to endogenous Rad9.

Using Edman radiosequencing, we identified four radiolabeled peptides, and within these peptides we identified six phosphorylation sites in S-tag Rad9 (Fig. 1, A and B, Ser-272, Ser-277, Ser-328, Ser-375, Ser-380, and Ser-387). Using mass spectrometry, we identified five phosphorylated sites (Fig. 1, A and B). Two of these sites (Ser-277 and Ser-328) were also identified by radiosequencing. The three additional sites (Ser-336, Ser-341, and Thr-355) were new. All the phosphorylation sites were located within the carboxyl-terminal 119-amino acid Rad9 tail. This portion of Rad9 has no homology with PCNA, is not required for Rad9 to interact with Hus1 and Rad1 (14, 40), but is involved in nuclear transport of the protein due to the presence of a nuclear localization sequence (40). Collectively, the radiosequencing and mass spectrometric analyses mapped nine phosphorylation sites. Mutation of all nine sites to alanine blocked the Rad9 mobility shift observed by SDS-PAGE (data not shown and Fig. 2), indicating that these studies identified all the phosphorylation sites responsible for the mobility shift.



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FIG. 1.
Mapping of Rad9 phosphorylation sites. A, the amino-terminal two-thirds of Rad9 contains two predicted PCNA folds that form a PCNA-like structure. The carboxyl terminus, dubbed the tail, contains all the mapped Rad9 phosphorylation sites. B, Rad9 phosphorylation sites, their surrounding sequences, and as described under "Experimental Procedures" used to identify the site. C, three different mutants were generated: Ser-272 mutated to alanine (S272A), all mapped phosphorylation sites except for Ser-272 mutated to alanine (8A) and all mapped phosphorylation sites mutated to alanine (9A).

 


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FIG. 2.
Characterization of Rad9-expressing mouse ES cell lines. Mrad–/– ES cells were transfected with plasmids encoding wild-type Rad9, Rad9-S272A, Rad9-8A, and Rad9-9A. Stable cell lines expressing approximately equivalent amounts of wild-type and mutant Rad9 proteins were identified. Two independent clones of each Rad9-expressing cell line were characterized alongside wild-type and Mrad9–/– ES cells. The clone names are indicated below the panels. Rad9 was immunoprecipitated from equivalent amounts of total protein and sequentially immunoblotted for Rad1, Hus1, Rad17, and Rad9. The lower band in the Rad1 blot is a nonspecific band.

 

Expression of Wild-type Rad9 and Mutant Rad9 in Mrad9/ ES Cells—To determine the role of Rad9 phosphorylation in cellular responses to genotoxins, we created Mrad9–/– mouse ES cells stably expressing human wild-type Rad9 or Rad9 mutants that could not be phosphorylated. (The full derivation of the Mrad9–/– cells will be described elsewhere.) To ensure that epitope tags would not affect the function of Rad9, untagged Rad9 was used for the remaining studies presented here. We generated the following untagged Rad9 mutants (Fig. 1C): 1) Rad9-S272A, which converted Ser-272, a previously identified DNA damage-induced phosphorylation site, to Ala, 2) Rad9-9A, which converted all the phosphorylation sites identified in this study (including Ser-272) to Ala residues, and 3) Rad9-8A, which retained Ser-272 but converted all the remaining phosphorylation sites to Ala residues. The Mrad9–/– ES cells did not express immunologically detectable full-length mouse Rad9, whereas the corresponding wild-type ES cells did (Fig. 2). We also did not detect any amino-terminally truncated Rad9 proteins by immunoblotting with antisera that recognize the carboxyl terminus of Rad9, indicating that the targeted Mrad9 loci are not producing truncated protein (data not shown). We transfected Mrad9–/– ES cells with expression vectors encoding wild-type human Rad9 and the three mutant Rad9s (Rad9-S272A, Rad9-8A, Rad9-9A). We identified two independently derived clonal cell lines expressing each form of Rad9 at approximately equivalent levels (Fig. 2). It is worth noting that although the transfected cell lines (expressing human Rad9s) appeared to express slightly higher levels of Rad9 than did wild-type ES cells (expressing mouse Rad9), this may result from different immunoreactivities of human Rad9 compared with mouse Rad9. (The antibodies were generated to human Rad9 antigen.) Consistent with the role of phosphorylation in Rad9's extensive mobility shift, the Rad9-8A and Rad9-9A mutants exhibited dramatically increased mobility that was identical to the mobility of phosphatase-treated Rad9 (data not shown).

Rad9 Phosphorylation Is Not Required for Interaction with Hus1, Rad1, and Rad17—Rad9 tail phosphorylation could affect Rad9 function at numerous points. Because Rad9 associates with Hus1 and Rad1 as part of the 9-1-1 complex, and with Rad17, which is the putative clamp loader for the 9-1-1 complex, we first assessed whether tail phosphorylation was required for interactions with these binding partners. We immunoprecipitated wild-type human Rad9 and the phosphomutant Rad9 proteins and immunoblotted the precipitates to detect interacting endogenous mouse Rad1, Hus1, and Rad17 in the reconstituted ES cell clones. All the Rad9 mutants interacted with Hus1, Rad1, and Rad17 as well as wild-type human and wild-type endogenous mouse Rad9 did (Fig. 2), demonstrating that Rad9 phosphorylation is not required for 9-1-1 complex formation or for the 9-1-1 complex to interact with the Rad17-containing clamp loader.

Rad9 Phosphorylation Regulates Survival following Treatment with HU and UV Light—Once we established that the Rad9 mutants associated with Hus1, Rad1, and Rad17, we then asked whether the mutants affected cellular responses to genotoxins. Mhus1–/– mouse embryonic fibroblasts (MEF) are sensitive to UV light and the replication inhibitor hydroxyurea (HU) (13). We therefore examined the role of Rad9 and Rad9 phosphorylation in ES cell clonogenicity following treatment with these agents. Mrad9–/– cells, wild-type ES cells (Rad9+/+), and two independently derived clones of Mrad9–/– cells expressing human wild-type Rad9 were exposed to increasing concentrations of HU for 24 h. Like Mhus1–/– MEFs, the Mrad9–/– ES cells were exquisitely sensitive to HU (Fig. 3A). Expression of wild-type human Rad9 in the Mrad9–/– mouse ES cells restored the HU resistance to near normal levels, demonstrating that human Rad9 complemented the defect in Mrad9–/– cells (Fig. 3A).



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FIG. 3.
Rad9 and Rad9 phosphorylation are required for resistance to HU and UV. A–D, ES cells were plated, and 4 h later treated with the indicated concentrations of HU. After 24 h, the cells were washed to remove HU, and fresh media was added. The cells were cultured for 14 days, and colonies were stained and counted. The experiment has been repeated three times with identical results. The results shown are from a representative experiment. Each point shows the mean of three samples, with error bars showing S.D. Two independent clones expressing wild-type or Rad9 mutants were compared. The clone names are indicated in parentheses. E, ES cells expressing wild-type Rad9 or Rad9-S272A were treated with 50 Gy IR, 10 mM HU, or 40 J/m2 UV, and cells were lysed 1 h later. Rad9 immunoprecipitates were sequentially immunoblotted for phospho-Rad9 (P-Ser272) and total Rad9. F, ES cells were treated with 15 J/m2 the following day. The cells were stained 72 h later. One representative cell line per clone is shown in this experiment. The clone names are indicated in parentheses.

 

Following HU treatment, we then assessed clonogenic survival of the Mrad9–/– cells expressing the three Rad9 phosphorylation-deficient mutants. Rad9 Ser-272 was originally reported to be phosphorylated in response to IR in an ATM-dependent manner (37). To determine whether other genotoxins also induce Ser-272 phosphorylation, we treated Mrad9–/– ES cells expressing wild-type human Rad9 with IR, HU, or UV. Immunoblotting of the Rad9 immunoprecipitates with anti-phospho-Rad9 (P-Ser-272) antibodies revealed that all three genotoxins induced Rad9 Ser-272 phosphorylation (Fig. 3E). As a control, we also treated Mrad9–/– ES cells expressing Rad9-S272A and found that Rad9-S272A did not react with the anti-phospho-Rad9 (P-Ser-272) antibodies after genotoxin exposure. Thus, we examined whether Mrad9–/– ES cells expressing Rad9-S272A exhibited increased sensitivity to HU. Surprisingly, these cells were indistinguishable from Mrad9–/– cells expressing wild-type Rad9 (Fig. 3B). In contrast, Mrad9–/– cells expressing Rad9-8A and Rad9-9A were nearly as sensitive to HU as the parental mutant cells (Fig. 3, C and D).

We also examined the role of Rad9 phosphorylation in cells treated with UV light. Wild-type and Mrad9–/– ES cells, and Mrad9–/– ES cells expressing wild-type human Rad9 and the three Rad9 phosphorylation mutants were exposed to 15 J/m2 of UV light. Three days later the plates were stained with Coomassie Blue. As expected, cells lacking Rad9 were very sensitive to UV damage (Fig. 3F, left panel). Introduction of wild-type or Rad9-S272A restored the UV resistance to essentially wild-type ES cell levels (Fig. 3F, middle panel). However, as with the HU experiments, cells expressing Rad9-8A and Rad9-9A were as sensitive to UV as the Mrad9-null cells (Fig. 3F, right panel). Collectively, these results demonstrated that Rad9 Ser-272 phosphorylation is not required for cellular responses that affect clonogenicity following replication inhibition or UV irradiation. In contrast, one or more of the phosphorylation sites mutated in the 8A and 9A mutants is required for normal responses to these genotoxic stresses.

Rad9 Promotes Cell Survival following IR, but Rad9 Phosphorylation Plays Only a Minor Role—Mhus1–/– MEFs are sensitive to both UV light and HU, but have near wild-type sensitivity to IR (45). Because Hus1 and Rad9, along with Rad1, form a heterotrimeric complex (1416), we anticipated that Mrad9–/– ES cells would be only modestly sensitive to IR. Surprisingly, when assessed by clonogenic assays, the Mrad9–/– ES cells were far more sensitive to IR than wild-type ES cells (Fig. 4A), and expression of wild-type Rad9 in the Mrad9–/– ES cells fully restored the wild-type levels of resistance (Fig. 4A). Like the results found with HU and UV, cells expressing Rad9-S272A were no more sensitive to IR than were cells expressing wild-type Rad9, suggesting that this phosphorylation site is not required for any Rad9 function that contributes to ES cell survival (Fig. 4B). In sharp contrast, cell lines expressing Rad9-8A and Rad9-9A exhibited near wild-type sensitivity to lower doses (≤4 Gy) of IR and modestly increased sensitivity at 8 Gy IR (Fig. 4, C and D). Taken together, these results indicate that although Rad9 plays an important role in the cellular response to IR, the Rad9 phosphorylation sites identified here are not central to this function.



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FIG. 4.
Rad9 but not Rad9 phosphorylation is required for resistance to IR. A–D, ES cell lines were plated and treated with the indicated doses of IR 4 h later. The experiment has been repeated three times with identical results. The results shown are from a representative experiment. Error bars indicate S.D. Two independent clones expressing wild-type or Rad9 mutants were compared. Clone names are indicated in parentheses.

 

Rad9 Phosphorylation Is Not Required for Chromatin Binding in Response to Genotoxic Stress—We then assessed which aspect of Rad9 function required phosphorylation. In the current model for Rad9 function, the Rad17-RFC clamp loader loads the 9-1-1 complex onto DNA at sites of damage (1, 3, 29, 30). In Fig. 2, we demonstrated that Rad9 phosphorylation did not affect interaction with Hus1 and Rad1, the other members of the 9-1-1 complex, or with Rad17. Because these interactions remained intact, we then asked whether Rad9 chromatin binding, a Rad17-dependent event (12), required Rad9 phosphorylation. To streamline further biochemical characterizations for the studies that follow, we selected a representative clone from each transfected cell line. We treated wild-type cells, Mrad9–/– cells, and Mrad9–/– cells expressing Rad9 and the three Rad9 phosphorylation mutants with UV or ionizing radiation. The cells were sequentially extracted with a low salt buffer to first remove unbound Rad9 and then with a high salt buffer that dislodges chromatin-bound Rad9 (28). Endogenous Rad9 inducibly bound to chromatin after UV and IR treatment (Fig. 5). Likewise, wild-type Rad9 and Rad9-S272A also bound chromatin after genotoxic stress, showing that Ser-272 phosphorylation is not required for chromatin binding. Similarly, Rad9-8A and Rad9-9A also inducibly associated with chromatin after genotoxic stress. However, we did note a slightly higher level of chromatin-bound Rad9 in the 8A and 9A mutants even in the absence of DNA damage, suggesting that there may a slightly higher level of basal genomic stress in these cells. Taken together, these data indicate that none of the phosphorylation sites targeted in this study is required for genotoxin-induced chromatin binding.



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FIG. 5.
Rad9 phosphorylation is not required for chromatin binding in response to IR or UV. Wild-type, Mrad9–/–, and Mrad9–/– cells expressing wild-type and Rad9 mutants were left untreated or treated with either 50 Gy IR (panel A) or 40 J/m2 UV radiation (panel B). One hour later the cells were harvested and fractionated into low and high salt (chromatin-bound) fractions to separate free and chromatin-bound Rad9. Rad9 was immunoprecipitated from equivalent amounts of total protein, except for wild-type ES cells, where Rad9 was immunoprecipitated from twice the amount of protein to facilitate detection of Rad9. The immunoprecipitates were immunoblotted for Rad9. The clones used in both experiments were wild-type Rad9 (B3-24), Rad9-S272A (3-9-15), Rad9-8A (D3-8), and Rad9-9A (E2-2).

 

Rad9 Is Not Required for IR-induced Chk2 or S-phase Checkpoint Activation—Rad9, Hus1, and Rad1 function early in a checkpoint signaling cascade. To determine what checkpoint signaling pathways lie downstream of Rad9 and are dependent upon Rad9 phosphorylation, we examined genotoxin-induced Chk1 and Chk2 activation, and downstream events that require Chk1 and Chk2. Chk2, which is phosphorylated and activated by PIKKs in response to genotoxins (8, 9), undergoes a phosphorylation-dependent mobility shift detected by SDS-PAGE. Wild-type and Mrad9–/– ES cells were exposed to increasing doses of IR, and the cell lysates were immunoblotted for Chk2 (Fig. 6A). At both low and high doses we did not see any effect of Rad9 status on IR-induced Chk2 phosphorylation. Therefore, even though the Mrad9–/– cells were more sensitive to IR, they did not have a Chk2 activation defect.



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FIG. 6.
Rad9 is not required for IR-induced Chk2 or S-phase checkpoint activation. A, wild-type or Mrad9–/– ES cells were treated with the indicated doses of IR. One hour later the cells were harvested and lysed. Chk2 was immunoblotted with an anti-Chk2 antiserum. B, wild-type or Mrad9–/– cells were pretreated with nothing or 3 mM caffeine and treated with 2, 4, or 10 Gy IR or with 30 J/m2 UV light. [methyl-3H]Thymidine was added 40 min later, and the cultures were incubated for an additional 20 min. [3H]Thymidine incorporation is plotted as percent of incorporation compared with untreated cells. Values are the mean from three independent experiments. Error bars indicate S.E.

 

IR activates an S-phase checkpoint that slows DNA synthesis and blocks firing of late DNA replication origins. An ATM-activated signaling pathway regulates this IR-induced S-phase response, and mutations in ATM disrupt this checkpoint and cause a characteristic phenotype of radio-resistant DNA synthesis (RDS), in which cells synthesize DNA even in the presence of DNA damage (4648). We treated cells with increasing doses of IR or with UV light and examined their ability to synthesize DNA (Fig. 6B). As expected, the UV-induced S-phase checkpoint was disrupted in cells lacking Mrad9. In contrast, the IR-induced S-phase checkpoint did not require Rad9 and was fully reversed by caffeine, a known inhibitor of ATM (4951). Collectively, these results demonstrate that Rad9 is not involved in the RDS response.

Rad9 Is Not Required for IR-, UV-, or HU-induced p53 Accumulation—We also examined whether Rad9 was required for accumulation of the DNA damage-responsive transcription factor p53. After genotoxic stress, p53 undergoes post-translational modifications, including phosphorylation by Chk2 and ATM, which participate in p53 stabilization and activation of p53 target genes (52). Basal p53 levels were low in untreated wild-type cells and slightly higher in untreated Mrad9-null cells (Fig. 7), consistent with the results observed in Mhus1–/– MEFs (13). When treated with UV or ionizing radiation, wild-type and Mrad9–/– ES cells accumulated similar levels of p53 after 2 and 8 h (Fig. 7), demonstrating that Rad9 is not required for p53 stabilization in response to these genotoxic stresses.



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FIG. 7.
Rad9 is not required for genotoxin-induced p53 accumulation. Wild-type and Mrad9–/– cells were treated with 20 Gy IR or 20 J/m2 UV radiation and harvested 2 or 8 h later. Equal amounts of total protein were separated by SDS-PAGE and sequentially immunoblotted for p53 and JNK (as a loading control).

 

Rad9 and Its Phosphorylation Are Required for Chk1 Activation—Replication inhibition and UV irradiation trigger ATR- and Hus1-dependent Chk1 phosphorylation on Ser-345 (13, 33), which is essential for Chk1 activation (2, 53). To assess whether Rad9 and its phosphorylation also participate in Chk1 activation, wild-type, Mrad9–/– ES cells, and Mrad9–/– ES cells expressing wild-type human Rad9 or the three Rad9 phosphorylation mutants were treated with HU, UV, or increasing amounts of IR. All three stimuli triggered robust Chk1 Ser-345 phosphorylation in wild-type ES cells (Fig. 8, A and B). In contrast, UV- and IR-induced Chk1 phosphorylation was eliminated, and HU-induced phosphorylation was dramatically reduced in Mrad9–/– cells. We also noted that the mobility of the residual phospho-Chk1 in HU-treated Mrad9–/– cells was altered compared with wild-type cells (Fig. 8 and data not shown). We then asked whether Rad9 phosphorylation was required for Chk1 activation. Wild-type Rad9 restored Chk1 phosphorylation after genotoxic damage, as did the S272A mutant (Fig. 8, C and D). Consistent with the defects in survival, Rad9-8A, and Rad9-9A did not restore Chk1 phosphorylation, indicating that Rad9 tail phosphorylation is required for downstream signaling from the 9-1-1 complex to Chk1 in response to genotoxic stress.



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FIG. 8.
Rad9 and Rad9 phosphorylation are required for genotoxin-induced Chk1 activation. Wild-type or Mrad9–/– ES cells were treated with 10 mM HU or 50 J/m2 UV radiation (panel A) or with increasing amounts of IR (panel B). Mrad9–/– and Mrad9–/– cells expressing wild-type Rad9 and Rad9 mutants were treated with 10 mM HU or 50 J/m2 UV radiation (panel C) or with 50 Gy IR (panel D). One hour later the cells were harvested and lysed, and equal amounts of total protein were sequentially immunoblotted with an antibody recognizing phosphorylated Chk1 (P-Ser-345) and an antibody recognizing total levels of Chk1. The clones used were wild-type Rad9 (B3-24), Rad9-S272A (3-9-15), Rad9-8A (D3-8), and Rad9-9A (E2-2).

 

Rad9 and Its Phosphorylation Are Required for UV-induced S-phase Checkpoint Activation—Chk1 participates in the genotoxin-activated S-phase checkpoint (54). Consequently, we asked whether Rad9 phosphorylation was required for S-phase arrest following DNA damage. We UV-irradiated wild-type ES cells, Mrad9–/– ES cells, and Mrad9–/– ES cells expressing Rad9-S272A, Rad9-8A, and Rad9-9A. Forty minutes after UV irradiation [3H]thymidine was added to detect DNA synthesis, and the cells were incubated for an additional 20 min prior to harvest. The level of [3H]thymidine incorporation decreased in wild-type cells to ~60% of control, whereas the [3H]thymidine incorporation in Mrad9–/– cells was only slightly inhibited by UV irradiation (Fig. 9), indicating that the S-phase checkpoint requires Rad9. Expression of wild-type Rad9 in the Mrad9–/– ES cells reconstituted the checkpoint, as did the S272A mutant. In contrast, both Rad9-8A and Rad9-9A incorporated [3H]thymidine at levels similar to Mrad9-null cells (Fig. 9). Taken together, these results demonstrate that Rad9 phosphorylation at Ser-272 is not required for the S-phase checkpoint; however, the additional phosphorylation sites in the Rad9 tail are required for this cellular response.



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FIG. 9.
The UV irradiation-induced S-phase checkpoint requires Rad9 and Rad9 phosphorylation. Wild-type, Mrad9–/–, and reconstituted ES cells were plated in 96-well plates. The clones used were wild-type Rad9 (B3-24), S272A (3-9-15), 8A (D3-8), and 9A (E2-2). The cells were treated with 20 J/m2 and [methyl-3H]thymidine was added 40 min later, and the cells were incubated for an additional 20 min. [methyl-3H]Thymidine incorporation is plotted as percent of incorporation relative to untreated cells. Values are the mean of three independent experiments. Error bars indicate S.D.

 


    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 
Mapping Rad9 Phosphorylation Sites—Several Rad9 phosphorylation sites have been identified previously (28, 3438). Our analysis confirmed these sites (except Tyr-28 and Thr-292) and found four additional sites (Ser-341, Ser-375, Ser-380, and Ser-387). Even though Ser-272 is a DNA damage-inducible site, we identified this site in the present radiosequencing analysis, possibly because the cells were radiolabeled with large amounts of radiolabel, a procedure that activates p53 and generates DNA damage (55, 56). Alternatively, Ser-272 may also be phosphorylated by ATR. ATR is activated during unperturbed S-phase progression in Xenopus egg extracts (57), raising the possibility that this site is phosphorylated at low levels during S-phase. In contrast, based on Rad9 mobility studies, the remaining sites that we identified are phosphorylated in the absence of exogenous genotoxic stress, suggesting that they are constitutively phosphorylated; however, we cannot rule out the possibility that one or more may be inducibly phosphorylated. Nonetheless, our results clearly demonstrate that these phosphorylation sites play essential role(s) in Chk1 activation and highlight the role of the Rad9 tail in the Chk1 activation process.

Rad9 is also phosphorylated on Thr-292, a site that is preferentially phosphorylated in mitotic cells (35). Even though our radiosequencing analysis extended through the peptide containing Thr-292, we did not observe phosphorylation of this site. Because we used an exponentially growing cell population, which contains few mitotic cells, this site may have escaped our analysis and was not included in our studies of Rad9 function in intact cells.

Additionally, Rad9 is inducibly phosphorylated on Tyr-28 in a c-Abl-dependent manner following DNA damage (38), although neither of our phosphorylation site analyses detected Tyr-28 phosphorylation. Possible explanations for this discrepancy are that the radiolabeling procedure did not cause sufficient damage to induce this phosphorylation or that the techniques used here were not sufficiently sensitive to detect this phosphorylation event.

The Rad9 Tail—All Rad9 orthologs identified to date possess a carboxyl-terminal tail. The tail is the least conserved portion of Rad9, and yet all Rad9 proteins identified have tails of varying length, suggesting that it has a role in Rad9 function. Despite the fact that all Rad9s possess a tail, its role in Rad9 function is unknown. A recent study in S. pombe demonstrated that the S. pombe Rad9 tail was required for checkpoint signaling (16). However, this study did not address how loss of the tail affected interaction of Rad9 with its interacting partners or Rad9 chromatin binding. Another analysis revealed that the human Rad9 tail contains a nuclear localization sequence, which is required for Rad9 to enter the nucleus (40). This observation raises the possibility that mutation of the tail phosphorylation sites might impair nuclear localization. However, both Rad9-8A and Rad9-9A inducibly associated with chromatin following UV and IR treatment, indicating that they were present in the nucleus at the time of damage. Thus, the present studies allowed us to examine the role of the tail in checkpoint activation without deleting the tail, a manipulation that would affect other aspects of Rad9 function.

Rad9 and Rad9 Phosphorylation in Checkpoint Activation—In many respects the Mrad9–/– ES cells and Mhus1–/– MEFs have nearly identical genotoxin-triggered cellular responses. First, both Hus1- and Rad9-deficient cells were sensitive to UV and HU (45). Second, the Mhus1–/– MEFs and the Mrad9–/– ES cells were defective in Chk1 activation in response to UV, HU, and IR (13). Third, neither Hus1 nor Rad9 was required for Chk2 activation or p53 accumulation induced by these genotoxins (13). Fourth, the Hus1- and Rad9-deficient cells have defects in the S-phase checkpoint triggered by bulky DNA lesions but they do not have defects in IR-induced S-phase checkpoint (58). Collectively, these results further support the model that Hus1 and Rad9 function together in the 9-1-1 signaling complex.

In contrast, survival following treatment with IR differed significantly between the cell lines. The Hus1 null MEFs were not substantially more sensitive to IR than paired control cells (45), and we anticipated that Mrad9–/– ES cells would also be relatively insensitive to IR. Surprisingly, our studies demonstrated that Mrad9–/– ES cells were very sensitive to IR, much like S. pombe lacking Rad9, suggesting that Rad9 either has roles outside the 9–1–1 complex or that in some cell types the 9–1–1 complex may participate in cellular responses to IR. Because IR-induced Chk2 and S-phase checkpoint activation were normal in the Mrad9–/– ES cells, the increased sensitivity to IR could not be explained by loss of these checkpoint signaling events. In contrast, IR-induced Chk1 activation was disrupted in the Mrad9-null ES cells, raising the possibility that the Chk1 signaling deficiency might contribute to the survival defect. However, cells expressing Rad9–8A and Rad9–9A also exhibited the Chk1-activation defect. Yet these cells exhibited near-wild-type IR sensitivity at lower doses of IR, demonstrating that, although Rad9 is important for survival following IR, Chk1 activation does not contribute to survival at lower doses of IR (≤4 Gy). Because the cells expressing Rad9-8A and Rad9-9A were more sensitive to a higher dose of IR (8 Gy) than cells expressing wild-type Rad9, it is possible that IR-induced Chk1 may contribute to cell survival under these conditions. These results demonstrate that Rad9 plays a key role in IR-treated cells that depends only modestly on Rad9 tail phosphorylation. However, this role does not depend on Chk1 or Chk2 activation, suggesting that Rad9 must participate in other IR-activated cellular responses that are independent of the Rad9 phospho-tail.

Rad9 is inducibly phosphorylated on Ser-272 in response to IR (37) as well as HU and UV radiation (Fig. 3). In the case of IR, this phosphorylation has been implicated in the regulation of G1 to S phase progression (37). Because ES cells lack a strong DNA damage-induced G1 checkpoint, we were unable to examine this phenotype. However, we did show that Ser-272 phosphorylation was not required for survival following treatment with HU, UV, or ionizing radiation. We also showed that HU- and UV-induced Chk1 activation and UV-induced activation of the S-phase checkpoint did not require Ser-272 phosphorylation. Collectively, these results suggest that Rad9 Ser-272 phosphorylation is not required for the cellular and biochemical responses assayed in our ES cell studies; however, they do not rule out a role for this phosphorylation in other cell types or in the G1 checkpoint.

In contrast, one or more of the additional Rad9 tail phosphorylation sites was essential for Chk1 activation. Because Rad9 mutants lacking tail phosphorylation sites form a 9-1-1 complex that associates with chromatin in response to genotoxins, these results demonstrated that 9-1-1 clamp binding was not sufficient for Chk1 activation. Moreover, they also demonstrate that the mutation of the phosphorylation sites did not disrupt the overall structure of Rad9; it is possible, however, that the defects in Rad9-8A and Rad9-9A are due to conformational alterations rather than changes in phosphorylation status. The precise roles of the non-SQ phosphorylation sites in Chk1 activation are not yet understood. Although many of these sites are likely phosphorylated constitutively (based on their retarded mobility when analyzed by SDS-PAGE (Fig. 2)), we cannot exclude the possibility that some of the sites may also be inducibly phosphorylated. The phosphorylation site sequences provide few clues as to potential kinases that might phosphorylate Rad9. Six of the sites are followed by Pro residues, suggesting that Pro-directed kinases such as MAP kinase or cyclin-dependent family members might target these sites, whereas two sites are embedded in regions rich in negatively charged amino acids, raising the possibility that they may be phosphorylated by CKII. Interestingly, while this work was under review, a study was published implicating protein kinase C (PKC) {delta} in Rad9 phosphorylation (59). However, because the published study did not identify Rad9 phosphorylation sites and none of the sites reported here is a consensus PKC{delta} site, it is unclear whether PKC{delta} targets the sites identified in the present work. Studies to identify the roles of specific phosphorylation sites and the kinases that phosphorylate them are presently underway.

A Checkpoint Signaling Model—The 9-1-1 complex and ATR, as a complex with its binding partner ATRIP (18), are independently recruited to DNA lesions (12, 31, 32), where these complexes cooperatively activate Chk1. Importantly, both the 9-1-1 complex and ATR are required for Chk1 activation. The present studies provide additional insight into this requirement. Our results demonstrate that formation of the 9-1-1 complex and genotoxin-induced chromatin binding of that complex are not sufficient for Chk1 activation; the Rad9 phospho-tail plays a critical role in Rad9-dependent Chk1 activation. Because neither ATR nor Chk1 co-precipitates with Rad9,2 the role of the Rad9 phospho-tail may be to facilitate the transient assembly of a multisubunit complex that participates in Chk1 activation (Fig. 10). Although additional studies are required to determine the precise role of Rad9 phosphorylation, our investigations highlight for the first time the key role played by the Rad9 phospho-tail in genotoxin-induced checkpoint activation.



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FIG. 10.
Model for the role of the Rad9 phospho-tail in Chk1 activation. See text for discussion.

 


    FOOTNOTES
 
Note Added in Proof—The mouse ES cells bearing a mutation in Mrad9 and described in this report were originally created and characterized by Kevin M. Hopkins, Wojtek Auerbach, Xiang Yuan Wang, M. Prakash Hande, Haiying Hang, Debra J. Wolgemuth, Alexandra L. Joyner, and Howard B. Lieberman. A more detailed characterization will appear in a separate paper.

* This work was funded by National Institutes of Health Grants CA84321 (to L. M. K.), GM52493 (to H. B. L.), CA89816 (to H. B. L.), the Mayo Clinic Foundation (to L. M. K.), and the Magnus Ehrnrooth and Oskar Öflund Foundations (to P. R.-M.). The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. Back

b Present address: Turku Centre for Biotechnology Biocity, 5th floor, Tykistokatu 6B, Fin-20520 Turku, Finland. Back

h Present address: Beyond Genomics, 40 Bear Hill Rd., Waltham, MA 02451. Back

j To whom correspondence should be addressed: Division of Developmental Oncology Research, Guggenheim 13, Mayo Clinic and Foundation, 200 First St. S.W., Rochester, MN 55905. Tel.: 507-284-3124; Fax: 507-284-3906; E-mail: karnitz.larry{at}mayo.edu.

1 The abbreviations used are: PIKK, phosphatidylinositol 3-kinase-related kinase; ES, embryonic stem cells; PBS, phosphate-buffered saline; MEF, mouse embryonic fibroblasts; HU, hydroxyurea; IR, ionizing radiation. Back

2 P. Roos-Mattjus, unpublished observations. Back


    ACKNOWLEDGMENTS
 
We thank Drs. J. Chen, S. Kaufmann, and J. Sarkaria for thoughtful discussions, insights, and critical reading of the manuscript. We thank B. Madden and Dr. D. McCormick of the Mayo Protein Core Facility for performing the Edman radiosequencing analyses. We also thank Dr. J. Chen for providing the anti-Chk2 antiserum and Dr. E. Lee for sharing the anti-phospho-Rad9 (P-Ser-272) antibodies.



    REFERENCES
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 ABSTRACT
 INTRODUCTION
 EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 REFERENCES
 

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