Roos-Mattjus et al 1 Phosphorylation of Human Rad9 Is Required ...

Roos-Mattjus et al

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Phosphorylation of Human Rad9 Is Required for Genotoxin-Activated Checkpoint

Signaling

Pia Roos-Mattjus1,9, Kevin M. Hopkins7, Andrea J. Oestreich3, Benjamin T. Vroman4,

Kenneth L. Johnson6, Stephen Naylor1,2,6,10, Howard B. Lieberman7, and Larry M.

Karnitz2,3,4,5,8

Department of Biochemistry and Molecular Biology1, the Department of

Molecular Pharmacology and Experimental Therapeutics2, the Program in Tumor

Biology3, the Divisions of Developmental Oncology Research4 and Radiation Oncology5,

and the Biomedical Mass Spectrometry and Functional Proteomics Facility6

Mayo Clinic and Foundation

Rochester, MN 55905, USA

and

The Center for Radiological Research, Columbia University7

630 West 168th Street

New York, NY 10032, USA.

8Corresponding author: Division of Developmental Oncology Research, Guggenheim 13,Mayo Clinic and Foundation, 200 First Street Southwest, Rochester, MN 55905, USATelephone: 507-284-3124FAX: 507-284-3906email: [email protected]

9Present Address:Turku Centre for BiotechnologyBiocity, 5th floorTykistokatu 6BFin-20520 TurkuFinland

10Present Address:Beyond Genomics40 Bear Hill RoadWaltham, MA 02451

Running title: Rad9 tail phosphorylation affects 9-1-1 function

Copyright 2003 by The American Society for Biochemistry and Molecular Biology, Inc.

JBC Papers in Press. Published on April 21, 2003 as Manuscript M301544200 by guest on M

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Abstract

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 Ser272

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.

Keywords: checkpoint/DNA damage/phosphorylation/Rad9

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Introduction

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

(1-6). The phosphatidylinositol-3 kinase-related kinases (PIKK) 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 (8-10).

In addition to the PIKKs, Rad1, Hus1, Rad9, and Rad17 (using

Schizosaccharomyces pombe nomenclature) are evolutionarily conserved proteins

essential for Chk1 activation (11-13). Rad9, Hus1 and Rad1 form a stable heterotrimeric

complex, called the 9-1-1 complex (14-17). Biochemical, biophysical, and molecular

modeling studies predict that the 9-1-1 complex resembles PCNA (14,18-21), 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).


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Recent studies demonstrated that ATR and the 9-1-1 complex collaboratively

induce Chk1 activation. In both humans and S. 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 (11-13,33), 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 (28,34-38). 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 Ser272. 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


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which Ser272 was converted 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.


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Experimental Procedures

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-Ser345) 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-Ser272) (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 (1x107) 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 ESGRO

(Chemicon International, Temecula, CA). Cells were grown at 37°C and 5% CO2. ES

cells were transfected with Lipofectamine 2000 according to manufacturer’s instructions

(Invitrogen). Stable clones were selected in medium containing G418 (0.2 mg/ml) and


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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 full derivation of the Mrad9-/- ES cells will be described elsewhere. 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, St. Louis, MO) was prepared fresh in phosphate-buffered

saline (PBS). Cells were irradiated with a 137Cs source at a dose rate of approximately

10 Gy/min (Mark I, JLS 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.


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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 ß-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

ß–glycerophosphate) for 10 min at 4ºC. 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-tag™

(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

phospho-deficient 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.


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Purification of S-tagged Rad9

A total of 2x109 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 hours 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

[32P]orthophosphate (ICN Biomedicals, Inc.). Cells were cultured an additional two

hours, 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 ß–glycerophosphate) for 10 min at 4ºC. 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 for 2 h at 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 ß–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


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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-mm 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).


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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 l-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% TFA, 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-mm 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-propanol/water

(80:10:10) containing 0.2% formic acid. A linear gradient from 0-50% B over 30 minutes

was performed. The liquid chromatography pumping system (Michrom UMA, Michrom


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BioResources Inc., Auburn, CA) was operated at 50 ml/min and split to a column flow of

0.2 mL/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 phospho-peptides if the mass of the 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

database 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 minutes at 37°C following treatment. [Methyl-

3H]thymidine (TRA120, Amersham Pharmacia) was then added at 2 µCi/well and the

cells were incubated for an additional 20 minutes. 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.


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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 to 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.


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Results

Mapping of Rad9 phosphorylation sites

Rad9 is extensively and constitutively phosphorylated even in cells that have not

been exposed to exogenous genotoxins (34-36). Rad9 is further inducibly

phosphorylated in response to DNA damage (34-38). 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 over-expressed 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. 1A and

1B, Ser272, Ser277, Ser 328, Ser375, Ser 380 and Ser387). Using mass spectrometry,

we identified five phosphorylated sites (Fig. 1A and 1B). Two of these sites (Ser277 and

Ser328) were also identified by radiosequencing. The three additional sites (Ser336,

Ser341 and Thr355) 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


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observed by SDS-PAGE (data not shown and Fig. 2), indicating that these studies

identified all the phosphorylation sites responsible for the mobility shift.

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 insure 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 Ser272, a previously identified DNA damage-induced phosphorylation

site, to Ala; (2) Rad9-9A, which converted all the phosphorylation sites identified in this

study (including Ser272) to Ala residues; and (3) Rad9-8A, which retained Ser272 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 to mouse Rad9.

(The antibodies were generated to human Rad9 antigen.) Consistent with the role of


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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 phospho-mutant 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 hr. Like Mhus1-/- MEFs, the Mrad9-/- ES cells were exquisitely sensitive to HU (Fig.


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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).

We then assessed clonogenic survival of the Mrad9-/- cells expressing the three

Rad9 phosphorylation-deficient mutants treated with HU. Rad9 Ser272 was originally

reported to be phosphorylated in response to IR in an ATM-dependent manner (37). To

determine whether other genotoxins also induce Ser272 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-Ser272) antibodies revealed

that all three genotoxins induced Rad9 Ser272 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-Ser272) 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 (Figs. 3C and 3D).

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


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were as sensitive to UV as the Mrad9 null cells (Fig. 3F, right panel). Collectively, these

results demonstrated that Rad9 S272 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 (14-16), 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

(Figs. 4C and 4D). 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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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 Ser272 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.

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


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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.

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 (46-48). 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 (49-51). 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

posttranslational 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


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(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 hr (Fig. 7), demonstrating that Rad9 is not required for p53 stabilization

in response to these genotoxic stresses.

Rad9 and its phosphorylation are required for Chk1 activation

Replication inhibition and UV irradiation trigger ATR- and Hus1-dependent Chk1

phosphorylation on Ser345 (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 Ser345 phosphorylation in wild-type ES cells

(Fig. 8A and 8B). 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 to 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. 8C and 8D).

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.

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


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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 minutes prior to harvest. The level of 3H-thymidine

incorporation decreased in wild-type cells to approximately 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 Ser272 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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Discussion

Mapping Rad9 phosphorylation sites

Several Rad9 phosphorylation sites have been identified previously (28,34-38).

Our analysis confirmed these sites (except Tyr28 and Thr292) and found four additional

sites (Ser341, Ser375, Ser380 and Ser387). Even though Ser272 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, Ser272 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 Thr292, a site that is preferentially

phosphorylated in mitotic cells (35). Even though our radiosequencing analysis extended

through the peptide containing Thr292, 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.


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Additionally, Rad9 is inducibly phosphorylated on Tyr28 in a c-Abl-dependent

manner following DNA damage (38), although neither of our phosphorylation site

analyses detected Tyr28 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 (18).

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


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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


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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 Ser272 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 Ser272 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

Ser272 phosphorylation. Collectively, these results suggest that Rad9 Ser272

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-


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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) d 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 d site, it is unclear whether PKC d 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 (60),

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 (P.R.-M., unpublished observations), 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-


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induced checkpoint activation.


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Acknowledgements

We wish to 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-Ser272) antibodies. This

work was funded by NIH grants CA84321 (L.M.K.), GM52493 (H.B.L.), CA89816

(H.B.L.), the Mayo Clinic Foundation (L.M.K.) and the Magnus Ehrnrooth and Oskar

Öflund foundations (P.R.-M.).


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1716


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Figure legends

Figure 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 the methods used to

identify the site. (C) Three different mutants were generated: Ser272 mutated to alanine

(S272A), all mapped phosphorylation sites except for Ser272 mutated to alanine (8A)

and all mapped phosphorylation sites mutated to alanine (9A).

Figure 2

Characterization of Rad9-expressing mouse ES cell lines. Mrad9-/- 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.

Figure 3

Rad9 and Rad9 phosphorylation are required for resistance to HU and UV. (A-D) ES

cells were plated, and four hours 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


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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 standard deviation. 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.

Figure 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 four hours later. The experiment

has been repeated three times with identical results. The results shown are from a

representative experiment. Error bars indicate standard deviation. Two independent

clones expressing wild-type or Rad9 mutants were compared. Clone names are

indicated in parentheses.

Figure 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


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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).

Figure 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 to untreated cells. Values

are the mean from three independent experiments. Error bars indicate standard error.

Figure 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 hours

later. Equal amounts of total protein were separated by SDS-PAGE and sequentially

immunoblotted for p53 and JNK (as a loading control).

Figure 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


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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-Ser345) 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).

Figure 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 standard deviation.

Figure 10

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


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Figure 3A B

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control 15 J/m2

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Mrad9-/- wt ES

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- HUUV - HUUVwt ES Mrad9-/-

P-Chk1 (S345)

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A

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Figure 8


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3 H-th

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UV 20 J/m2


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L. Johnson, Steve Naylor, Howard B. Lieberman and Larry M. KarnitzPia Roos-Mattjus, Kevin M. Hopkins, Andrea J. Oestreich, Benjamin T. Vroman, Kenneth

signalingPhosphorylation of human Rad9 is required for genotoxin-activated checkpoint

published online April 21, 2003J. Biol. Chem.

10.1074/jbc.M301544200Access the most updated version of this article at doi:

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