Protein Phosphatases Regulate DNA-dependent Protein Kinase Activity*

Pauline DouglasDagger, Greg B. G. Moorhead, Ruiqiong Ye, and Susan P. Lees-Miller§

From the Department of Biological Sciences, University of Calgary, Calgary, Alberta T2N 1N4, Canada

Received for publication, December 26, 2000, and in revised form, March 12, 2001


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

DNA-dependent protein kinase (DNA-PK) is a complex of DNA-PK catalytic subunit (DNA-PKcs) and the DNA end-binding Ku70/Ku80 heterodimer. DNA-PK is required for DNA double strand break repair by the process of nonhomologous end joining. Nonhomologous end joining is a major mechanism for the repair of DNA double strand breaks in mammalian cells. As such, DNA-PK plays essential roles in the cellular response to ionizing radiation and in V(D)J recombination. In vitro, DNA-PK undergoes phosphorylation of all three protein subunits (DNA-PK catalytic subunit, Ku70 and Ku80) and phosphorylation correlates with inactivation of the serine/threonine protein kinase activity of DNA-PK. Here we show that phosphorylation-induced loss of the protein kinase activity of DNA-PK is restored by the addition of the purified catalytic subunit of either protein phosphatase 1 or protein phosphatase 2A (PP2A) and that this reactivation is blocked by the potent protein phosphatase inhibitor, microcystin. We also show that treating human lymphoblastoid cells with either okadaic acid or fostriecin, at PP2A-selective concentrations, causes a 50-60% decrease in DNA-PK protein kinase activity, although the protein phosphatase 1 activity in these cells was unaffected. In vivo phosphorylation of DNA-PKcs, Ku70, and Ku80 was observed when cells were labeled with [32P]inorganic phosphate in the presence of the protein phosphatase inhibitor, okadaic acid. Together, our data suggest that reversible protein phosphorylation is an important mechanism for the regulation of DNA-PK protein kinase activity and that the protein phosphatase responsible for reactivation in vivo is a PP2A-like enzyme.


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The reversible phosphorylation of proteins, catalyzed by protein kinases and protein phosphatases, is a major mechanism for the regulation of many eukaryotic cellular processes, including metabolism, muscle contraction, pre-mRNA splicing, and cell cycle control (1). The eukaryotic protein phosphatases comprise several families of enzymes that catalyze the dephosphorylation of intracellular phosphoproteins. The protein phosphatases responsible for dephosphorylation of serine and threonine residues in the cytoplasmic and nuclear compartments of eukaryotic cells are encoded by the PPP and PPM gene families, which are defined by distinct amino acid sequences and tertiary structures (2). The type 1, 2A, and 2B protein phosphatases (PP1, PP2A, and PP2B, respectively)1 belong to the PPP family. This family also includes a growing list of novel protein phosphatases (e.g. protein phosphatase 4, 5, 6, and 7) (2, 3).

Inhibitors of protein (serine/threonine) phosphatases include endogenous proteins that regulate protein phosphatases in eukaryotic cells, such as inhibitor-1, DARPP 32, and inhibitor-2, which specifically inhibit PP1 (4, 5). Several toxins, drugs, and tumor promotors, including okadaic acid, microcystins, tautomycin, nodularin, cantharidin, endothall, calyculin A, and fostriecin, are also highly specific inhibitors of members belonging to the PPP family of serine/threonine protein phosphatases. These toxins bind to the same site on the enzymes but with different relative affinities (6, 7) and have been fundamental to understanding the role of protein phosphorylation in various physiological processes and deciphering which type of protein phosphatase is responsible for a given cellular event (6).

The differential sensitivities to these inhibitors have provided methods to identify and quantitate the levels of PP1 and PP2A in cell and tissue extracts (8). Several of these inhibitors, including okadaic acid and fostriecin, are membrane-permeable and potently inhibit phosphatase activity in intact cells. Due to their differential affinities to PP1 and PP2A and their distinct permeation properties, these two inhibitors can inhibit PP1 and PP2A in a highly selective manner (9). We have used these inhibitors at PP2A-selective concentrations to investigate the regulation of DNA-dependent protein kinase (DNA-PK) by protein phosphorylation in vitro and in vivo.

DNA-PK is composed of a catalytic subunit (DNA-PKcs) and a heterodimeric DNA end-binding protein, Ku70/Ku80. DNA-PKcs and Ku are required for the repair of ionizing radiation induced DNA damage via the process of nonhomologous end joining (reviewed in Ref. 10). Although the exact role of DNA-PKcs and Ku in nonhomologous end joining has yet to be determined, in vitro, DNA-PK acts as a serine/threonine protein kinase and phosphorylates a variety of protein substrates including p53 and the 32-kDa subunit of DNA replication protein A (reviewed in Refs. 10 and 11). Several in vitro DNA-PK phosphorylation sites (e.g. serine 15 of human p53 and serines and threonines in the amino-terminal 30 amino acids of the replication protein A 32-kDa subunit) are phosphorylated in vivo in response to DNA damage (12-14); however, it is not clear whether DNA-PK plays a direct role in these processes in vivo. DNA-PK also undergoes phosphorylation of all three of its protein components in vitro, and phosphorylation of the DNA-PK complex correlates with loss of protein kinase activity and disruption of DNA-PKcs from the Ku-DNA complex (15). The DNA-PK phosphorylation sites on DNA-PKcs are presently unknown; however, Ku70 is phosphorylated by DNA-PK in vitro predominantly at serine 6, while Ku80 is phosphorylated on at least three carboxyl-terminal sites, including serines 577 and 580 and threonine 715 (16). Although the physiological substrates of DNA-PK are unknown, a conserved aspartic acid residue in the kinase domain of DNA-PKcs is required to rescue the radiosensitive phenotype associated with deficiency of DNA-PKcs in rodent cells (17), strongly suggesting that DNA-PK acts as a protein kinase in vivo. Ku is phosphorylated in vivo (18), although the sites of in vivo phosphorylation have yet to be identified.

Here we have examined the effects of protein dephosphorylation on the activity of in vitro DNA-PK-mediated phosphorylation of DNA-PKcs and Ku. We show that dephosphorylation of self-phosphorylated DNA-PK results in reactivation of DNA-PK protein kinase activity. Moreover, we show that DNA-PK protein kinase activity is significantly reduced in cells that have been treated with the protein phosphatase inhibitors okadaic acid and fostriecin and that inhibition of protein phosphatase activity by okadaic acid significantly increases the in vivo phosphorylation state of DNA-PKcs, Ku70, and Ku80. Together, our data show that DNA-PK protein kinase activity can be regulated by reversible protein phosphorylation in vitro and that DNA-PK is a target of protein kinases and protein phosphatases in vivo.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Reagents-- Bovine serum albumin, phenylmethylsulfonyl fluoride, Tris base, EGTA, leupeptin, pepstatin, and wortmannin were from Sigma. A Mono Q (HR5/5) column and 32P-labeled inorganic phosphate were from Amersham Pharmacia Biotech. [gamma -32P]ATP was from PerkinElmer Life Sciences. Okadaic acid (OA) and microcystin-LR were from Calbiochem. Dithiothreitol was from BDH. Fostriecin was a kind gift from Dr. Michel Roberge (University of British Columbia).

Cell Culture and Inhibitor Treatment-- Lymphoblastoid cells (BT) and MO59J cells were maintained in RPMI or Dulbecco's modified Eagle's medium, respectively (Life Technologies, Inc.) supplemented with 10% fetal calf serum in an atmosphere of 5% CO2. Log phase cells (1 × 106 cells/10-ml dish) were incubated in media containing OA or fostriecin. OA was prepared by dissolving in dimethyl sulfoxide, and fostriecin was dissolved in phosphate-buffered saline containing 0.1 mM ascorbic acid. Control incubations included that same amount of vehicle alone.

Preparation of S10 and P10 Extracts-- Cells were harvested either by centrifugation (BT) or trypsinization (MO59J), washed twice in phosphate-buffered saline, and lysed by a single freeze thaw cycle as described previously (19). Cytoplasmic (S10) and nuclear (P10) extracts were prepared as described (20). Extracts were snap frozen in liquid nitrogen and stored at -80 °C. Protein concentrations were determined using a dye binding assay (Bio-Rad) with bovine serum albumin as a standard.

Preparation of Recombinant PP1 and Purification of PP2A-- Human PP1gamma 1 cDNA was polymerase chain reaction-amplified from a human (brain) cDNA library (CLONTECH) incorporating an NdeI site at the initiator codon (ATG) and a HindIII site immediately following the stop codon. The polymerase chain reaction product was cloned into pCRTOPO (Invitrogen), and the sequence was verified by DNA sequencing. The modified cDNA was then cloned into the NdeI/HindIII sites of the pCW vector (a kind gift from Dr. F. W. Dalhqvist, University of Oregon) and introduced into Escherichia coli DH5alpha by standard techniques.

Cells were grown in LB medium plus 1 mM MnCl2, and, after reaching an A600 of 0.3, cells were induced with 0.1 mM IPTG for 16 h and then pelleted by centrifugation at 4000 × g for 30 min. Cells from 0.75 liters of suspension were resuspended in 15 ml of 50 mM Hepes-KOH, pH 7.5, 100 mM KCl, 1 mM EDTA, 2 mM MnCl2, 5% glycerol, 0.1% beta -mercaptoethanol, 0.1 mM phenylmethylsulfonyl fluoride and snap frozen in liquid nitrogen and then stored at -80 °C. Frozen cells were thawed and put through a French press cell (SIM Aminco) one time at 1100 p.s.i., and the extract was clarified by centrifugation at 35,000 × g for 30 min. The clarified extract was filtered through a 0.45-µm filter and loaded onto a 20-ml column of SP-Sepharose (Amersham Pharmacia Biotech) at 3 ml/min. The column was equilibrated in buffer A (50 mM Tris-HCl, pH 7.5, 0.1 mM EGTA, 5% glycerol, 0.1% beta -mercaptoethanol, and 1 mM MnCl2). PP1gamma 1 was eluted with a gradient from 0-500 mM NaCl over 200 ml with 5-ml fractions. PP1gamma 1 was further purified on a Mono-Q column equilibrated in buffer A. The column was developed at 1 ml/min from 0-500 mM NaCl over 40 ml, and 1-ml fractions were collected. Peak fractions were dialyzed into buffer A containing 50% glycerol and 50 mM NaCl and stored at -20 °C. The catalytic subunit of protein phosphatase 2A was purified from bovine heart as described (21).

Preparation of 32P-Labeled Glycogen Phosphorylase a and Protein Phosphatase Assays-- 32P-Labeled glycogen phosphorylase a containing 1.0 mol of phosphate/mol of subunit was prepared using phosphorylase b (22) and phosphorylase kinase (23) purified from rabbit skeletal muscle by the methods described. Details of the assay can be found in Ref. 4. Briefly, assays were performed at 30 °C in a total volume of 30 µl that included 10 µl of protein phosphatase diluted as required into buffer B (50 mM Tris-HCl, pH 7.5 (20 °C), 0.1 mM EGTA, 0.1% (v/v) beta -mercaptoethanol, 1 mg/ml bovine serum albumin) plus 10 µl of buffer C (50 mM Tris-HCl, pH 7.5 (20 °C), 0.1 mM EGTA, and 0.03% (v/v) Brij-35 plus inhibitors/activators as required). The assays were initiated by the addition of 10 µl of 30 µM 32P-labeled glycogen phosphorylase a. One unit is defined as the amount of enzyme that catalyzed the release of 1 µmol of phosphate in 1 min. In all cases, dephosphorylation of substrates was kept less than 20% to ensure the linearity of the assay. Rabbit skeletal muscle inhibitor-2 was purified according to Cohen et al. (24) and used at a final concentration of 200 nM in assays. All assays that included protein phosphatase inhibitors were incubated at 30 °C for 15 min before initiating the assay with substrate. Inhibitor-2 was diluted in assay buffer C immediately before use.

Protein Purification and DNA-PK Activity Assays-- The DNA-PKcs and Ku subunits of DNA-PK were purified from human placenta as described previously (25). Kinase assays using purified proteins or cell extracts from MO59J (3 µg) or lymphoblastoid cells (3 µg) were as described (19) except that the synthetic peptide substrate used was PESQEAFADLWKK (26). The peptide, PESEQAFADLWKK, which is not phosphorylated by DNA-PK, was used as a control (26). Reactions contained 25 mM Hepes-NaOH, pH 7.5, 0.25 mM synthetic peptide, 100 mM KCl, 10 mM MgCl2, 1 mM dithiothreitol, 0.2 mM EGTA, 0.1 mM EDTA plus 10 µg/ml sonicated calf thymus DNA, and 0.25 mM ATP containing stabilized [gamma -32P]ATP (specific activity 500-1000 dpm/pmol) and were started by the addition of purified DNA-PK proteins (concentration as indicated). Reactions were at 30 °C for 5 min, and DNA-PK protein kinase activity was calculated as nmol of phosphate incorporated into the peptide substrate/min/mg of protein. Synthetic peptides were synthesized and purified by high pressure liquid chromatography by the Alberta Peptide Institute (Edmonton, Alberta, Canada).

Autophosphorylation of DNA-PK-- Purified DNA-PKcs and Ku proteins were preincubated at 30 °C as described for activity assays, except that synthetic peptide was not present. Radiolabeled ATP was present where indicated at 0.25 mM. After 0-10 min, aliquots were removed and analyzed by SDS-PAGE followed by autoradiography. In order to reassay samples for remaining protein kinase activity, aliquots corresponding to 5-10% (v/v) or the percentage indicated of the preincubation reaction were removed and reassayed under standard kinase assay conditions with a full complement of synthetic peptide, DNA, and radiolabeled ATP as described above. For "add-back" experiments, after the indicated times, either the free catalytic subunit of PP1 (50 milliunits/ml final concentration), the free catalytic subunit of PP2A (50 milliunits/ml final concentration) or protein extracts from lymphoblastoid or MO59J cells (treated with Me2SO, OA, or fostriecin) were added to aliquots from preincubation reactions containing phosphorylated DNA-PK. At timed intervals, aliquots corresponding to 5-10% (v/v) of this reaction were removed and reassayed under standard kinase assay conditions with a full complement of synthetic peptide, DNA, and radiolabeled ATP. In identical experiments, aliquots were removed and analyzed by SDS-PAGE, followed by autoradiography.

The stoichiometry of phosphorylation of in vitro phosphorylated DNA-PKcs was estimated by excising the phosphorylated bands from the dried Coomassie-stained gels and measuring 32P incorporation by Cerenkov counting in a scintillation counter. The specific activity of the radioactive ATP was calculated in cpm/pmol, and the amount of DNA-PKcs and Ku was estimated from the predicted molecular masses (~470, 80, and 70 kDa, respectively) and the protein concentration as determined by the Bio-Rad protein assay.

In Vivo Labeling and Immunoprecipitation of DNA-PKcs and Ku-- BT cells were grown to approximately midlog phase, and then cells were transferred to 15-ml conical tubes and pelleted by centrifugation at 1500 × g for 5 min. Cells were then rinsed twice with phosphate-free RPMI medium. For each experiment, ~2 × 106 cells were resuspended in 1 ml of phosphate-free medium containing 10% fetal calf serum and incubated with okadaic acid (1 µM) or an equivalent volume of Me2SO for 1 h in a humidified CO2 incubator at 37 °C under 5% CO2. Cells were allowed to incorporate 32P-labeled inorganic phosphate (0.75 millicuries/ml) for a further 1 h, and then the label was removed and cells were washed twice in ice-cold phosphate-buffered saline and lysed in immunoprecipitation buffer containing 20 mM Tris-HCl, pH 7.4, 250 mM NaCl, 1 mM EDTA, 1% (v/v) Nonidet P-40, 2 µg/ml aprotinin, 2 µg/ml leupeptin, 1 mM phenylmethylsulfonyl fluoride, 1 mM Na3VO4, and 10 mM NaF. DNA-PKcs and Ku were immunoprecipitated as described previously (15) and transferred to nitrocellulose membrane. The membrane was exposed to x-ray film overnight at -80 °C and then probed with an antibody to either Ku70 or DNA-PKcs as described in Fig. 6. Approximately 2 × 107 cpm of lysate was used for each immunoprecipitation reaction.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

We have previously shown that incubation of purified DNA-PKcs and Ku heterodimer with DNA and Mg-ATP results in phosphorylation of all three subunits of the DNA-PK holoenzyme and time-dependent loss of protein kinase activity (15). Loss of DNA-PK protein kinase activity did not occur when the purified protein was incubated with Mg-ATP in the absence of DNA or with DNA, magnesium, and the nonhydrolyzable ATP analogue AMP-PNP (15), suggesting that loss of DNA-PK protein kinase activity correlates with phosphorylation of one or more of the DNA-PK subunits.

An important prediction of these results is that treatment with a protein phosphatase would reverse the phosphorylation-induced loss of DNA-PK protein kinase activity. In order to test this hypothesis, purified DNA-PKcs and Ku were first incubated with DNA and Mg-ATP in order to produce phosphorylated, inactive DNA-PK. Following this 10-min incubation period, the purified catalytic subunit of either PP1 or PP2A was added to the reactions, and DNA-PK protein kinase activity was assayed. The addition of the catalytic subunit of either PP1 or PP2A resulted in a complete (PP1) or partial (PP2A) restoration of DNA-PK protein kinase activity (Fig. 1, A and B, open symbols). In contrast, when the catalytic subunit of PP1 or PP2A was preincubated with microcystin, a potent inhibitor of both protein phosphatases, no significant increase in DNA-PK protein kinase activity was observed (Fig. 1, A and B, closed symbols). Moreover, both PP1 and PP2A catalytic subunits were capable of removing phosphate from DNA-PKcs, Ku70, or Ku80 (Fig. 2, A and B). These results strongly support our hypothesis that the loss of DNA-PK protein kinase activity in vitro is due to reversible protein phosphorylation.


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Fig. 1.   Incubation of protein phosphatase 1 and 2A catalytic subunits with phosphorylated inactivated DNA-PK complex restores DNA-PK protein kinase activity. A, DNA-PK complex consisting of DNA-PKcs (0.013 µg/µl) and Ku (0.004 µg/µl) (molar ratio 1:1) was incubated in a final volume of 33 µl, under standard assay conditions as described previously (15). At 0, 2, 5, and 10 min, 2-µl aliquots (equivalent to 0.1 µg of total protein) were removed and assayed under standard assay conditions as described previously (15). At 10 min, indicated by the arrow, either recombinant PP1 (final concentration of 50 milliunits/ml (open squares)) or recombinant PP1 (final concentration of 50 milliunits/ml) that was preincubated with microcystin-LR (1 µM final concentration) (closed squares) was added to the incubation mixture. After these additions, at timed intervals 2-µl aliquots were removed and assayed under standard DNA-PK assay conditions as described previously (15). B, DNA-PK was incubated exactly as described for A; however, the catalytic subunit of PP2A (purified from bovine heart) (50 milliunits/ml) was added in place of recombinant PP1.


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Fig. 2.   The catalytic subunits of PP1 and PP2A are capable of dephosphorylating in vitro autophosphorylated DNA-PK. DNA-PKcs (0.04 µg/µl) and Ku (0.013 µg/µl) (1 µg of DNA-PK total protein) were preincubated in 50 mM Hepes-NaOH, pH 7.5, 50 mM KCl, 10 mM magnesium chloride, 1 mM dithiothreitol, 0.2 mM EDTA, 10 µg/ml calf thymus DNA, and 0.25 mM [gamma -32P]ATP in a total volume of 20 µl for 10 min. Following the first incubation period, various concentrations of recombinant PP1 catalytic subunit (A) or PP2A catalytic subunit (purified from bovine heart) (B) were added to the reaction at 15-s intervals (as shown), and incubations were continued for a further 10 min. Following the second incubation period, microcystin-LR (1 µM final) was added at 15-s intervals to stop the dephosphorylation reactions, and samples were boiled in SDS sample buffer. Samples were then analyzed by SDS-PAGE on 10% acrylamide gels followed by autoradiography. Shown are separate exposures for DNA-PKcs (5 h at -80 °C with intensifying screens) and Ku (overnight at -80 °C with intensifying screens).

The activity of most known protein phosphatases is regulated by association with regulatory subunits that can target the protein phosphatase to certain subcellular locations or directly affect its activity by allosteric or other effects. It was therefore important to determine if the protein kinase activity of DNA-PK was affected by phosphorylation in vivo. Specific protein phosphatases can be inhibited by treatment of cells with particular cell-permeable protein phosphatase inhibitors. For example, treatment of human cells with okadaic acid at 1 µM results in loss of PP2A activity but not PP1 activity (27). Human lymphoblastoid (BT) cells were incubated with either Me2SO (control), okadaic acid, or fostriecin at PP2A-selective concentrations and were assayed for protein phosphatase activity using radiolabeled phosphorylase a as a substrate. Extracts were assayed either with no additions (Fig. 3A, solid bars), with the addition of the PP1 inhibitor, inhibitor-2, at 200 nM final (Fig. 3A, open bars) or inhibitor-2 (200 nM final concentration) plus 5 nM okadaic acid (Fig. 3A, hatched bars). In this experiment, the activity of both PP1 and PP2A is shown in assays with no additions (solid bars), whereas in assays that contained inhibitor-2, only PP2A-like protein phosphatases would be active (open bars), and in assays that contained okadaic acid (at a concentration of 5 nM final) and inhibitor-2 (hatched bars), neither of the protein phosphatases would be active. These data show that of the total phosphorylase phosphatase activity in BT cells, ~40% is PP1 protein phosphatase activity and 60% is PP2A-like protein phosphatase activity (Fig. 3A, DMSO, open bars compared with closed bars). Treatment of cells with either okadaic acid or fostriecin, which, at the specified concentrations used, is predicted to inhibit only PP2A-like protein phosphatase activity, resulted in an ~50% loss of phosphorylase phosphatase activity (Fig. 3A, OA or fostriecin, solid bar). The further addition of inhibitor-2 (200 nM) to these extracts abolished the majority of the remaining protein phosphatase activity, and this is unaltered by the further addition of 5 nM OA to these extracts. This indicates that treatment of cells with either OA or fostriecin abolishes PP2A-like activity, leaving PP1 unaffected. Having established this, the same extracts were used to assay for DNA-PK protein kinase activity, and the DNA-PK protein kinase activity was reduced by ~50% in the extracts from cells that had been treated with either okadaic acid or fostriecin (Fig. 3B). No change in the amount of DNA-PKcs protein was detected after treatment of cells with either okadaic acid or fostriecin (Fig. 3C). These data support a model in which DNA-PK protein kinase activity is regulated by reversible protein phosphorylation in vivo and indicate that, in vivo, protein phosphatases are required for maintaining DNA-PK in a highly active state. Consequently, this model would predict that DNA-PK is phosphorylated in vivo, and as the phosphorylation state of DNA-PK increases, its activity decreases.


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Fig. 3.   Effects of okadaic acid and fostriecin on protein phosphatase and DNA-PK activity in lymphoblastoid cells. A, lymphoblastoid cells (BT) were treated with control (Me2SO; DMSO), okadaic acid (1 µM final concentration), or fostriecin (100 µM final) for 2 h. Cells were harvested, and S10 and P10 fractions were prepared as described under "Experimental Procedures." For protein phosphatase assays, equal volumes of S10 and P10 fractions added back together, to give 15 µg of total protein, and the combined lysates were diluted 50-fold in protein phosphatase assay buffer B (see "Experimental Procedures") and assayed for protein phosphatase activity with 32P-labeled phosphorylase a as a substrate. Assays were carried out with no additions (solid bars), with the addition of rabbit skeletal muscle inhibitor-2 at a 200 nM final concentration in assay (open bars), or with the addition of inhibitor-2 at 200 nM (final concentration) plus OA (5 nM final concentration) (hatched bars). Protein phosphatase activity was calculated as a percentage of total protein phosphatase activity from control (DMSO) cell extracts. B, DNA-PK protein kinase activity from the P10 extracts of cell lysates from A was assayed under standard kinase assay conditions in the presence of synthetic peptide (PESQEAFADLWKK). 2 µl of extract (equivalent to 3 µg of total protein) was preincubated in 50 mM Hepes-NaOH, pH 7.5, 50 mM KCl, 10 mM magnesium chloride, 1 mM dithiothreitol, 0.2 mM EDTA, 10 µg/ml calf thymus DNA, and synthetic peptide in a total volume of 18 µl. Reactions were started at 15-s intervals with the addition of 2 µl of ATP containing [gamma -32P]ATP (final concentration in the assay of 0.25 mM ATP, 1 µCi of [gamma -32P]ATP). DNA-PK protein kinase activity was calculated as a percentage of activity from control (DMSO) cell extracts. Shown are the mean ± S.E. values of three separate experiments. Where not shown, S.E. was <3.0%. C, 15 µg of total protein from the P10 extracts from the experiment shown in Fig. 3B were analyzed by Western blot using an antibody to DNA-PKcs (DPK1). Lane 1, Me2SO (DMSO)-treated cells; lane 2, okadaic acid-treated cells; lane 3, fostriecin-treated cells. This control experiment showed that the observed reduction in DNA-PK activity was not due to a reduction in DNA-PK protein levels.

Our data therefore suggest that a protein phosphatase activity acts on DNA-PK in vivo. It was therefore important to show that in cell extracts there exists a protein phosphatase activity capable of removing phosphate from DNA-PK. Purified DNA-PK was phosphorylated and inactivated by incubation with DNA and radioactively labeled Mg-ATP. The phosphorylation reactions were stopped by the addition of wortmannin to 10 µM, and in vitro phosphorylated DNA-PK protein was incubated with extracts from cells that were either untreated, treated with okadaic acid, or treated with fostriecin. The results show that extracts from untreated cells contained an activity capable of removing phosphate from in vitro phosphorylated DNA-PKcs and that the presence of inhibitor-2 (a PP1 inhibitor) had no inhibitory effect on the dephosphorylation reaction (Fig. 4, compare lane 1 with lanes 2 and 3). Significantly, when cells were treated with okadaic acid or fostriecin, at PP2A-selective concentrations, no significant loss of DNA-PK phosphorylation was observed (Fig. 4, lanes 4-7). Again, treatment of extracts with the PP1 inhibitor, inhibitor-2, had no effect on dephosphorylation of DNA-PKcs, strongly suggesting that the phosphatase in these extracts responsible for dephosphorylating in vitro autophosphorylated DNA-PKcs is a PP2A-like enzyme. Identical results were observed for Ku70 and Ku80 (data not shown). Although the data shown in Fig. 1 suggests that PP1 is more efficient than PP2A at dephosphorylating and activating DNA-PK in vitro, it is important to note that these experiments were carried out using the free catalytic subunits of either PP1 or PP2A. These enzymes therefore lack their regulatory subunit(s) and consequently would be expected to have different properties from the naturally occurring enzymes.


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Fig. 4.   Add-back of extracts treated with protein phosphatase inhibitors to phosphorylated-inactivated DNA-PKcs. Purified DNA-PK (equivalent to 1 µg of protein) was incubated in the presence of DNA, MgCl2, and [gamma -32P]ATP as described in the legend to Fig. 1. After 10 min, cell lysates (5 µg), prepared exactly as in Fig. 3A, were added to reactions. Where indicated, cell lysates were pretreated with 200 nM inhibitor-2 (+) or with an equal volume of buffer (50 mM Hepes-NaOH, pH 7.5) (-). Incubations were continued for a further 10 min, and then reactions were stopped by the addition of microcystin-LR (1 µM final concentration), and samples were analyzed by SDS-PAGE on 10% acrylamide gels followed by autoradiography. Lane 1, no additions (N/A); lanes 2 and 3, cell extracts treated with Me2SO; lanes 4 and 5, cell extracts treated with okadaic acid (1 µM final concentration); lanes 6 and 7, cell extracts treated with fostriecin (100 µM final concentration).

In order to determine whether cell extracts that had been treated with okadaic acid or fostriecin were incapable of reactivating phosphorylated DNA-PK, purified DNA-PK was phosphorylated and inactivated in vitro and then incubated with extracts from cells that were either untreated or treated with PP2A-selective concentrations of okadaic acid or fostriecin. Since BT cells contain abundant DNA-PK protein kinase activity, which would complicate interpretation of results, a human cell line that lacks DNA-PKcs protein (MO59J) was used for these experiments (28). Protein extracts from untreated MO59J cells contained an activity that was capable of reactivating DNA-PK, whereas treatment of MO59J cells with either okadaic acid or fostriecin significantly decreased the ability of these extracts to reactivate DNA-PK (Fig. 5). These data again support our hypothesis that human cells contain a PP2A-like enzyme that can both remove phosphate groups from in vitro phosphorylated DNA-PK and reverse phosphorylation-induced loss of protein kinase activity.


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Fig. 5.   Reactivation of phosphorylated inactivated DNA-PK, using extracts from MO59J cells treated with protein phosphatase inhibitors. MO59J cells were treated with control (Me2SO; DMSO), okadaic acid (1 µM final concentration), or fostriecin (100 µM final concentration) for 2 h. Cells were harvested, and S10 and P10 extracts were prepared exactly as described in the legend to Fig. 3A. Equal volumes of lysates (equivalent to 7 µg of protein) were combined and added back to purified DNA-PK complex consisting of DNA-PKcs (0.013 µg/µl) and Ku (0.004 µg/µl) (molar ratio 1:1) that had been preincubated for 10 min in the presence of ATP as described previously (15). Column 1, purified active DNA-PK (assayed before a preincubation with ATP); column 2, phosphorylated inactivated DNA-PK that was preincubated with ATP for 10 min; column 3, MO59J extract treated with Me2SO added to phosphorylated inactivated DNA-PK; column 4, MO59J extract treated with OA added to phosphorylated inactivated DNA-PK; column 5, MO59J extract treated with fostriecin added to phosphorylated inactivated DNA-PK.

Although our data show that a PP2A-like enzyme can dephosphorylate DNA-PK that has been phosphorylated in vitro, it was important to determine whether DNA-PK is phosphorylated in vivo. Ku70 and Ku80 have previously been shown to be phosphorylated on serine in vivo (18); however, to date, in vivo phosphorylation of DNA-PKcs has not been reported. Human lymphoblastoid (BT) cells were therefore incubated with 32P-labeled inorganic phosphate in phosphate-free RPMI in the absence or presence of okadaic acid (1 µM final). DNA-PKcs and Ku70/Ku80 subunits were immunoprecipitated with a monoclonal antibody (42-27) and a polyclonal antibody to recombinant Ku70 (Ab68) respectively, transferred to nitrocellulose membrane, and exposed to x-ray film. Treatment with okadaic acid was found to cause a dramatic increase in the in vivo phosphorylation of DNA-PKcs (Fig. 6A), indicating that DNA-PK undergoes reversible phosphorylation in vivo. In addition, treatment of cells with okadaic acid increased the endogenous levels of phosphorylation of both Ku70 and Ku80 (Fig. 6C), suggesting that both DNA-PKcs and Ku are modified by reversible phosphorylation in vivo. Western blotting of the membrane with antibodies to either DNA-PKcs (Figs. 6B) or Ku70 (Fig. 6D) shows that equal amounts of protein were immunoprecipitated.


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Fig. 6.   DNA-PKcs and Ku70/Ku80 are phosphorylated in vivo in the presence of OA. Human lymphoblastoid cells were incubated in the presence of Me2SO (control; lane 1) or OA (1 µM final concentration; lane 2) for 1 h. [32P]Orthophosphate was then added to these cells, and they were incubated for a further 1 h. Extracts were prepared as described under "Experimental Procedures." A monoclonal antibody (42-27) was used to immunoprecipitate DNA-PKcs, and a mouse polyclonal antibody made to recombinant Ku70 (Ab68) was used to immunoprecipitate Ku70/Ku80 subunits. The immunoprecipitated DNA-PKcs and Ku70/Ku80 subunits were analyzed by SDS-PAGE followed by electroblotting to nitrocellulose. The nitrocellulose was first analyzed by autoradiography (A and C), followed by Western analysis of the same blot using a polyclonal antibody (DPK1) specific to DNA-PKcs and a rabbit polyclonal Ku70 antibody (Ab31A) specific to Ku70 (B and D, respectively). Similar results were obtained in three separate experiments whether okadaic acid was added together with the [32P]orthophosphate or the okadaic acid was added to cells for 1 h prior to the addition of [32P]orthophosphate (data not shown).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Purified DNA-PKcs, Ku70, and Ku80 undergo DNA-dependent phosphorylation in vitro, and that phosphorylation is associated with loss of DNA-PK protein kinase activity (15). We have previously mapped the major in vitro Ku phosphorylation sites, and identification of the in vitro phosphorylation sites in DNA-PKcs is in progress.2 In this study, we show that the catalytic subunit of either PP1 or PP2A is capable of dephosphorylating DNA-PKcs, Ku70, and Ku80 in vitro and that this results in restoration of DNA-PK protein kinase activity. These data therefore strongly suggest that phosphorylation of one or more of the DNA-PK subunits is directly responsible for the loss of protein kinase activity. We estimate that in vitro, DNA-PKcs was phosphorylated on at least seven sites after 1 h of incubation under the conditions used in these experiments. Phosphorylation occurs predominantly on serine residues (15). Studies with synthetic peptides indicate that the protein phosphorylation consensus for DNA-PKcs is serine or threonine followed by glutamine (19). However, DNA-PK can also phosphorylate serines that are followed by tyrosine (16, 26), or valine (29), suggesting that the phosphorylation recognition motif may be more complex than originally proposed. The cDNA sequence of DNA-PKcs predicts 29 serine or threonine residues that are in an SQ or TQ motif, any of which could be a potential autophosphorylation site. The Ku heterodimer is also phosphorylated at several sites in vitro, one major site on Ku70 and at least three on Ku80 (16). At this time, it is not known if one single phosphorylation event is sufficient to inactivate DNA-PK or whether several phosphorylation events contribute to loss of protein kinase activity. We previously showed that DNA-PK-phosphorylated Ku70 is able to bind DNA and that phosphorylated DNA-PKcs dissociates from DNA-bound Ku (15), suggesting that the loss of kinase activity is due to inactivation of DNA-PKcs and not Ku. However, at this time, we cannot exclude the possibility that phosphorylation of Ku also plays a role in regulating the activity of DNA-PK.

We show, for the first time, that DNA-PKcs is phosphorylated in vivo. In preliminary experiments, low levels of phosphorylation of DNA-PKcs were observed in the absence of okadaic acid3; however, here we show that the addition of okadaic acid significantly increased the phosphorylation state of DNA-PKcs in vivo (Fig. 6A). These data suggest that DNA-PKcs is the target of protein phosphatases and protein kinases in vivo and that the activity of protein phosphatases is required to maintain DNA-PK in a highly active state. A prediction of this hypothesis is that DNA-PK would be expected to be largely dephosphorylated in its active form and highly phosphorylated in its inactive form. We have shown that, in vitro, autophosphorylation results in loss of DNA-PK protein kinase activity, again suggesting that DNA-PK is active in a dephosphorylated form and inactive in its phosphorylated form.

In vitro, co-immunoprecipitation experiments suggest that autophosphorylation of DNA-PKcs results in dissociation of phosphorylated DNA-PKcs from phosphorylated DNA-bound Ku (15). Again, this is consistent with phosphorylation resulting in an inactive form of DNA-PK. It remains to be seen whether DNA-PK is regulated by autophosphorylation or phosphorylation by other protein kinases in vivo. These observations also have implications for the repair of DNA double strand breaks by nonhomologous end joining in vivo. We speculate that Ku and DNA-PKcs assemble at the site of a DNA strand break in an ordered fashion, thus generating the active protein kinase. Subsequently, phosphorylation of DNA-PK, either by itself or by another protein kinase, may serve to inactivate DNA-PK and possibly remove DNA-PKcs from the site of the DNA lesion.

DNA-PKcs is a member of a growing family of serine/threonine protein kinases that bears significant amino acid similarity to the catalytic domain of phosphoinositide 3 kinases (30-32). Interestingly, the p110 subunit of the class 1A phosphoinositide-3 kinases has serine/threonine protein kinase activity toward the Src homology 2 domain-containing targeting subunit, p85, and increased incorporation of phosphate into p85 is associated with a dramatic decrease in phosphoinositide 3-kinase activity (33). Also, autophosphorylation of the p110delta subunit of the class II phosphoinositide 3 kinases down-regulates its lipid kinase activity (34). Together, these data suggest that phosphorylation-dependent inactivation may be a characteristic of the phosphoinositide 3-kinase-like family of enzymes.

We have demonstrated the presence of a protein phosphatase activity in crude extracts of human cells that is capable of dephosphorylating DNA-PKcs, Ku70, and Ku80 and restoring the kinase activity of phosphorylated inactivated DNA-PK. This activity has properties expected of a PP2A-like enzyme, since its activity is inhibited at PP2A-selective concentrations of okadaic acid and fostriecin. Further studies will be required in order to identify the protein phosphatase responsible for dephosphorylating DNA-PK in vivo.

    ACKNOWLEDGEMENTS

We thank Seong-Cheol Son for supplying highly purified DNA-PKcs and Ku for these studies. Doug Chan is acknowledged for preliminary in vivo labeling experiments. We also thank Dr. Michel Roberge for providing a sample of fostriecin and Dr. Jim Lees-Miller for reading the manuscript.

    FOOTNOTES

* This work was supported by Canadian Institutes of Health Research Grant 13639 (to S. P. L. M.) and the Natural Sciences and Engineering Council of Canada (to G. M.).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Dagger Postdoctoral Fellow of the Alberta Heritage Foundation for Medical Research.

§ Senior Scholar of the Alberta Heritage Foundation for Medical Research. To whom correspondence should be addressed. Tel.: 403-220-7628; Fax: 403-289-9311; E-mail: leesmill@ucalgary.ca.

Published, JBC Papers in Press, March 16, 2001, DOI 10.1074/jbc.M011703200

2 Y. Yu, D. Chan and S. P. Lees-Miller, unpublished results.

3 D. Chan and S. P. Lees-Miller, unpublished data.

    ABBREVIATIONS

The abbreviations used are: PP1, PP2A, and PP2B, protein phosphatase 1, 2A, and 2B, respectively; DNA-PK, DNA-dependent protein kinase; DNA-PKcs, DNA-PK catalytic subunit; OA, okadaic acid; PAGE, polyacrylamide gel electrophoresis; AMP-PNP, 5'-adenylyl-beta ,gamma -imidodiphosphate.

    REFERENCES
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ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
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