s-phase arrest by reactive nitrogen species is bypassed by okadaic acid, an inhibitor of protein...

/freeradbiomed

Free Radical Biology & M

Original Contribution

S-phase arrest by reactive nitrogen species is bypassed by okadaic acid,

an inhibitor of protein phosphatases PP1/PP2A

Priya Ranjan, Nicholas H. Heintz *

Department of Pathology and Vermont Cancer Center, University of Vermont College of Medicine, Burlington, VT 05405, USA

Received 17 March 2005; revised 3 June 2005; accepted 8 August 2005

Available online 14 November 2005

Abstract

In mammalian cells DNA damage activates a checkpoint that halts progression through S phase. To determine the ability of nitrating agents to

induce S-phase arrest, mouse C10 cells synchronized in S phase were treated with nitrogen dioxide (NO2) or SIN-1, a generator of reactive

nitrogen species (RNS). SIN-1 or NO2 induced S-phase arrest in a dose- and time-dependent manner. As for the positive controls adozelesin and

cisplatin, arrest was accompanied by phosphorylation of ATM kinase; dephosphorylation of pRB; decreases in RF-C, cyclin D1, Cdc25A, and

Cdc6; and increases in p21. Comet assays indicated that RNS induce minimal DNA damage. Moreover, in a cell-free replication system, nuclei

from cells treated with RNS were able to support control levels of DNA synthesis when incubated in cytosolic extracts from untreated cells,

whereas nuclei from cells treated with cisplatin were not. Induction of phosphatase activity may represent one mechanism of RNS-induced arrest,

for the PP1/PP2A phosphatase inhibitor okadaic acid inhibited dephosphorylation of pRB; prevented decreases in the levels of RF-C, cyclin D1,

Cdc6, and Cdc25A; and bypassed arrest by SIN-1 or NO2, but not cisplatin or adozelesin. Our studies suggest that RNS may induce S-phase arrest

through mechanisms that differ from those elicited by classical DNA-damaging agents.

D 2005 Elsevier Inc. All rights reserved.

Keywords: Cell cycle checkpoint; Oxidative stress; Retinoblastoma protein; C10 cells; Okadaic acid; Free radicals

Reactive nitrogen species (RNS) such as nitrogen dioxide

(NO2), nitric oxide, and peroxynitrite (ONOO�) have been

implicated in the pathophysiology of inflammatory lung

diseases such as asthma, chronic obstructive pulmonary disease,

cystic fibrosis, acute respiratory distress syndrome, and idio-

pathic pulmonary fibrosis [1,2]. Among the various cell types

which comprise the lung, epithelial cells of the alveolar structure

seem to be a major target for RNS-mediated injury [2]. Alveolar

epithelial cells are of two types: type I and type II cells. It is now

well established that type II epithelial cells are responsible for

regeneration of alveolar epithelium, as repair of damaged

alveolar surface is dependent on their ability to replicate and

provide progenitor cells that have the potential to undergo

transition into type I cells [3–5]. Irrespective of source, which

can be environmental (such as air pollution) or endogenous

0891-5849/$ - see front matter D 2005 Elsevier Inc. All rights reserved.

doi:10.1016/j.freeradbiomed.2005.08.049

Abbreviations: RNS, reactive nitrogen species; ATM, ataxia telangiectasia

mutated; NO2, nitrogen dioxide; CDK, cyclin-dependent kinase; pRB,

retinoblastoma protein; SIN-1, 3-morpholinosydnonimine; OKA, okadaic acid;

CDDP, cisplatin.

* Corresponding author. Fax: +1 802 656 8892.

E-mail address: [email protected] (N.H. Heintz).

(inducible NO synthase activity due to inflammation), RNS can

induce cell injury in the airway by targeting various cell

components and inducing covalent modification of macromo-

lecules [6,7].

The cellular proteins targeted by RNS may include signaling

molecules, transcription factors, the cytoskeleton, and path-

ways of cell cycle regulation. Recent reports suggest that RNS

can activate the cell membrane death-receptor FAS pathway to

induce S-phase arrest and apoptosis [8,9]. RNS has also been

shown to induce nitration of tyrosine moieties, which has been

shown to occur in patients with asthma and other inflammatory

lung diseases in a manner which directly correlates with the

severity of the disease [10].

To defend against damage induced by various stressors,

proliferating cells activate signaling pathways (or checkpoints)

that induce cell cycle arrest, which in turn protect cells against

the genotoxic consequences of continuing through the cell

cycle in the presence of damage [11]. When DNA damage or

oxidative stress is encountered during the S phase of the cell

cycle, two major responses are prevention of entrance into

mitosis and suppression of further DNA replication [12].

edicine 40 (2006) 247 – 259

www.elsevier.com/locate

P. Ranjan, N.H. Heintz / Free Radical Biology & Medicine 40 (2006) 247–259248

However, the signaling mechanisms by which DNA damage is

detected remain incompletely described. Recent evidence

suggests that DNA damage and a variety of other stressors

induce rapid activation of ATM (ataxia telangiectasia mutated)

and ATR (Rad3-related kinase), which then phosphorylate

Chk2 and Chk1, respectively. Once activated by ATM/ATR,

Chk1 and Chk2 phosphorylate Cdc25 phosphatase and thereby

target the phosphatase for proteasome-mediated degradation.

This prevents Cdc25 from activating downstream pathways

required for S-phase progression [11,12].

Progression of cells into and through S phase is a tightly

controlled process that involves cyclins, cyclin-dependent

kinases (CDKs), CDK inhibitors, retinoblastoma protein, and

other protein factors [13,14]. Many of these proteins require

active phosphorylation and dephosphorylation for G1/S tran-

sition and progression through S phase, thereby implicating the

involvement of specific kinases and phosphatases. The

retinoblastoma tumor suppressor protein (pRB) is a negative

regulator of cell proliferation [15]. pRB is expressed through-

out the cell cycle, but its antiproliferative activity is neutralized

by phosphorylation during the G1/S transition. In the hypopho-

sphorylated form, pRB binds to the members of the E2F family

of transcription factors, thereby negatively regulating transcrip-

tion of E2F-dependent genes that are required for entry into and

transition through S phase [13–15]. During the G1 to S

transition, phosphorylation of pRB is initiated by cyclin D-

dependent kinases and is completed by cyclin E–Cdk2 and

cyclin A–Cdk2. CDKs are activated by isoforms of Cdc25, a

dual phosphatase that dephosphorylates CDKs [16,17]. In

mammals Cdc25A is considered to be a critical regulator of

G1/S transition [18]. CDK activity also is negatively regulated

by cyclin-dependent kinase inhibitors, including p21Cip1,

p27Kip1, and p57Kip1 [19].

In addition to regulating the G1- to S-phase transition, pRB

and its homolog p107 also play a role in controlling progression

through S phase in response to DNA damage [20,21]. One

mechanism involves inhibition of Cdk2 activity and disruption

of PCNA function [22]. Studies with phosphorylation mutants

indicate pRB is reactivated by dephosphorylation [23], which is

catalyzed by specific serine–threonine phosphatases [24].

Whereas the regulation of pRB by CDKs has been studied

extensively, the role(s) of protein phosphatases in controlling

pRB is only partially understood. Interestingly, both PP1 and

PP2A phosphatases have been shown to be involved in

regulating phosphorylation of pRB as well as S-phase progres-

sion [25–28].

Though various biological effects of RNS have been

reported, very few reports explain the effect of these agents

on the checkpoint mechanisms of proliferating cells. In the

present study, we demonstrate that RNS can trigger an intra-S-

phase checkpoint in lung alveolar type II epithelial cells

through mechanisms that correlate with the phosphorylation

state of pRB. Inhibition of dephosphorylation of pRB and other

proteins by okadaic acid, an inhibitor of PP1/PP2A phospha-

tases, not only rescued cells from RNS-induced S-phase arrest,

but also prevented degradation of cyclin D1, Cdc25A, RF-C,

and Cdc6, all of which decline during RNS-induced S-phase

arrest. Further, experiments with a cell-free DNA replication

assay suggest that arrest is not due to inhibition of replication

forks at the level of elongation due to damaged DNA

templates, but rather may involve posttranslational modifica-

tion of proteins that control DNA synthesis.

Materials and methods

Cell culture, synchronization, and RNS treatment

Murine type II alveolar C10 cells were maintained in CMRL

medium supplemented with 10% fetal bovine serum (FBS)

containing 100U/ml penicillin and 100 Ag/ml streptomycin [29].

To synchronize cells in S phase, cells were first arrested in G0/

G1 by incubation in medium containing 0.2% FBS for 72 h and

then were induced to reenter the cell cycle by adding Dulbecco’s

modified Eagle’s medium containing 10% FBS for 16 h. Cell

cycle progression was evaluated by flow cytometry as described

previously [30]. For exposure to RNS, S-phase cells (6.25� 105

cells/60-mm dish) were treated with 8 ppm NO2 using a rocking

platform [29] or with different concentrations of SIN-1 (3-

morpholinosydnonimine; Calbiochem), a generator of RNS.

After 3 h of exposure in S phase, cells were either harvested or

allowed to recover in fresh medium for 3 or 6 h. Okadaic acid

(Sigma) was added during exposure and recovery as described in

the text. Progression through the cell cycle was monitored by

flow cytometry [30]. Cells were also treated with cisplatin or the

alkylating agent adozelesin as positive controls.

Immunoblotting

Immunoblotting was performed with total cell lysates as

described previously [29,30]. Briefly, cells were rinsed with PBS

and lysed in 2� SDS sample buffer, and equal amounts of

soluble protein were resolved by electrophoresis on 8 or 14%

SDS–polyacrylamide gels. Proteins were then transferred to

Immobilon-P membranes (Millipore) by electroblotting. Blots

were blocked in 4% nonfat dry milk or 4% BSA fraction V in

TBS-T as described [30]. Blots were then probed with the

indicated primary antibodies and then with the appropriate

horseradish peroxidase-coupled secondary antibody. Signals

were detected by the ECL system (Amersham, Piscataway, NJ,

USA). h-Actin was used as a loading control. Antibodies to h-actin, cyclin D1, RF-C, and Cdc6 were purchased from Santa

Cruz; pRB and p21 from BD Pharmingen; and total ATM and

phospho-ATM from Upstate. Antibody to Cdc25A was a gift

from W. Burhans (Roswell Park Cancer Institute, Buffalo, NY,

USA).

Nitrite/nitrate assay

Cell-free culture supernatants were assayed for nitrite

concentrations by a microplate assay method as described

previously [31]. Briefly, 100 Al of culture supernatant was

incubated with an equal volume of Griess reagent (one part 1%

sulfanilamide in 2.5% H3PO4 plus one part 0.1% naphthy-

lethylene diamine dihydrochloride in distilled water) at room

P. Ranjan, N.H. Heintz / Free Radical Biology & Medicine 40 (2006) 247–259 249

temperature for 10 min. The absorbance was taken at 540 nm in

a microtiter plate reader. Nitrite concentration was quantified

by using sodium nitrite as standard.

In vitro run-on replication assay

The ability of cells to support DNA synthesis was monitored

using a run-on replication assay described previously [32]. For

isolation of nuclear and cytoplasmic extracts, treated or

untreated C10 cells in 150-cm2 plates were washed with ice-

cold PBS and rinsed twice with ice-cold hypotonic buffer (20

mM N-2-hydroxyethylpiperazine-N V-2-ethanesulfonic acid

(Hepes)–KOH (pH 7.5), 5 mM KCl, 1.5 mM MgCl2, 0.1 mM

dithiothreitol) and plates were drained extensively before

harvesting with a rubber policeman. Under these conditions

each 100-mm plate yielded approximately 200 Al of cell lysate.The cells were incubated on ice for 10 min and disrupted by

Dounce homogenization until 95% of the cells were broken (20

strokes of a B pestle), and the cell lysate was incubated on ice for

30 min before centrifugation at 10,000 g for 10 min at 4-C. Thesupernatant (cytosol fraction) was removed and either used

immediately or frozen at�80-C. Concentration of protein in thecytosolic extracts was determined using a Bio-Rad protein assay

kit. The nuclear pellet was suspended in 60 Al (1�105 to 5� 105

nuclei/Al) of hypotonic buffer per plate with or without 10%

sucrose (nuclear fraction) or directly in the cytosolic extracts

from cells subjected to the indicated treatments. Phase-contrast

microscopy was used to confirm the integrity of nuclei at each

stage.

DNA synthesis assay

Reaction mixtures contained (final concentration) 30 mM

Hepes–KOH (pH 7.5); 7 mM MgCl2; 0.8 mM DTT; 100 AMeach dTTP, dGTP, and dCTP; 25 AM [a-32P]dATP; 200 AMeach CTP, GTP, and UTP; 4 mM ATP; 40 mM creatine

phosphate; and 20 Ag of creatine phosphokinase (rabbit

muscle type I; Sigma Chemical Co.) per milliliter. Standard

reaction mixtures were prepared by adding 50 Al of a 5�reaction buffer mix containing all the above components and

5 � 106 nuclei suspended with 200 Al of cytosolic extract.

Reaction mixtures were prepared on ice and incubated at

37-C for 1 h. Reactions were terminated by addition of an

equal volume of lysis buffer (40 mM EDTA, 1.2% SDS, 100

mM NaCl, 50 mM Tris–Cl (pH 8.0)). The lysate (500 Al)was incubated with proteinase K, and DNA was purified

from the lysate using phenol–chloroform extraction and

ethanol precipitation. Equivalent amounts of cellular DNA

were resolved on agarose gels, and the gels were stained with

ethidium bromide, photographed, dried, and exposed to X-ray

film for 24 h. Incorporation of [a-32P]dATP into DNA was

quantified with a phosphoimager (Bio-Rad).

Comet assays

Cells were harvested by scraping with a rubber policeman

and resuspended in PBS. Twenty microliters of cell suspension

containing 15,000 cells was mixed with 85 Al of melted low-

melt agarose, layered on agarose-precoated microscope slides,

and allowed to solidify on ice for 10 min. Another layer of

low-melt agarose was added and slides were processed for

comet assays as described [33]. Briefly, slides with the

agarose-embedded cells were first subjected to a lysis step

(1-h incubation at 4-C in 1% SDS, 2.5 M NaCl, 100 mM

EDTA, 1% Triton X-100, 10% dimethyl sulfoxide) and then

placed for 20 min in an ice-cold electrophoresis chamber

containing alkaline electrophoresis buffer (300 mM NaOH, 1

mM EDTA) to allow DNA denaturation. The electrophoresis

was conducted for 20 min at 25 V and 300 mA, and the slides

were washed with neutralization buffer (40 mM Tris–HCl, pH

7.4), stained with ethidium bromide overnight, and analyzed on

a fluorescence microscope provided with epifluorescence and

equipped with a rhodamine filter (excitation wavelength 546

nm, barrier 580 nm). The images of 100 randomly chosen cells

per slide were captured and analyzed with a digital camera

system. DNA damage was determined by measuring the length

of DNA migration (total comet length) using an eyepiece

micrometer.

Statistical analysis

Results are expressed as means T standard deviation (SD) of

at least three independent experiments. The statistical signif-

icance of difference between test groups was analyzed by two-

tailed Student’s t test. The level of significance was considered

to be p < 0.05.

Results

The C10 cell cycle

To synchronize cells in the S phase, mouse type II

alveolar lung C10 cells first were arrested in G0/G1 by

incubation in 0.2% FBS for 72 h and then incubated in fresh

medium with 10% FBS to induce cell cycle reentry. Cell

cycle progression was analyzed by flow cytometry (Fig. 1A

and Table 1). Cells began to enter the S phase by 12 h, and

by 16 h the majority of the cycling population was in S

phase. By 18 h cells had begun to accumulate in G2/M, and

by 21 h had returned to G1 (Fig. 1A). Western blot analysis

showed that pRB was dephosphorylated in serum-starved

C10 cells (Fig. 1B, lane 1). By 9 h after serum stimulation

pRB was largely phosphorylated (lane 4), and this pattern of

hyperphosphorylation was maintained until entry into G2/M

at 18 h (Fig. 1B, lanes 5–8). By 21 h, when cells had begun

to reenter G1 (Fig. 1A), dephosphorylation of ¨50% of pRB

was evident (Fig. 1B, lane 9).

Compared to the loading control h-actin, progression

through the cell cycle also was associated with increased

expression of cyclin D1, Cdc25A, Cdc6, and RF-C (Fig. 1B).

Expression of CDK inhibitor p21 was observed in serum-

starved cells, and its levels were reduced only at later time

points when cells had begun to reenter G1 (Fig. 1B, lanes

8 and 9). These data are consistent with synchronization of

Fig. 1. Kinetics of cell cycle progression in synchronized C10 cells after serum

stimulation. (A) C10 cells were arrested in G0/G1 by incubation in 0.2% FBS for

72 h (denoted 0 hr) and then were induced to reenter the cell cycle by incubation

inmedium containing 10%FBS. Cultures were harvested at indicated time points

and analyzed for cell cycle progression by flow cytometry. By 16 h, the majority

of the population was in S phase. (B) At the indicated times, extracts of C10 cells

were prepared and analyzed for expression of pRB, cyclin D1, Cdc25A, Cdc6,

RF-C, and p21 by immunoblotting. Note that phosphorylation of pRB at the G1

restriction point preceded entry into the S phase. Data shown are representative of

three independent experiments.


C10 cells by serum deprivation and document normal cell

cycle progression over a 24-h period in response to 10%

FBS.

Table 1

Effect of reactive nitrogen species on S-phase progression

Nitrite (AM) 16 h S-phase cells

% S % G2/M

Medium 8.6 T 1.6a 58.6 T 5.8 11.8 T 2.9

NO2 (8 ppm) 461 T 6.3* � �SIN-1 (0.5 mM) 287.3 T 10.6* � �SIN-1 (1 mM) 441 T 9.7* � �Cisplatin (25 AM) 5.1 T 1.3 � �Adozelesin (20 nM) 9.6 T 2.1 � �Synchronized S-phase C10 cells were treated with NO2, SIN-1, CDDP, or adozeles

acid (300 nM) and then incubated in fresh medium for another 6 h with or without

supernatants were assayed for nitrite by the method described under Materials and m

of S phase; values for the percentage of cells in S and G2/M are shown.a The numbers represent means T SD of triplicate sets.

* p < 0.05 versus values for untreated control.

RNS induce an S-phase arrest

To determine the effects of RNS on S phase, serum-starved

C10 cells were incubated in fresh medium with 10% FBS for

16 h, conditions that reproducibly resulted in 45–70% S-phase

cells (Figs. 1A and 2A). S-phase cultures then were treated

with SIN-1, cisplatin, or adozelesin, or exposed directly to pure

NO2 gas, for 3 h. Cells were harvested for analysis at the end of

exposure (denoted 3 hr exp) or after recovery for 3 or 6 h in

fresh medium containing 10% FBS. Progression out of S phase

was monitored by flow cytometry, and the levels of various

proteins in cell extracts were assessed by immunoblotting.

In control cells, incubation of S-phase C10 cells (e.g., cells

serum stimulated for 16 h) for additional periods of time

allowed cells to exit the S phase and collect in G2/M and G1.

For example, after an additional 6–9 h of culture all S-phase

cells had reentered G1 (Fig. 2A). Exposure to 0.5 mM SIN-1

for 3 h induced a delay in S-phase progression, as the majority

of S-phase cells were able to transit G2/M and reenter G1 after

incubation in fresh medium for 6 h (Fig. 2A). However,

treatment with 1 mM SIN-1 caused ¨49% of the cell

population to remain in S phase, even after an additional 6

h of recovery in fresh medium (Fig. 2A and Table 1). Indeed, in

response to 1.0 mM SIN-1, flow cytometry suggested that cells

continued to enter, but not exit, the S phase. Exposure to 8 ppm

pure NO2 gas for 3 h caused S-phase arrest, and after an

additional 6-h recovery period, cells remained in S phase (Fig.

2A and Table 1).

The nitrite/nitrate level in the culture supernatants accumu-

lated in a dose- and time-dependent manner with SIN-1

treatment (Table 1 and data not shown). For example, the

nitrite levels in the culture medium after the 3-h exposure

period with 0.5 and 1 mM SIN-1 were found to be ¨287 and

441 AM, respectively. After exposure to 8 ppm NO2 for 3 h, the

nitrite level was found to be ¨461 AM (Table 1).

S-phase cells were also treated with the DNA-damaging

agents cisplatin (25 AM) and adozelesin (20 nM) as positive

controls. Both agents arrested cells in S phase, and both

prevented cells from exiting S phase after a 6-h recovery period

(Fig. 2A).

6 h recovery

�OKA +OKA (300 nM)

% S % G2/M % S % G2/M

27.3 T 2.9 26.2 T 3.1 35.2 T 4.1 22.3 T 3.9

47.8 T 5.6 16.2 T 3.6 24.8 T 2.9 23.2 T 6.1

37.2 T 2.7 21.6 T 4.8 26.4 T 3.1 24.6 T 3.7

49.1 T 9.1 12.5 T 3.5 32.3 T 4.4 23.4 T 4.1

51.0 T 7.9 11.8 T 2.7 48.2 T 8.5 14.6 T 5.5

59.3 T 8.2 5.1 T 1.1 57.2 T 7.2 7.2 T 3.1

in at the indicated concentrations for 3 h in the presence or absence of okadaic

OKA. Cells were analyzed for cell cycle by flow cytometry. Cell-free culture

ethods after 3 h of exposure. Flow cytometry was used to assess progression out

Fig. 2. Reactive nitrogen species inhibit S-phase progression. (A) C10 cells synchronized in S phase (16 h post-serum stimulation) were treated with the nitrating agent

SIN-1 or NO2 or with the DNA-damaging agent cisplatin (CDDP) or adozelesin, at the indicated concentrations. After 3 h of exposure, cells were harvested or washed

and allowed to recover for an additional 3 or 6 h in freshmediumwith 10%FBS. Flow cytometry was used to assess cell cycle progression. See Table 1 for quantification

of percentage S and G2/M cells under these conditions. (B) S-phase cells treated with SIN-1, CDDP, or adozelesin (Adz) at the indicated concentrations were also

analyzed for expression of pRB, cyclin D1, Cdc25A, Cdc6, RF-C, and p21 by immunoblotting. h-Actin was used as a loading control. (C) S-phase C10 cells were

exposed directly to NO2 (8 ppm) for 3 h and cell extracts were prepared (denoted 3 hr exp). Replicate cultures were allowed to recover from NO2 exposure for 3 or 6

h before preparation of cell extracts. As controls, serum-stimulated cells were harvested in S phase (16 h) or after completion of S phase (16 + 6 h). Cell extracts were

analyzed for expression of pRB, cyclin D1, Cdc25A, Cdc6, RF-C, and p21 by immunoblotting as before. S-phase arrest was associated with dephosphorylation of pRB;

decreased expression of cyclin D1, Cdc25A, Cdc6, and RF-C; and increased levels of p21. Data shown are representative of three independent experiments.


Effects of RNS on cell cycle and S-phase modulators

We next investigated the effects of RNS on cell-cycle

regulators involved in progression through S phase. Whereas

pRB is a negative regulator of progression through G1, it has

also been shown to be involved in S-phase progression [20–

23]. Treatment of S-phase C10 cells for 3 h with 0.5 or 1 mM

SIN-1 resulted in partial dephosphorylation of pRB, as did

exposure to CDDP and adozelesin (Fig. 2B, lanes 2–5). NO2

was the most effective agent in inducing dephosphorylation of

pRB (Fig. 2C), which was almost completely dephosphory-

lated by the end of the 3-h exposure period (lane 3) and

remained so during 6 h of recovery in fresh medium (lane 5).

Activation of the S-phase checkpoint is associated with

active degradation of cyclin D1 [34] and Cdc25A [35].

Treatment of S-phase C10 cells with SIN-1, cisplatin,

adozelesin, or NO2 also resulted in a time-dependent decrease

in the levels of cyclin D1 and Cdc25A (Figs. 2B and 2C),

indicating arrest is accompanied by proteolytic degradation of

these factors. The status of the S-phase replication factors RF-C

and Cdc6 was also investigated during arrest and recovery.

Decreased levels of these factors were observed over time in

response to all four agents (Figs. 2B and 2C).

The CDK inhibitor p21 has been shown to be up-regulated

by p53 in response to DNA damage [36] or by oxidants in a

p53-independent manner [37]. Because p21 can inhibit pRB

phosphorylation by inhibiting CDKs, and can interact with

PCNA to inhibit DNA synthesis [22], we examined the status

of p21 in RNS-induced S-phase arrest. In S-phase C10 cells

treatment with SIN-1, cisplatin, or adozelesin (Fig. 2B), or

exposure to NO2 gas (Fig. 2C), resulted in increased levels of

p21 in a dose- and time-dependent manner. Together these data


indicate that cisplatin, adozelesin, and RNS induce S-phase

arrest through pathways that include dephosphorylation of pRB,

degradation of cyclin D1 and Cdc25A, and induction of p21.

Inhibition of serine–threonine phosphatases prevents S-phase

arrest induced by RNS

Phosphorylation and dephosphorylation of pRB play a role

in controlling progression through the cell cycle under normal

as well as stress conditions [38,39]. Dephosphorylation of pRB

has been observed in response to various genotoxic stimuli [20],

and serine–threonine phosphatases have been implicated as

major pRB phosphatases (see introduction). In an attempt to

modulate the phosphorylation status of pRB in RNS-induced S-

phase arrest, cells were treated with okadaic acid (OKA), a

serine–threonine phosphatase inhibitor. Treatment of S-phase

C10 cells with doses of 100–300 nM OKA did not prevent cells

from exiting S phase (Fig. 3A and Table 1), although cell death

was observed at concentrations higher than 350 nM (data not

shown). As expected, cells exposed to 0.5 mM SIN-1 showed

minimal response to OKA treatment, for in the presence or

absence of OKA cells most cells were able to complete S phase

and reenter G1 after 6 h of recovery (Fig. 3A and Table 1). In

contrast, S-phase cells treated with 1.0 mM SIN-1 showed a

graded response to OKA, and only in the presence of 200–300

nM OKAwere cells able to bypass the S-phase arrest (Fig. 3A).

OKA also prevented NO2-induced S-phase arrest in C10 cells in

a dose-dependent manner (Fig. 3B). Whereas 100 nM OKAwas

insufficient to prevent arrest by NO2, cells treated with NO2 in

the presence of 300 nM OKA were observed in G2/M and G1

after 3 h of recovery and in G1 after 6 h (Fig. 3B). In cells

exposed to NO2 in the presence of 100 nM OKA, cells seemed

to continue to enter, but not exit, S phase during recovery (Fig.

3B). In contrast, OKAwas not able to prevent S-phase arrest by

either cisplatin or adozelesin under any condition (Table 1 and

data not shown), even though at 300 nM OKA a fraction of the

pRB in CDDP-treated cells remained phosphorylated (Fig. 3C,

lane 12). In contrast, addition of OKA prevented dephosphory-

lation of pRB in cells treated with either SIN-1 or NO2 in a

dose-dependent manner (Fig. 3C, lanes 1–8).

OKA also prevented RNS-induced degradation or down-

regulation of Cdc25A, cyclin D1, Cdc6, and RF-C in a dose-

dependent fashion (Fig. 3C). Interestingly, decreases in the

levels of cyclin D1, Cdc25A, Cdc6, and RF-C were completely

prevented by 300 nM OKA in cells treated with cisplatin (Fig.

3C, lanes 9–12), even though cells treated with cisplatin failed

to exit S phase (Table 1). In contrast, OKA did not alter the levels

of p21 expression, which remained elevated in response to RNS

or cisplatin (Fig. 3C). Other phosphatase inhibitors, such as

sodium vanadate and sodium fluoride, did not prevent S-phase

arrest by RNS at any concentration tested (data not shown).

RNS and inhibition of DNA synthesis in vitro

S-phase arrest by DNA damage depends on the extent and

duration of the DNA damage, cell type, and cell cycle stage.

Exposure of S-phase C10 cells to RNS induces an intra-S-

phase checkpoint, but the immediate targets of RNS, which

have limited cell permeability, are not known. To assess the

role of DNA damage in S-phase arrest by RNS, we examined

the ability of nuclei from treated cells to support chromosomal

DNA synthesis in vitro using a cell-free, run-on replication

assay [32]. Previously it was demonstrated that cytosolic

fractions from S phase contain cellular proteins that support

high rates of replication fork elongation in vitro [32]. One

advantage to the reconstituted cell-free system is that no new

initiation events occur in isolated nuclei in vitro [32], providing

a method for selectively examining effects on the elongation of

preexisting replication forks.

First we examined the effect of SIN-1 (1 mM) or CDDP (25

AM) on the reconstituted nuclei plus cytosol replication assay

(Fig. 4). S-phase C10 cells were treated with different

concentrations of SIN-1 or cisplatin for the indicated periods

of time, nuclei and cytosolic extracts were prepared, and these

fractions were then reconstituted for the in vitro replication

assay as described under Materials and methods. Fig. 4A

shows representative results of experiments in which nuclei

and cognate cytosol from S-phase cells treated with 1 mM SIN-

1 or 25 AM cisplatin were assayed for DNA replication in vitro.

No inhibition of DNA synthesis was observed in vitro after 1

h of treatment with either agent (Fig. 4A, lanes 3 and 4).

However, treatment for 3 h resulted in complete inhibition of

DNA replication when nuclei and cytosol were reassembled for

in vitro DNA synthesis assay (Fig. 4A, lanes 5 and 6). Note

that after treatment with either SIN-1 or cisplatin for 3 h, or for

3 h with up to 6 h of recovery (Fig. 4A, lanes 7–10), there was

no evidence of DNA degradation as judged by agarose gel

electrophoresis.

In order to study the contribution of cytosol to inhibition, S-

phase cells were treated with varying concentrations of SIN-1

or cisplatin for 3 h, and cytosolic extracts then were prepared

and incubated with nuclei from untreated S-phase cells (Fig.

4B). As before, control S-phase nuclei incubated with cytosol

from untreated cells supported DNA replication (Fig. 4B, lane

1). Interestingly, cytosol from cells treated with SIN-1 showed

a biphasic response with regard to the ability to support DNA

synthesis. Cytosol from cells treated with 0.25 mM SIN-1

supported a significant increase in DNA synthesis in nuclei

from untreated S-phase cells compared to controls (Fig. 4B,

lane 3). In contrast, cytosol from S-phase cells treated with

higher doses of SIN-1 (0.5 or 1 mM) markedly inhibited DNA

replication (Fig. 4B, lanes 4 and 5). Cytosol from cells treated

with 10 AM cisplatin also stimulated synthesis above control

levels (Fig. 4B, lane 6), whereas cytosol from cells treated with

higher doses inhibited DNA replication in S-phase nuclei from

untreated cells (Fig. 4B, lanes 7 and 8).

In contrast to the similar biphasic responses observed with

cytosol from cells treated with SIN-1 and cisplatin, nuclei from

cells treated with SIN-1 showed significant differences from

nuclei from cells treated with cisplatin in their ability to support

DNA synthesis. In these experiments, nuclei from cells treated

with various doses of either SIN-1 or cisplatin were incubated

with cytosol from untreated S-phase cells, and replication

activity was measured as before. As shown in Fig. 4C, nuclei

Fig. 3. The protein serine– threonine phosphatase inhibitor okadaic acid prevents RNS-induced S-phase arrest. (A) Serum-stimulated C10 cells collected in S phase

(denoted as 16-h time point) were treated with okadaic acid (OKA) at the indicated concentrations, with or without treatment with 0.5 or 1.0 mM SIN-1, for 3 h. Cells

were then allowed to recover in fresh medium with or without OKA for an additional 6 h. Flow cytometry was used to assess cell cycle progression. (B) As in A,

serum-stimulated C10 cells collected in S phase (denoted as 16 h) were treated with or without OKA at the indicated concentrations, with or without exposure to

8 ppm NO2. After 3 h of exposure (left) cells were harvested for analysis by flow cytometry as before. Note that after 3 h in the absence of NO2 or OKA the control

S-phase cells entered G2/M. Replicate cultures were allowed to recover from exposure in fresh medium with or without 100–300 nM OKA for 3 or 6 h, as indicated.

Flow cytometry was used to assess cell cycle progression. (C) S-phase cells were treated with OKA and SIN-1, NO2, or CDDP at the indicated concentrations for

3 h and then were allowed to recover in fresh medium for 6 h, with or without OKA as indicated. Cell extracts were analyzed for expression of pRB, cyclin D1,

Cdc25A, Cdc6, RF-C, and p21 by immunoblotting as before. OKA inhibited dephosphorylation of pRB and restored the expression levels of cyclin D1, Cdc25A,

Cdc6, and RF-C in a dose-dependent manner, but had little effect on the levels of p21. Data shown are representative of three independent experiments.


from S-phase C10 cells treated with concentrations of SIN-1 up

to 1.0 mM, when incubated with cytosol extract from untreated

S-phase cells, supported DNA synthesis at levels indistin-

guishable from those of nuclei from untreated cells (compare

lane 1 to lanes 2–5). In contrast, nuclei from S-phase cells

treated with 25 or 40 AM CDDP were markedly compromised

in their ability to support DNA synthesis in vitro when

incubated with cytosol extracts from untreated cells (Fig. 4C,

lanes 7 and 8), although lower doses did allow DNA

replication (lane 6). These results suggest that SIN-1 affects

the ability of the cytosolic extract to support DNA replication

in normal S-phase nuclei without compromising the ability of

the nucleus to serve as a template. In contrast, cisplatin targets

both replication factors in the cytosol and the nuclear DNA

template. In support of these data, incubation of cytosol extract

with 0.5 mM SIN-1 for 1 h in vitro completely abolished the

ability of the extract to stimulate DNA replication, whereas

cisplatin had little effect (data not shown).

Fig. 4. Inhibition of nuclear DNA synthesis by RNS. (A) Nuclei were isolated from S-phase C10 cells treated with SIN-1 (1.0 mM) or cisplatin (25 AM) for the

indicated time periods and incubated with their cognate cytosolic extracts in the nuclear run-on replication assay in the presence of [32P]dATP. Genomic DNA

from each reaction was purified and resolved by agarose gel electrophoresis. After staining with ethidium bromide (top), the gel was dried and radiolabeled DNA

was visualized by autoradiography (middle). A phosphoimager was used to quantify the amount of [32P]dATP incorporated into genomic DNA (bottom). See

Materials and methods for experimental details. Note that under these conditions SIN-1 and CDDP were equally effective over time at inhibiting genomic DNA

synthesis. (B) Nuclei were isolated from untreated S-phase C10 cells and incubated with control cytosol (lane 1) or cytosol from S-phase cells treated with the

indicated concentrations of SIN-1 (lanes 2–5) or CDDP (lanes 6–8). Samples were processed for agarose gel analysis as in A. (C) Cytosol isolated from

untreated S-phase C10 cells was incubated with nuclei isolated from control S-phase cells (lane 1) or from cells treated with the indicated concentrations of SIN-1

(lanes 2–5) or cisplatin (lanes 6–8), and DNA synthesis was evaluated as before. Nuclei from cells treated with SIN-1 supported DNA synthesis as well as

nuclei from untreated cells, whereas CDDP inhibited the ability of nuclei to support in vitro DNA synthesis in a dose-dependent fashion. Total signal

incorporated into genomic data was quantified with a phosphoimager and is plotted as CPM/Ag DNA. Data shown are means T SD of three independent

experiments.


Cisplatin, but not RNS, induces DNA strand breaks

The results of the in vitro DNA replication assay indicate

that the targets that mediate S-phase arrest by RNS and

cisplatin may differ. To examine the extent of DNA damage in

cells treated with SIN-1, NO2, or cisplatin, we quantified the

DNA breaks induced by these agents using the comet assay

[33]. The comet assay, which is a sensitive method for early

detection of low levels of DNA breaks in individual cells,

involves embedding individual cells in agarose on a micro-

scopic slide and measuring the degree of migration of nuclear

DNA after application of an electrical field. The extent of

DNA migration is proportional to the number of breaks in

DNA and its evaluation allows indirect measurement of the

DNA damage at the single-cell level. As shown in Figs. 5A

and 5B, exposure to NO2 or treatment with SIN-1 for 3 h did

not induce significant increases in DNA breaks in C10 cells as

assessed after 6 to 48 h of recovery. In contrast, treatment with

cisplatin at 25 AM for 3 h resulted in significant levels of DNA

breaks by 24 h of recovery, and the level of breaks increased

further by 48 h (Figs. 5A and 5B). The comet assays shown in

Fig. 5 were performed under alkaline conditions that report

single-stranded DNA breaks. Comet assays conducted under

neutral buffer conditions to assess the level of double-stranded

DNA breaks induced by these agents failed to detect double-

stranded breaks in RNS-treated cells by 48 h, whereas this type

of damage was readily evident in cells treated with cisplatin by

48 h (data not shown).

Fig. 5. RNS induce minimal DNA damage as assessed by comet assays. (A) Synchronized S-phase C10 cells treated with SIN-1, NO2, or cisplatin for 3 h as

before and then allowed to recover in fresh medium for the indicated periods of time. Cells were processed for comet assays and nuclear DNA was visualized by

staining with ethidium bromide. (B) The degree of DNA damage from each treatment modality was quantified by measuring the length of DNA migration (or

comet). Data shown are means T SD of three replicate samples and are representative of three independent experiments. *p < 0.05 versus values of control

cultures.


RNS induce phosphorylation of ATM

Results of comet assays indicated that RNS may not

induce detectable DNA breaks in S-phase C10 cells,

suggesting that activation of the S-phase checkpoint may

not be entirely dependent on DNA damage. We therefore

investigated phosphorylation of the checkpoint kinase ATM,

which, in addition to DNA damage, has been reported to be

rapidly activated by chromatin modifications in response to

stress [40]. Treatment of C10 cells with SIN-1 for 3

h resulted in phosphorylation of ATM, and ATM remained

phosphorylated in RNS-treated cells during recovery, where-

as the total level of ATM remained unchanged (Fig. 6A).

The positive control cisplatin also induced ATM phosphor-

ylation. Similar effects were observed in NO2 exposure

experiments (Fig. 6B), in which ATM phosphorylation was

evident after 3 h of treatment and gradually decreased during

recovery.

Discussion

To defend against potential damage induced by DNA

damage, mammalian cells activate regulatory mechanisms that

stop proliferating cells in the G1 or G2 phase of the cell cycle

[11,12]. The induction of G1 or G2 arrest prevents replication

or segregation of damaged DNA and hence contributes to the

maintenance of genomic integrity. DNA damage in S phase

induces signaling cascades that block new initiation events,

thereby inhibiting DNA synthesis [12]. Inhibition of progres-

sion through S phase may be transient or permanent, depending

on the type and severity of stimuli [41]. Recent work shows,

however, that deregulation of growth control by oncogenes

induces the DNA damage response, which may represent an

important event in the generation of genomic instability and

neoplastic transformation [42,43]. The source of DNA damage

under these altered growth conditions is unknown, but does not

seem to be linked to reactive oxygen species [42].

Fig. 6. RNS induces phosphorylation of ATM kinase. (A) S-phase C10 cells

were treated with SIN-1 (0.5 or 1 mM) or CDDP (25 AM) for 3 h as before, and

then cells were allowed to recover for the times indicated. Levels of phospho-

ATM (pATM) in cell extracts were detected by immunoblotting. Total levels of

ATM (ATM) were assessed in the same cell extracts using a different gel, with

h-actin serving as a loading control. (B) S-phase cells were treated with NO2

(8 ppm) for 3 h and allowed to recover for the indicated periods of time, and

cell extracts were analyzed for ATM phosphorylation by immunoblotting as

above. As in A, total levels of ATM were assessed in the same extracts using a

different gel, with h-actin serving as a loading control.


Agents which can induce the intra-S-phase checkpoint

include cisplatin, alkylating agents like adozelesin, etoposide,

and free radicals. Among these agents, reactive nitrogen

species have drawn increasing attention in recent years because

of their involvement in the pathophysiology of pulmonary

diseases [1,2]. Though various biological effects of RNS have

been reported [2], the mechanisms by which cells activate

checkpoints in response to RNS-induced potential damage in

lung alveolar cells remain largely undefined. In the present

investigation we demonstrate that RNS induce an intra-S-phase

checkpoint in lung type II C10 epithelial cells through

mechanisms that may differ from those induced by cisplatin

or adozelesin.

Flow cytometry data show that exposure to NO2 or

treatment with SIN-1 results in S-phase arrest in synchronized

C10 cells in a dose- and time-dependent manner and that

these effects are apparent after as little as 3 h of treatment

with NO2. A dose of SIN-1 (0.5 mM) that induces arrest in

G0/G1 [29] induced a temporary delay in S-phase progres-

sion, with cells recovering the ability to continue through the

cell cycle during a 6-h recovery period (Fig. 2A). Arrest

induced by exposure to either 1.0 mM SIN-1 or 8 ppm NO2,

as for cisplatin and adozelesin, was accompanied by changes

in expression of several markers associated with induction of

the S-phase checkpoint by DNA damage, i.e., dephosphory-

lation of pRB and decreased expression of cyclin D1 and

Cdc25A. We also observed decreased levels of RF-C and

Cdc6, which is degraded during apoptosis [35], which were

not measured directly here. Common alterations in the

expression levels of these markers initially suggested that

RNS, like cisplatin and adozelesin, induce S-phase arrest

through a common mechanism. However, several lines of

evidence suggest that S-phase arrest by RNS may be mediated

through additional mechanisms.

First, the effect of the phosphatase inhibitor okadaic acid

on maintenance of the S-phase checkpoint differed between

RNS and cisplatin. One possible target of OKA may be pRB,

which has been shown to be required for intra-S-phase

response to DNA damage [20–23]. The function of

retinoblastoma protein in controlling G1/S phase transition

and progression through S phase is regulated by phosphor-

ylation on serine and threonine residues [13–15]. Whereas

the role of CDKs in phosphorylation and inactivation of pRB

during the G1- to S-phase transition has been characterized in

detail, the roles of protein phosphatases in regulating pRB

function are not well understood. Protein phosphatases PP1

and PP2A have been implicated as major pRB phosphatases

both in vivo and in vitro and have been shown to play a

major role in cell cycle progression through S phase in many

cell types [25,28]. In the present study, okadaic acid, a cell-

permeable agent which inhibits PP1 and PP2A serine–

threonine phosphatase in a concentration-dependent manner

[44], was able to rescue NO2- or SIN-1-induced S-phase

arrest at doses that are reported to inhibit PP1 [44]. In

contrast, at any dose okadaic acid was ineffective in rescuing

cells treated with cisplatin from S-phase arrest, despite the

fact that at 300 nM okadaic acid a fraction of pRB remained

phosphorylated in cells treated with cisplatin. The reason for

this discrepancy is unknown, but may be related to

differences in the activity of replication forks detected in

the in vitro assay (Fig. 4).

Interestingly, OKA also prevented degradation or down-

regulation of cyclin D1, Cdc25A, Cdc6, and RF-C without

altering the levels of p21, including in cells treated with

cisplatin (Fig. 3C), suggesting the levels of these proteins in S

phase are regulated in manner independent of p21. Our

studies suggest that RNS may induce a protein serine–

threonine phosphatase activity that is predominantly respon-

sible for dephosphorylation of pRB and that dephosphoryla-

tion of pRb (and perhaps other targets) is required for S-phase

arrest. Further studies will be required to determine if the

phosphatases are indeed PP1 and PP2A or involve other

proteins.

Second, run-on replication assays suggest that the molecular

targets and the mechanism(s) for induction of the S-phase

checkpoint by RNS differ from those of cisplatin and

adozelesin. If S-phase arrest is mediated at the level of

initiation and not elongation, inhibition of S-phase progression

should correlate with dose-dependent inhibition of replication

fork progression in nuclei from RNS-treated cells in the cell-

free nuclear replication assay, as is observed with cisplatin (Fig.

4). In this system, which does not support initiation of DNA

synthesis at replication origins and requires the activity of

DNA polymerase a, replication is stimulated three- to eightfold

by cytoplasmic factors from S-phase cells [32]. Whereas


cytosol from S-phase C10 cells treated with 1.0 mM SIN-1 (or

cytosolic extracts from S-phase cells that were treated with

SIN-1 in vitro) did not support DNA replication in untreated

nuclei (Fig. 4), nuclei from the same cells displayed control

levels of replication activity when incubated in cytosolic

extracts from untreated S-phase C10 cells. In contrast, both

cytosolic extracts and nuclei from cells treated with cisplatin

showed dose-dependent inhibition of DNA synthetic activity.

Although both RNS and cisplatin induced S-phase arrest, our

data indicate that cisplatin targets both the nuclear template and

factors in the cytosol, whereas RNS seems to preferentially

target factors in the cytosol that are required to support DNA

replication.

Identification of the protein targets of RNS in the cytosolic

extract may be required to establish the mechanism of S-phase

arrest by RNS. One of the major mechanisms by which DNA

replication is controlled involves the regulated assembly of

prereplicative complexes (pre-RCs) at origins of replication.

Pre-RCs require sequential assembly of a number of proteins,

including ORC, Cdc6, and MCM proteins [45,46]. Cdc6 is also

required during S phase and has been shown to be degraded in

mammalian cells during apoptosis [47]. Although apoptosis

was not measured directly here, cells with a sub-G1 DNA

content were not observed by flow cytometry (Figs. 2 and 3).

Nonetheless, Cdc6 may represent one of the targets of RNS, for

PP2A has been shown to dephosphorylate Cdc6 by direct

physical interaction [48]. The phosphorylation status of RF-C

has also been reported to play a critical role in DNA elongation

[49].

Activation of the S-phase checkpoints is initiated in

response to DNA damage, though recent evidence suggests

that the checkpoint can be activated even in the absence of

DNA strand breaks [40]. The mechanisms by which eukaryotic

cells sense DNA strand breaks or stress signals remain to be

elucidated, but the rapid induction of ATM kinase activity after

ionizing radiation suggests it acts at an early stage of signal

transduction in mammalian cells [50]. In normal cells, ATM is

held inactive as a dimer or higher order multimer with the

kinase domain bound to a region surrounding serine 1981 that

is contained within the ‘‘FAT’’ domain. In response to a stress

signal, most ATM molecules within the cell get rapidly

phosphorylated on this site. This phosphorylation event does

not directly regulate the activity of kinase, but instead disrupts

ATM oligomers, which in turn allows accessibility of

substrates to the ATM kinase domain [40]. Changes in higher

order chromatin structure may represent an early checkpoint

signal [40]. Interestingly, the target amino acid for ATM

autophosphorylation has been shown to be a serine that is

regulated by PP2A [51], indicating that the role of serine–

threonine phosphatases is not limited to regulation of pRB

phosphorylation, but may also be to control early signaling

events in S-phase checkpoint activation. Alternatively, protein

phosphatases may facilitate cell cycle reentry by depho-

sphorylating Chk1 at a site phosphorylated by the ATR

homologue Rad3 in response to DNA damage, which results

in Chk1 inactivation and checkpoint release [52]. PP2A also

has been reported to regulate Chk2 activation, a key player of

checkpoint activation signaling pathway [53]. Further studies

will be required to clarify the effects of RNS on these and

other checkpoint mediators.

Our data indicate that RNS induce phosphorylation of

ATM and S-phase arrest in C10 cells in the absence of

detectable DNA strand breaks, suggesting that the primary

targets of RNS may be signaling proteins, chromatin

structure, or other factors involved in regulating ATM.

Serine–threonine phosphatases induced by RNS may repre-

sent one mechanism of arrest, suggesting that the phosphor-

ylation status of key mediators of S phase such as pRB and

Cdc6 may be involved. Mechanisms for checkpoint activation

that do involve extensive DNA damage may provide an

avenue for preventing cell cycle progression in response to

low levels of ROS and RNS. It is also possible that

deregulation of the redox status of the cell by RNS [29]

may contribute to induction of the DNA damage response

and thereby play a role not only in cell cycle arrest, but also in

neoplastic transformation.

Acknowledgments

We thank Bill Burhans for advice, adozelesin, and anti-

bodies; Y. Janssen-Heininger for technical assistance with the

NO2 exposures; and the laboratories of B. Mossman and A. van

der Vliet for fruitful discussions. This work was supported by a

grant from the NIH to N.H.H. (ES09673).

References

[1] MacNee, W.; Donaldson, K. Exacerbations of COPD: environmental

mechanisms. Chest 117:390S–397S; 2000.

[2] van der Vliet, A.; Eiserich, J. P.; Shigenaga, M. K.; Cross, C. E. Reactive

nitrogen species and tyrosine nitration in the respiratory tract: epiphe-

nomena or a pathobiologic mechanism of disease? Am. J. Respir. Crit.

Care Med. 160:1–9; 1999.

[3] Harris, J. B.; Chang, L. Y.; Crapo, J. D. Rat lung alveolar type I epithelial

cell injury and response to hyperoxia. Am. J. Respir. Cell Mol. Biol.

4:115–125; 1991.

[4] Miller, B. E.; Hook, G. E. Hypertrophy and hyperplasia of alveolar type

II cells in response to silica and other pulmonary toxicants. Environ.

Health Perspect. 85:15–23; 1990.

[5] Witschi, H. Responses of the lung to toxic injury. Environ. Health

Perspect. 85:5–13; 1990.

[6] Sadeghi-Hashjin, G.; Folkerts, G.; Henricks, P. A.; Verheyen, A. K.; van

der Linde, H. J.; van Ark, I.; Coene, A.; Nijkamp, F. P. Peroxynitrite

induces airway hyperresponsiveness in guinea pigs in vitro and in vivo.

Am. J. Respir. Crit. Care Med. 153:1697–1701; 1996.

[7] McConnell, R.; Berhane, K.; Gilliland, F.; London, S. J.; Vora, H.; Avol,

E.; Gauderman, W. J.; Margolis, H. G.; Lurmann, F.; Thomas, D. C.; et

al. Air pollution and bronchitic symptoms in Southern California

children with asthma. Environ. Health Perspect. 107:757–760; 1999.

[8] Shrivastava, P.; Pantano, C.; Watkin, R.; McElhinney, B.; Guala, A.;

Poynter, M. L.; Persinger, R. L.; Budd, R.; Janssen-Heininger, Y.

Reactive nitrogen species-induced cell death requires Fas-dependent

activation of c-Jun N-terminal kinase. Mol. Cell. Biol. 24:6763–6772;

2004.

[9] N’cho, M.; Brahmi, Z. Evidence that Fas-induced apoptosis leads to S

phase arrest. Hum. Immunol. 62:310–319; 2001.

[10] Ichinose, M.; Sugiura, H.; Yamagata, S.; Koarai, A.; Shirato, K.

Increase in reactive nitrogen species production in chronic obstructive


pulmonary disease airways. Am. J. Respir. Crit. Care Med. 162:

701–706; 2000.

[11] Abraham, R. T. Cell cycle checkpoint signaling through the ATM and

ATR kinases. Genes Dev. 15:2177–2196; 2001.

[12] Bartek, J.; Lukas, C.; Lukas, J. Checking on DNA damage in S phase.

Nat. Rev. Mol. Cell Biol. 5:792–804; 2004.

[13] Bartek, J.; Bartkova, J.; Lukas, J. The retinoblastoma protein pathway in

cell cycle control and cancer. Exp. Cell Res. 237:1–6; 1997.

[14] Weinberg, R. A. The retinoblastoma protein and cell cycle control. Cell

81:323–330; 1995.

[15] Harbour, J. W.; Dean, D. C. The Rb/E2F pathway: expanding roles and

emerging paradigms. Genes Dev. 14:2393–2409; 2000.

[16] Kristjansdottir, K.; Rudolph, J. Cdc25 phosphatases and cancer. Chem.

Biol. 11:1043–1051; 2004.

[17] Busino, L.; Chiesa, M.; Draetta, G. F.; Donzelli, M. Cdc25A

phosphatase: combinatorial phosphorylation, ubiquitylation and proteo-

lysis. Oncogene 23:2050–2056; 2004.

[18] Vigo, E.; Muller, H.; Prosperini, E.; Hateboer, G.; Cartwright, P.;

Moroni, M. C.; Helin, K. CDC25A phosphatase is a target of E2F

and is required for efficient E2F-induced S phase. Mol. Cell. Biol.

19:6379–6395; 1999.

[19] Fischer, P. M.; Endicott, J.; Meijer, L. Cyclin-dependent kinase

inhibitors. Prog. Cell Cycle Res. 5:235–248; 2003.

[20] Knudsen, K. E.; Booth, D.; Naderi, S.; Sever-Chroneos, Z.; Fribourg,

A. F.; Hunton, I. C.; Feramisco, J. R.; Wang, J. Y.; Knudsen, E. S.

RB-dependent S-phase response to DNA damage. Mol. Cell. Biol.

20:7751–7763; 2000.

[21] Kondo, T.; Higashi, H.; Nishizawa, H.; Ishikawa, S.; Ashizawa, S.;

Yamada, M.; Makita, Z.; Koike, T.; Hatakeyama, M. Involvement of

pRB-related p107 protein in the inhibition of S phase progression in

response to genotoxic stress. J. Biol. Chem. 276:17559–17567;

2001.

[22] Sever-Chroneos, Z.; Angus, S. P.; Fribourg, A. F.; Wan, H.; Todorov, I.;

Knudsen, K. E.; Knudsen, E. S. Retinoblastoma tumor suppressor

protein signals through inhibition of cyclin-dependent kinase 2 activity

to disrupt PCNA function in S phase. Mol. Cell. Biol. 21:4032–4045;

2001.

[23] Chew, Y. P.; Ellis, M.; Wilkie, S.; Mittnacht, S. pRB phosphorylation

mutants reveal role of pRB in regulating S phase completion by a

mechanism independent of E2F. Oncogene 17:2177–2186; 1998.

[24] Mittnacht, S. Control of pRB phosphorylation. Curr. Opin. Genet. Dev.

8:21–27; 1998.

[25] Yan, Y.; Mumby, M. C. Distinct roles for PP1 and PP2A in

phosphorylation of the retinoblastoma protein: PP2A regulates the

activities of G1 cyclin-dependent kinases. J. Biol. Chem. 274:

31917–31924; 1999.

[26] Lazzereschi, D.; Coppa, A.; Minicione, G.; Lavitrano, M.; Frago-

mele, F.; Colletta, G. The phosphatase inhibitor okadaic acid

stimulates the TSH-induced G1–S phase transition in thyroid cells.

Exp. Cell Res. 234:425–433; 1997.

[27] Nakamura, K.; Koda, T.; Kakinuma, M.; Matsuzawa, S.; Kitamura,

K.; Mizuno, Y.; Kikuchi, K. Cell cycle dependent gene expressions

and activities of protein phosphatases PP1 and PP2A in mouse

NIH3T3 fibroblasts. Biochem. Biophys. Res. Commun. 187:507–514;

1992.

[28] Berndt, N.; Ludlow, J. W. Interaction between the retinoblastoma protein

and protein phosphatase 1 during the cell cycle. Methods Mol. Biol.

281:17–32; 2004.

[29] Yuan, Z.; Schellekens, H.; Warner, L.; Janssen-Heininger, Y.; Burch, P.;

Heintz, N. H. Reactive nitrogen species block cell cycle re-entry through

sustained production of hydrogen peroxide. Am. J. Respir. Cell Mol.

Biol. 28:705–712; 2003.

[30] Burch, P. M.; Yuan, Z.; Loonen, A.; Heintz, N. H. An extracellular

signal-regulated kinase 1- and 2-dependent program of chromatin

trafficking of c-Fos and Fra-1 is required for cyclin D1 expression

during cell cycle reentry. Mol. Cell. Biol. 24:4696–4709; 2004.

[31] Ranjan, P.; Shrivastava, P.; Singh, S. M.; Sodhi, A.; Heintz, N. H.

Baculovirus P35 inhibits NO-induced apoptosis in activated macro-

phages by inhibiting cytochrome c release. J. Cell Sci. 117:3031–3039;

2004.

[32] Heintz, N. H.; Stillman, B. W. Nuclear DNA synthesis in vitro is

mediated via stable replication forks assembled in a temporally specific

fashion in vivo. Mol. Cell. Biol. 8:1923–1931; 1988.

[33] Giovannelli, L.; Cozzi, A.; Guarnieri, I.; Dolara, P.; Moroni, F. Comet

assay as a novel approach for studying DNA damage in focal cerebral

ischemia: differential effects of NMDA receptor antagonists and

poly(ADP-ribose) polymerase inhibitors. J. Cereb. Blood Flow Metab.

22:697–704; 2002.

[34] Agami, R.; Bernards, R. Distinct initiation and maintenance mechanisms

cooperate to induce G1 cell cycle arrest in response to DNA damage.

Cell 102:55–66; 2000.

[35] Mailand, N.; Falck, J.; Lukas, C.; Syljuasen, R. G.; Welcker, M.; Bartek,

J.; Lukas, J. Rapid destruction of human Cdc25A in response to DNA

damage. Science 288:1425–1429; 2000.

[36] Prives, C.; Hall, P. A. The p53 pathway. J. Pathol. 187:112–126; 1999.

[37] Esposito, F.; Russo, L.; Chirico, G.; Ammendola, R.; Russo, T.; Cimino,

F. Regulation of p21/waf1/cip1 expression by intracellular redox

conditions. IUBMB Life 52:67–70; 2001.

[38] Dou, Q. P.; An, B.; Will, P. L. Induction of a retinoblastoma phosphatase

activity by anticancer drugs accompanies p53-independent G1 arrest and

apoptosis. Proc. Natl. Acad. Sci. USA 92:9019–9023; 1995.

[39] Alberts, A. S.; Thorburn, A. M.; Shenolikar, S.; Mumby, M. C.;

Feramisco, J. R. Regulation of cell cycle progression and nuclear affinity

of the retinoblastoma protein by protein phosphatases. Proc. Natl. Acad.

Sci. USA 90:388–392; 1993.

[40] Bakkenist, C. J.; Kastan, M. B. DNA damage activates ATM through

intermolecular autophosphorylation and dimer dissociation. Nature

421:499–506; 2003.

[41] Helt, C. E.; Cliby, W. A.; Keng, P. C.; Bambara, R. A.; O’Reilly,

M. A. ATM and ATR exhibit selective target specificities in response

to different forms of DNA damage. J. Biol. Chem. 280:1186–1192;

2005.

[42] Bartkova, J.; Horejsi, Z.; Koed, K.; Kramer, A.; Tort, F.; Zieger, F.;

Guldberg, P.; Sehested, M.; Nesland, J. M.; Lukas, C.; Orntoft, T.;

Lukas, J.; Bartek, J. DNA damage response as a candidate anti-

cancer barrier in early human tumorigenesis. Nature 434:864–870;

2005.

[43] Gorgoulis, V. G.; Vassiliou, L.-V. F.; Karakaidos, P.; Zacharatos, P.;

Athanassios, K.; Liloglou, T.; Venere, M.; Ditullio, R. A., Jr.;

Kastrinakis, N. G.; Levy, B.; Kletsas, D.; Yoneta, A.; Herlyn, M.;

Kittas, C.; Halazonetis, T. D. Activation of the DNA damage

checkpoint and genomic instability in human precancerous lesions.

Nature 434:907–913; 2005.

[44] Schonthal, A. Okadaic acid—A valuable new tool for the study of signal

transduction and cell cycle regulation? New Biol. 4:16–21; 1992.

[45] Diffley, J. Regulation of early events in chromosome replication. Curr.

Biol. 14:778–786; 2004.

[46] Takeda, D. Y.; Dutta, A. DNA replication and progression through the S

phase. Oncogene 24:2827–2843; 2005.

[47] Blanchard, F.; Rusiniak, M. E.; Sharma, K.; Sun, X.; Todorov, I.;

Castellano, M. M.; Gutierrez, C.; Baumann, H.; Burhans, W. C.

Targeted destruction of DNA replication protein Cdc6 by cell death

pathways in mammals and yeast. Mol. Biol. Cell. 13:1536–1549;

2002.

[48] Yan, Z.; Fedorov, S. A.; Mumby, M. C.; Williams, R. S. PR48, a

novel regulatory subunit of protein phosphatase 2A, interacts with

Cdc6 and modulates DNA replication in human cells. Mol. Cell.

Biol. 20:1021–1029; 2000.

[49] Munshi, A.; Cannella, D.; Brickner, H.; Salles-Passador, I.; Podust, V.;

Fotedar, R.; Fotedar, A. Cell cycle-dependent phosphorylation of the

large subunit of replication factor C (RF-C) leads to its dissociation from

the RF-C complex. J. Biol. Chem. 278:48467–48473; 2003.

[50] Canman, C. E.; Lim, D. S.; Cimprich, K. A.; Taya, Y.; Tamai, K.;

Sakaguchi, K.; Appella, E.; Kastan, M. B.; Siliciano, J. D. Activation of

the ATM kinase by ionizing radiation and phosphorylation of p53.

Science 281:1677–1679; 1998.


[51] Goodarzi, A. A.; Jonnalagadda, J. C.; Douglas, P.; Young, D.; Ye, R.;

Moorhead, G. B.; Lees-Miller, S. P.; Khanna, K. K. Autophosphorylation

of ataxia– telangiectasia mutated is regulated by protein phosphatase 2A.

EMBO J. 23:4451–4461; 2004.

[52] Den Elzen, N. R.; O’Connell, M. J. Recovery from DNA damage

checkpoint arrest by PP1-mediated inhibition of Chk1. EMBO J.

23:908–918; 2004.

[53] Dozier, C.; Bonyadi, M.; Baricault, L.; Tonasso, L.; Darbon, J. M. Regu-

lation of Chk2 phosphorylation by interaction with protein phosphatase

2A via its BV regulatory subunit. Mol. Biol. Cell. 96:509–517; 2004.

s-phase arrest by reactive nitrogen species is bypassed by okadaic acid, an inhibitor of protein...

Documents