1 Vascular Biology Research Center and Division of Hematology, University of Texas-Houston Medical School, Houston, Texas 77030; and 2 Institute of Biomedical Sciences, Academia Sinica, Taipei 115, Taiwan
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ABSTRACT |
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Cyclooxygenase-2 (COX-2) is continuously expressed in most
cancerous cells where it appears to modulate cellular proliferation and
apoptosis. However, little is known about the contribution of
transient COX-2 induction to cell cycle progression or programmed cell
death in primary cells. In this study we determined whether COX-2
regulates proliferation or apoptosis in human fibroblasts. COX-2 mRNA, protein, and prostaglandin E2
(PGE2) were not detected in quiescent cells but were
expressed during the G0/G1 phase of the cell
cycle induced by serum. Inhibition of COX-2 did not alter G0/G1 to S phase transition or induce
apoptosis at concentrations that diminished PGE2.
Addition of interleukin-1 to serum enhanced COX-2 expression and
PGE2 synthesis over that by serum alone but had no effect
on the progression of these cells into S phase. Furthermore,
platelet-derived growth factor drove the G0 fibroblasts into the cell cycle without inducing detectable levels of COX-2 or
PGE2. Collectively, these data show that transient COX-2
expression in primary human fibroblasts does not influence cell cycle progression.
serum; platelet-derived growth factor; interleukin-1; proliferation; apoptosis; cyclooxygenase-2
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INTRODUCTION |
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CYCLOOXYGENASE (COX)
metabolizes arachidonic acid to prostaglandins (PGs) and thromboxane
(20) and exists in two isoforms (23).
Constitutively expressed COX-1 is present in most tissues where it
synthesizes PGs continuously to maintain physiological functions
(23, 29). In contrast, COX-2 is induced by proinflammatory stimuli, cytokines, and mitogens and synthesizes a large quantity of
PGs (8, 10, 12). COX-2 is crucial to the inflammatory response (4, 28), and nonsteroidal anti-inflammatory drugs (NSAIDs) selective for its inhibition are now used clinically to treat
inflammatory arthropathies. However, evidence is now emerging to show
that COX-2 is also involved in tumorigenesis. For instance, COX-2 is
constitutively expressed in various cancerous cells and tissues
(13, 24), while in APC716 mice, a model of
familial adenomatous polyposis bearing a COX-2 deletion, there was a
dramatic reduction in intestinal polyp size and number
(16). In the same model, mice treated with a selective COX-2 inhibitor had a reduced polyp formation (19).
Indeed, epidemiological studies have demonstrated a 50% reduction in
the rate of mortality from colorectal cancer in patients taking NSAIDs (6).
It has been shown that forced COX-2 overexpression arrests rat intestinal epithelial cells at G1 and confers resistance to apoptosis (5). COX-2 overexpression in ECV-304 cells is associated with a decrease in cells in S phase and an accumulation in G0/G1, while in COS-7 and human embryonic kidney 293 cells there is an increase in cells in G2/M (26). Additionally, H-ras-transfected rat intestinal epithelial cells constitutively overexpressing COX-2 traverse the cell cycle in the absence of serum and show significantly reduced proliferation after treatment with a selective COX-2 inhibitor (21). Collectively, these results suggest that continuous COX-2 expression causes phenotypic changes typical of cancerous cells by regulating, at least in part, cell cycle progression. However, despite the mounting evidence that pathological overexpression of COX-2 enhances proliferation and reduces apoptosis in cancerous or transformed cells, comparatively little is known about the role of transient COX-2 expression in primary cells. Therefore, in the present study we examined whether COX-2 regulates G0/G1 to S phase transition and programmed cell death in human foreskin fibroblasts (HFF), a well-characterized model for cell cycle studies. Our results show that COX-2 expression after serum stimulation was restricted to the G0/G1 phase of the cell cycle and that its inhibition did not alter G0/G1 to S transition or result in programmed cell death.
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MATERIALS AND METHODS |
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Cell culture. HFF were obtained from American Type Culture Collection (Manassas, VA) and cultured on 10-cm plates in DMEM supplemented with 10% fetal bovine serum (FBS), 100 µg/ml streptomycin, and 100 U/ml penicillin. For all experiments, cells were washed twice with PBS and incubated in FBS-free medium for 24 h. FBS-free medium was replaced with medium containing 10% FBS to initiate the cell cycle. All tissue culture reagents were from Life Technologies. To evaluate the effects of COX inhibitors on cell cycle progression, NS-398 (Calbiochem), indomethacin, aspirin, or sodium salicylate (Sigma) was added with 10% FBS to G0 cells in culture.
Cell cycle analysis. The cell cycle of the 24-h serum-starved HFF was initiated by adding 10% FBS to fresh medium. At indicated time points, cells were trypsinized, washed in ice-cold PBS, and fixed using ice-cold acetone-free methanol (methanol:PBS ratio was 2:1). Cellular DNA was stained with 0.01% propidium iodide (Calbiochem) in PBS containing 0.1% Triton X-100 and 0.037% EDTA followed by the addition of 100 U/ml RNase (Worthington Biochemical) to remove RNA. Samples were filtered through a 35-µm nylon mesh, and the percentage of cells in sub-G0/G1, G0/G1, S, and G2/M phase was assessed using flow-assisted cell sorting (FACS) analysis (Becton Dickinson). Data were analyzed using ModFit software (Verity software).
Northern blotting. RNA was isolated from cultured cells by using RNA-STAT 60 (TEL-TEST; Friendswood, TX). RNA (25-30 µg) was fractionated on 1% agarose and was transferred to a positively charged nylon membrane. As a COX-2 probe, agarose gel-purified, full-length, 1.9-kb COX-2 cDNA was used (31). Hybridization and detection by autoradiography were performed according to a procedure previously reported (31).
Western blotting. Cell pellets were lysed with PBS (pH 7.4) containing 0.1% Triton X-100, 0.01% EDTA, 1 mM phenylmethylsulfonyl fluoride, 1.5 mM pepstatin A, and 0.2 mM leupeptin. Lysates were centrifuged at 13,000 rpm for 10 min. The supernatants were boiled for 5 min with equal volumes of 2× gel-loading buffer (100 mM Tris, 10% mercaptoethanol, 20% glycerol, 4% SDS, 2 mg/ml bromphenyl blue). The protein concentration of the supernatant was determined by the bicinchoninic acid assay method (Pierce Chemical). Protein (5 µg) was applied to and separated on 10% SDS-polyacrylamide minigels (Hoefer Scientific Instruments) using the Laemmli buffer system and transferred to polyvinylidine difluoride membranes (Amersham Pharmacia Biotech). Nonspecific IgGs were blocked with 5% nonfat dried milk containing 1 mg/ml globulin-free bovine serum albumin and incubated with specific antibodies to COX-1 and COX-2 (both 1:1,000; Santa Cruz Biotechnology). Protein bands were detected using enhanced chemiluminescence (Amersham Pharmacia Biotech).
COX activity. Because PGE2 is the major metabolite of COX enzyme catalysis in fibroblasts, we measured its levels in the culture medium by an enzyme immunoassay detection kit (Amersham Pharmacia Biotech).
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RESULTS |
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COX-2 and COX-1 expression during serum-driven cell cycle
progression.
We have recently shown by flow cytometry that, after FBS deprivation
for 24 h, 90% HFF were in the G0/G1 phase
of the cell cycle, which entered into S phase 16 h after the
addition of 10% FBS, and, by 24 h, >50% of cells were in S
phase (7). These results are consistent with previous
reports that the vast majority of HFF cultured in medium deprived of
serum are quiescent (G0) (17). COX-2 mRNA was
undetectable in quiescent cells, and, after addition of FBS, low levels
were measurable at 1 and 2 h, and the level peaked at 4 h
(Fig. 1A). COX-2 mRNA levels
declined thereafter, becoming barely detectable at 12 h.
The COX-2 protein was also expressed in a time-dependent manner after
serum addition. It peaked at 6-8 h after serum treatment and
declined thereafter (Fig. 1B). In contrast, COX-1 protein
was constitutively expressed in quiescent cells, increased slightly
after the addition of FBS, peaked at 12 h, and remained at the
same level up to 32 h (Fig. 1C). Thus COX-1 is involved
in a housekeeping function during cell cycle progression, while COX-2
expression, which coincides with G1 phase of the cell
cycle, could play a role in cell cycle regulation.
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Effect of COX inhibitors on cell cycle progression and
apoptosis.
Expression of COX-2 and PGE2 synthesis in the
G0/G1 phase of the cell cycle implicates a role
for COX-2 in regulating G0/G1 to S transition.
To discern this, we treated quiescent cells with NS-398 at the same
time as FBS stimulation. Despite an almost complete obliteration of
PGE2 synthesis by NS-398 (Fig. 2A), proportions of cells that progressed through G1 and entered S phase
were unaltered by this selective COX-2 inhibitor compared with the
controls (Fig. 3A). Because
indomethacin was previously reported to arrest cells in G1
(1), we also treated quiescent cells with this dual COX inhibitor in the presence of FBS but observed no change in the percentages of cells in G0/G1 compared with
controls (Fig. 3B). Western blot analysis revealed that
neither NS-398 nor indomethacin inhibited COX-2 protein expression
(data not shown). We also evaluated the effect of aspirin, which was
shown previously to inhibit COX-2 mRNA and protein levels in these
cells (31), on cell cycle progression. Aspirin
pretreatment had no significant effect on altering percentages of cells
entering the S phase. Similarly, sodium salicylate (104
M) did not change cell cycle progression (data not shown).
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Effects of interleukin-1 on COX-2 induction, PGE2
synthesis, and cell cycle progression induced by serum.
To examine further the role of COX-2 in fibroblast proliferation, we
treated quiescent HFF with a combination of serum and the
proinflammatory cytokine interleukin-1
(IL-1
; 1 nM), a well-known inducer of COX-2 and PGE2 synthesis. It was found that,
while this combination of stimuli caused a sustained increase in COX-2 protein expression (Fig. 5A)
and PGE2 synthesis (Fig. 5B) over and above that
of serum alone during G0/G1, there was no
corresponding alteration in cell cycle progression into S phase (Fig.
5C).
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Platelet-derived growth factor drove the cell cycle without
inducing COX-2 expression.
Platelet-derived growth factor (PDGF) is known to drive quiescent cells
into the cell cycle. We determined whether PDGF-driven cell cycle
progression was similarly accompanied by COX-2 expression as serum. To
our surprise, despite a similar time course of cell cycle progression
as serum (Fig. 6A), PDGF at a
submaximal concentration (1 ng/ml) did not induce COX-2 protein
expression during the 24-h period (Fig. 6B) or increase
PGE2 synthesis (Fig. 6C), in contrast to the
reported data of COX-2 induction by PDGF at a higher concentration (10 ng/ml) (30). Our results with submaximal concentrations of
PDGF further support the notion that COX-2 expression is not essential
for cell cycle progression.
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DISCUSSION |
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In this study, we determined COX-2 expression during
synchronized cell cycle progression by using a fibroblast model that has been widely used to characterize the cellular and molecular events
and control mechanisms of the cell cycle (17). We used FBS
and PDGF to drive quiescent cells into the cell cycle, and the results
show a striking difference in COX-2 expression. COX-2 is expressed in
the G1 phase of FBS-driven cells, whereas, in PDGF-driven
cell cycle progression, COX-2 protein was undetectable up to 32 h
after PDGF treatment. These results suggest that COX-2 expression is
not an integral part of cell cycle progression but rather is induced in
G0 cells by FBS, which is coincidental with the
G0 phase of the cell cycle. It is possible that cell cycle progression and COX-2 expression are induced by different factors in
FBS. IL-1 greatly increases COX-2 expression and PGE2
production induced by serum but has no effect on serum-driven cell
cycle progression, despite the production of large quantities of
PGE2. We have recently shown that addition of IL-1
alone
without FBS to quiescent fibroblasts does not drive the quiescent cells
into the cell cycle, and yet it induces COX-2 expression
(7). These results support the notion that cells in
G0 phase are highly responsive to exogenous stimuli in
COX-2 expression. Taken together, these results suggest that COX-2
expression is not inherently tied to cell cycle progression but is an
important property of G0 cells in response to exogenous
stimuli including serum, IL-1
, and phorbol esters.
Our results further suggest that COX-2 expression is not essential for
cell cycle progression. Selective inhibition of COX-2 with NS-398
obliterates almost the entire PGE2 synthesis without changing the duration of G1 and the proportion of cells
that enter the S phase of the cell cycle. Aspirin, which inhibits
COX-1/COX-2 activities and suppresses COX-2 expression
(31), also has no effect on the cell cycle. This is
further supported by cell cycle progression induced by PDGF despite a
lack of COX-2 induction or PGE2 synthesis. Thus our results
are in striking contrast to those from persistent transfection of
COX-2, which causes cell cycle arrest. One possible explanation for the
difference is that persistent COX-2 overexpression during the S and
G2/M phases of the cell cycle causes DNA damage, which
signals cell cycle arrest. It has been reported that proliferative
cells are more susceptible to DNA damage and mutation (2).
COX-2 overexpression is accompanied by generation of metabolites,
including malondialdehyde, which causes oxidative stress to DNA
(18), and inhibition of COX-2 by nebutame in asynchronized
cells, which reduces the level of DNA oxidation (25). We
have recently observed that that COX-2 expression in response to
phorbol 12-myristate 13-acetate and IL-1 is subdued in proliferative
fibroblasts compared with quiescent cells (7). These
results suggest that COX-2 expression is controlled in a cell
cycle-dependent manner in normal cells. Under the control mechanism,
COX-2 is transiently expressed predominantly in cells at quiescent or
early G1 of the cell cycle. Work is in progress to
elucidate the control mechanism.
It was reported that indomethacin induces G1 arrest and
causes apoptosis in both COX-2-positive and COX-2-negative cell
lines (22), as well as induces apoptosis in
transformed murine fibroblasts in the absence of both COX isoforms
(32). Indomethacin has been shown to bind and activate
peroxisome proliferator-activated receptor- (PPAR
)
(14), a nuclear receptor that, when activated in human colorectal cancer cell lines, causes G1 arrest and
apoptosis (3). However, in the present study,
indomethacin had no effect on either cell cycle progression or
apoptosis in nontransformed primary HFF. It may be speculated
that activation of the PPAR
is also without effect on cellular
proliferation and programmed cell death in HFF. Thus there are striking
differences in response to indomethacin between normal and transformed
cells. These differences may reflect derangements of the endogenous
control of COX-2-dependent and COX-2-independent cell cycle arrest,
apoptosis resistance, and cell proliferation in transformed and
cancer cells, in contrast to a tight control of COX-2 expression in
normal cells. Transient expression of COX-2 during
G0/G1 phase of the cell cycle by serum is
likely to play a physiological role such as wound healing. It has been
shown by microarray that several genes including COX-2, transforming
growth factor-
(TGF-
), and metalloproteinase-1 (MMP-1) are
expressed in mid G0/G1 in serum-stimulated
quiescent human dermal diploid fibroblasts (9). COX-2 and
PGE2 have been reported to modulate the expression of
TGF-
(11) and MMP-1 (15) in asynchronous
cells. However, we did not observe such modulation in our synchronized
fibroblast model, since NS-398 at doses reducing PGE2
synthesis by >95% had no effect on TGF-
or MMP-1 levels stimulated
by serum (data not shown). Our results suggest that serum stimulates
quiescent cells to express COX-2 and several other genes that act in
concert to promote wound healing and other pathophysiological processes.
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ACKNOWLEDGEMENTS |
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We thank Dr. Jeffrey J. Yen for assistance in cell cycle analysis and in the preparation of this manuscript.
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FOOTNOTES |
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This work was supported by National Institutes of Health Grants P50 NS-23327 and R01 HL-50675 (to K. K. Wu) and by Taiwan Academia Sinica.
Address for reprint requests and other correspondence: K. K. Wu, Vascular Biology Research Center and Division of Hematology, Univ. of Texas-Houston Medical School, 6431 Fannin St., MSB 5.016, Houston, TX 77030 (E-mail: Kenneth.K.Wu{at}uth.tmc.edu).
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.
Received 27 November 2000; accepted in final form 21 February 2001.
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