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Effects of antirheumatic treatments on the prostaglandin E2 biosynthetic pathway.

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ARTHRITIS & RHEUMATISM
Vol. 52, No. 11, November 2005, pp 3439–3447
DOI 10.1002/art.21390
© 2005, American College of Rheumatology
Effects of Antirheumatic Treatments on the
Prostaglandin E2 Biosynthetic Pathway
Marina Korotkova,1 Marie Westman,1 Karina R. Gheorghe,2 Erik af Klint,1 Christina Trollmo,1
Ann Kristin Ulfgren,1 Lars Klareskog,1 and Per-Johan Jakobsson1
Objective. Microsomal prostaglandin E synthase
1 (mPGES-1) is up-regulated in experimental arthritis
and markedly expressed in synovial tissue biopsy samples from patients with rheumatoid arthritis (RA). This
study was carried out to determine the effects of tumor
necrosis factor (TNF) blockers and glucocorticoids on
mPGES-1 and cyclooxygenase (COX) expression, as
well as biosynthesis of PGE2 in rheumatoid joints.
Methods. In vitro effects of TNF blockers and
dexamethasone on the PGE2 biosynthetic pathway were
examined in RA synovial fluid mononuclear cells
(SFMCs) by flow cytometry. PGE2 levels in culture
supernatants were measured by enzyme immunoassay.
Expression of enzymes responsible for PGE2 synthesis
ex vivo was evaluated by immunohistochemistry in
synovial biopsy samples obtained from 18 patients
before and after treatment with TNF blockers and from
16 patients before and after intraarticular treatment
with glucocorticoids. Double immunofluorescence was
performed using antibodies against mPGES-1, COX-1,
COX-2, and CD163.
Results. Double immunofluorescence revealed
that mPGES-1 and COX-2 were colocalized in SFMCs
as well as in RA synovial tissue cells. The addition of
either TNF blockers or dexamethasone suppressed
lipopolysaccharide-induced mPGES-1 and COX-2 expression in synovial fluid monocyte/macrophages in
vitro and decreased the production of PGE2. Intraarticular treatment with glucocorticoids significantly reduced both mPGES-1 and COX-2 expression in arthritic synovial tissue ex vivo. The number of COX-1–
expressing cells in synovial tissue was also significantly
decreased by glucocorticoid treatment. In contrast, neither mPGES-1 nor COX-2 expression in synovial tissue
was significantly suppressed by anti-TNF therapy.
Conclusion. These data are the first to demonstrate the effects of antirheumatic treatments on
mPGES-1 expression in RA and suggest that the inhibition of PGE2 biosynthesis, preferably by targeting
mPGES-1, might complement anti-TNF treatment for
optimal antiinflammatory results in RA.
Rheumatoid arthritis (RA) is a severe, chronic
disease characterized by systemic and local inflammation leading to joint destruction and functional impairment. The proinflammatory cytokines tumor necrosis
factor ␣ (TNF␣) and interleukin-1␤ (IL-1␤) play key
roles in initiating and driving RA. These cytokines
induce the production of prostaglandin E2 (PGE2),
which contributes to several of the pathologic features of
RA, such as pain, inflammation, and bone destruction
(1–3).
In the PGE2 biosynthetic pathway, cyclooxygenase 1 (COX-1) and COX-2 both catalyze the conversion
of arachidonic acid into PGH2. Subsequently, terminal
PGE synthases catalyze the formation of PGE2 from
PGH2. Several PGE synthases have been cloned and
characterized, including microsomal PGE synthase 1
(mPGES-1) (4,5), mPGES-2 (6), and cytosolic PGES
(cPGES) (7). Importantly, mPGES-1 is strongly induced
by proinflammatory stimuli and is functionally linked to
Supported by grants from the King Gustaf V 80 Years
Foundation, Alex and Eva Wallströms Foundation, Pfizer Inc., the
Swedish Society of Medicine, Freemason Lodge “Barnhuset” in Stockholm, Börje Dahlin Foundation, the Swedish Medical Research Council, and the Swedish Rheumatism Association.
1
Marina Korotkova, MD, PhD, Marie Westman, BSc, Erik af
Klint, MD, Christina Trollmo, PhD, Ann Kristen Ulfgren, PhD, Lars
Klareskog, MD, PhD, Per-Johan Jakobsson, MD, PhD: Karolinska
Institutet, Karolinska University Hospital, Stockholm, Sweden; 2Karina R. Gheorghe, MD: Karolinska Institutet, Stockholm, and Karolinska Institutet, Novum, Huddinge, Sweden.
Dr. Klareskog has obtained research grants from and worked
as a scientific advisor for Wyeth, Schering-Plough, and Abbott.
Address correspondence and reprint requests to Marina
Korotkova, MD, PhD, Rheumatology Research Laboratory, CMM
L8-04, Karolinska University Hospital, Stockholm S-17176, Sweden.
E-mail: Marina.Korotkova@cmm.ki.se.
Submitted for publication February 9, 2005; accepted in
revised form July 27, 2005.
3439
3440
COX-2 in various cells (8–10). It has also been reported
to function with COX-1 (9,11). In contrast, mPGES-2
and cPGES are constitutively expressed and therefore
are likely to be active during basal PGE2 production
(6,7). Although cPGES has been predominantly linked
to COX-1 (7), mPGES-2 efficiently couples with both
COX-1 and COX-2 (6).
Recent studies in rodents have provided convincing evidence for the important role of mPGES-1 in the
pathogenesis of inflammatory arthritis (12–14). Furthermore, the induction of both mPGES-1 and COX-2 was
observed in primary rheumatoid synovial cells after
treatment with proinflammatory cytokines in vitro (10).
We have recently demonstrated significant expression of
mPGES-1 in synovial tissue biopsy samples from patients with RA (15). Since mPGES-1 represents the
downstream and rate-limiting enzyme in the induced
state of PGE2 biosynthesis, it may be a therapeutic target
in RA.
Conventional treatment of RA includes nonsteroidal antiinflammatory drugs (NSAIDs), glucocorticoids, and disease-modifying antirheumatic drugs
(DMARDs). NSAIDs and glucocorticoids are well
known to suppress PGE2 generation. NSAIDs specifically inhibit COX activity (16), and treatment with COX
inhibitors effectively relieves pain and inflammation.
However, these drugs cause serious adverse effects, such
as gastrointestinal toxicity (17). Selective inhibition of
COX-2 has fewer gastrointestinal side effects, but may
lead to increased risk of thrombosis and cardiovascular
disease (18,19). Treatment with glucocorticoids efficiently suppresses inflammation in RA, but is also
associated with a number of adverse reactions (20).
One of the antiinflammatory effects of glucocorticoids is associated with suppression of PGE2 biosynthesis by the inhibition of phospholipase A2 activity (21)
and down-regulation of COX-2 expression (22). Marked
suppression of COX-2 expression by dexamethasone was
observed in freshly explanted RA synovial tissues and
cultured synoviocytes (22). Recently, the suppressive
effects of dexamethasone on mPGES-1 messenger
RNA, protein expression, and enzyme activity in synovial fibroblasts have been demonstrated in vitro (10).
Whether mPGES-1 might be an additional target for
glucocorticoid antiinflammatory action in RA patients
has not been reported.
New biologic DMARDs, blockers of TNF, display high clinical efficacy and retard joint damage in
patients with RA (23), although in most cases this
treatment does not induce complete remission. AntiTNF therapy is based on TNF binding and suppression
KOROTKOVA ET AL
of inflammation by inhibiting its downstream effects.
Anti-TNF therapy down-regulates the cytokine/
chemokine cascade (24–26), reduces inflammatory cell
migration into the RA joint (26), and decreases serum
matrix metalloproteinase levels (27,28). However,
whether anti-TNF therapy can have an effect on the
PGE2 biosynthetic pathway has not been investigated to
date.
In the present study, we examined the effects of
TNF blockers (infliximab and etanercept) and glucocorticoids on the prostaglandin E2 biosynthetic pathway.
We used both an in vitro system to study the expression
of mPGES-1 and related enzymes in synovial fluid
mononuclear cells (SFMCs) and an in vivo system in
which we analyzed synovial tissue biopsy samples from
RA patients obtained before and after therapy with TNF
blockers or glucocorticoids.
PATIENTS AND METHODS
Patients and tissue samples. Eighteen patients who
met the American College of Rheumatology (formerly, the
American Rheumatism Association) diagnostic criteria for RA
were recruited into the study (29). At study entry, all patients
had a moderate to high level of disease activity (Disease
Activity Score ⬎4.8) (30). In the first group, 8 patients (7
women and 1 man, median age 44 years, range 35–59 years)
received a subcutaneous injection of 25 mg etanercept (Wyeth
Europa, Maidenhead, UK) twice a week. In 3 patients, etanercept was combined with prednisolone, in 4 patients etanercept was combined with DMARDs and prednisolone, and in 1
patient etanercept was combined with DMARDs alone. All
patients except 1 received NSAIDs. None of these 8 patients
received intraarticular steroid therapy. Synovial tissue biopsy
specimens were obtained during arthroscopy from patients
before and a median of 8 weeks (range 7–10 weeks) after
initiation of treatment with etanercept.
In the second group, 10 patients (7 women and 3 men,
median age 55 years, range 25–74 years) received intravenous
infusions of infliximab (Schering-Plough, Stockholm, Sweden)
at a dose of 3 mg/kg at 0, 2, and 6 weeks. All patients received
infliximab in combination with DMARDs, and all patients
except 1 were treated with NSAIDs. Five patients also received
prednisolone. None of these 10 patients received intraarticular
steroid therapy. Synovial tissue biopsy specimens were obtained during arthroscopy from patients before and a median
of 10 weeks (range 8–16 weeks) after initiation of treatment
with infliximab.
Sixteen patients with inflammatory arthritis (8 women
and 8 men, median age 41 years, range 20–73 years) received
intraarticular injections of 40 mg triamcinolone hexacetonide
(Lederspan; Wyeth Lederle, Solna, Sweden). Patients with RA
(7 patients) and other inflammatory arthritides (3 patients with
monarthritis, 2 with juvenile idiopathic arthritis, 1 with psoriatic arthritis, 1 with spondylarthritis, 1 with polyarthritis, and
1 with oligoarthritis) showed clinical signs of active arthritis,
including swelling and pain in the joint, which were reduced in
mPGES-1 AND ANTIRHEUMATIC TREATMENT
all patients after intraarticular corticosteroid treatment. Nine
patients received intraarticular corticosteroids alone, 3 patients received intraarticular corticosteroids combined with
DMARDs, 1 patient received intraarticular corticosteroids
combined with DMARDs and prednisolone, and 3 patients
received intraarticular corticosteroids combined with prednisolone. All patients except 3 received NSAIDs. None of
these 16 patients received TNF blockers. Synovial biopsy
specimens were obtained during arthroscopy before and 9–12
days after the intraarticular injection.
Synovial fluid was collected by aspiration from an
additional 8 patients with active RA (4 women and 4 men,
median age 55 years, range 38–65 years). All patients received
DMARDs, 3 in combination with TNF blockers and prednisolone, and 1 in combination with TNF blockers. All patients
except 2 received NSAIDs.
This study was approved by the Ethics Committee at
the Karolinska University Hospital, Solna, Stockholm, Sweden.
Cell isolation. SFMCs were isolated using a discontinuous density gradient (Ficoll-Paque; Pharmacia, Uppsala,
Sweden). Cells were cultured in RPMI 1640 medium supplemented with 100 units/ml penicillin–streptomycin, 2 mM glutamine, 10 mM HEPES (all from Gibco Invitrogen, Lidingo,
Sweden), and 5% human pooled serum (blood bank, Karolinska Hospital, Solna, Stockholm, Sweden) in the presence of
100 ng/ml lipopolysaccharide (LPS; Sigma, St. Louis, MO) at
37°C in a humidified atmosphere containing 5% CO2, for 42
hours. Where indicated, the cultures were treated using
10⫺6–10⫺9M dexamethasone (Sigma), 0.1–100 ␮g/ml infliximab, or 0.1–100 ␮g/ml etanercept. Preliminary experiments
have shown that these treatments affect mPGES-1 expression
in a dose-dependent manner, and maximal effects were observed at concentrations of 10⫺6M dexamethasone, 100 ␮g/ml
etanercept, and 100 ␮g/ml infliximab. Unstimulated control
cells were always cultured in parallel. Culture supernatants
were harvested and stored at ⫺70°C. For immunostaining,
SFMCs were cultured in chamber slides (Nalge Nunc International, Naperville, IL) in the presence of 100 ng/ml LPS, fixed
with 2% formaldehyde (Sigma) for 20 minutes, and stored at
⫺70°C.
Flow cytometric analysis. Staining of SFMCs for flow
cytometry was performed after blocking of nonspecific binding
with phosphate buffered saline (PBS) supplemented with 5%
human serum. Cells were incubated with peridin chlorophyll
protein–conjugated mouse monoclonal anti-CD14 antibody
(Becton Dickinson, San Jose, CA) for 20 minutes at 4°C,
washed with PBS, and fixed with 4% paraformaldehyde
(Sigma) for 10 minutes. For intracellular staining, cells were
permeabilized using PBS supplemented with 0.1% saponin
(Reidel de Haen, Seelze, Germany) and 1% bovine serum
albumin (Sigma) for 10 minutes. For intracellular staining of
COX-1 and COX-2, cells were incubated with mouse monoclonal fluorescein isothiocyanate–conjugated anti–COX-1 and
phycoerythrin-conjugated anti–COX-2 antibodies (Becton
Dickinson) or isotype-matched irrelevant antibodies, respectively, for 45 minutes. For intracellular staining of mPGES-1,
cells were incubated with rabbit polyclonal antiserum raised
against purified human mPGES-1 (15) or isotype-matched
irrelevant control for 45 minutes, then with allophycocyaninconjugated goat anti-rabbit IgG (heavy and light chain;
3441
Molecular Probes, Eugene, OR) for 30 minutes. Analyses
were performed using a FACSCalibur (Becton Dickinson) and
CellQuest software (Becton Dickinson). Scatter properties
were used to identify the monocyte population. Quadrants
were set on the respective isotype controls. Results are expressed as a percentage of the total number of gated monocytes expressing CD14 and producing the respective enzymes.
Measurement of PGE2. The ability of SFMCs to
produce PGE2 was assessed in the supernatants of control and
LPS-stimulated cells cultured for 42 hours with or without
inhibitors. In addition, the conversion of exogenous arachidonic acid (Nu-Check Prep, Elysian, MN) to PGE2 by SFMCs,
cultured under the conditions described above, was assessed in
4 patients. For this purpose, cultured SFMCs were harvested,
washed twice in calcium-free PBS, assessed for viability, and
counted by trypan blue dye exclusion. Cells were then resuspended in PBS supplemented with Ca2⫹ and Mg2⫹. Arachidonic acid was added to a final concentration of 20 ␮M and
cells were stimulated with 2 ␮M calcium ionophore A23187
(Calbiochem-Novabiochem, San Diego, CA) for 30 minutes at
37°C. The incubation was terminated by adding 1M HCl to
attain a pH of 3. After centrifugation, the supernatants were
collected and stored at ⫺80°C until analyzed.
In a preliminary experiment, the supernatants were
analyzed using a reverse-phase high-performance liquid chromatography (RP-HPLC) system with ultraviolet detection at
195 nm and online radioactivity detection using a Radiomatic
Flo-One beta detector (Packard, Downers Grove, IL). The
mobile phase was water, acetonitrile, and trifluoroacetic acid
(70:30:0.007, by volume), and the eicosanoid compounds were
separated using a Nova-Pak C-18 column (Waters, Milford,
MA). Prostaglandin products were identified by comparison
with the retention time of synthetic standards (Cayman Chemical, Ann Arbor, MI). Aliquots of the supernatants were
measured using a PGE2 enzyme immunoassay (EIA) kit
(Cayman Chemical). PGE2 concentrations were assayed in
duplicate or triplicate and were read against a standard curve.
The results are expressed in ng per 1 ⫻ 106 cells.
Immunohistochemical analysis. Tissue samples were
snap frozen in prechilled isopentane and stored at ⫺70°C until
sectioned. Serial cryostat sections (8 ␮m) were fixed with 2%
formaldehyde (Sigma) for 20 minutes and stored at ⫺70°C.
Sections were incubated with rabbit polyclonal antiserum
raised against purified human mPGES-1 (15) or with the
following antibodies (all purchased from Cayman Chemical):
rabbit polyclonal anti-cPGES, rabbit polyclonal anti–COX-1,
and mouse monoclonal anti–COX-2. The staining procedure
has been described previously (31). Negative control experiments were performed using isotype-matched irrelevant antibodies and omitting the primary antibodies. Staining in synovial tissue was completely abolished by preincubation of anti–
mPGES-1 serum with mPGES-1 protein and preincubation of
commercial antibodies with respective blocking peptides (Cayman Chemical), which confirmed the specificity of the staining.
Stained sections were examined using a Polyvar II
microscope (Reichert-Jung, Vienna, Austria) and photographed with a digital camera (300F; Leica, Cambridge, UK).
Positive staining was indicated as brown deposits. The positive
staining was assessed quantitatively using computer-assisted
image analysis and expressed as percentages of the total area
of counterstained tissue. Double immunofluorescence staining
3442
KOROTKOVA ET AL
was performed using anti-human mPGES-1 antiserum and
mouse monoclonal anti-human COX-2 (Cayman Chemical),
anti-human COX-1 (Wako, Neuss, Germany), or anti-human
CD163 (BerMac3 clone; Dako, Glostrup, Denmark) antibodies. The staining procedure has been described previously (15).
Briefly, after blocking with an avidin–biotin kit (Vector, Peterborough, UK), the sections were incubated overnight with
primary antibodies. Thereafter, the sections were incubated
with biotinylated goat anti-rabbit IgG (heavy and light chain;
Vector), followed by incubation with streptavidin-conjugated
fluorophore Alexa Fluor 488 (Molecular Probes, Leiden, The
Netherlands). After blocking using the avidin–biotin kit, the
sections were incubated with biotinylated horse anti-mouse
IgG (heavy and light chain; Vector), followed by incubation
with streptavidin-conjugated fluorophore Alexa Fluor 546
(Molecular Probes). Some sections were also incubated with
4⬘,6-diamidino-2-phenylindole (KPL, Gaithersburg, MD) for 5
minutes.
Statistical analysis. Data were analyzed by Friedman’s
test for repeated measures, followed by Wilcoxon’s signed rank
test and Bonferroni correction for multiple comparisons. P
values less than 0.05 were considered significant.
RESULTS
Induction of mPGES-1 in SFMCs. SFMCs were
isolated and cultured in chamber slides in the presence
of LPS. LPS-induced expression of mPGES-1 was detected in SFMCs with macrophage-like morphology
(Figure 1A). Double immunofluorescence revealed the
expression of mPGES-1 in cells expressing CD163, a
member of the scavenger receptor family, which is highly
specific for cells of the mononuclear phagocyte lineage
(Figure 1B). In CD163⫹ SFMCs, no expression of
mPGES-1 was evident (Figure 1C).
Expression of mPGES-1 in synovial fluid monocytes was also evaluated by flow cytometric analysis
(Figure 2). In monocytes that had not been exposed to
LPS, mPGES staining was detected at a low level, while
stimulation with LPS substantially increased mPGES-1
expression in CD14⫹ monocytes (Figure 2A). LPSinduced mPGES-1 expression in CD14⫹ monocytes was
almost completely suppressed by dexamethasone. In
addition, both infliximab and etanercept reduced the
percentage of CD14⫹ monocytes that expressed
mPGES-1 (Figure 2A). In order to clarify a functional
coupling between mPGES-1 and COX-1 or COX-2, we
analyzed the expression of these enzymes in synovial
fluid monocytes. Figure 2B shows coordinated upregulation of mPGES-1 and COX-2 in CD14⫹ monocytes by LPS and suppression by dexamethasone. In
addition, the percentage of CD14⫹ monocytes stained
for mPGES-1, as well as for COX-2, was similarly
decreased after treatment with infliximab or etanercept.
Figure 1. Induction of microsomal prostaglandin E synthase 1
(mPGES-1) in synovial fluid mononuclear cells (SFMCs) by lipopolysaccharide. A, Green immunofluorescence staining of mPGES-1–
positive SFMCs (original magnification ⫻ 250). B and C, Double
fluorescence staining, showing mPGES-1–positive (green), CD163⫹
(red), and double-stained (yellow) SFMCs (4⬘,6-diamidino-2phenylindole–counterstained, original magnification ⫻ 500). D, Double fluorescence staining, showing mPGES-1–positive (green), cyclooxygenase 2 (COX-2)–positive (red), and double-stained (yellow)
SFMCs (original magnification ⫻ 500).
In contrast, the percentage of CD14⫹ monocytes that
expressed COX-1 was not increased by LPS stimulation
or suppressed by treatment with dexamethasone or TNF
blockers. Double immunofluorescence confirmed coexpression of mPGES-1 and COX-2 in SFMCs stimulated
with LPS (Figure 1D).
PGE2 generation by SFMCs. As demonstrated,
42 hours after LPS treatment both mPGES-1 and
COX-2 were induced in SFMCs. PGE2 accumulation in
the supernatant at this point was enhanced in LPStreated cells (mean ⫾ SEM 8.6 ⫾ 2.7 ng/106 cells) (n ⫽
8 patients) compared with control cells, which mainly
expressed COX-1 (mean ⫾ SEM 0.7 ⫾ 0.2 ng/106 cells).
Treatment with dexamethasone or TNF blockers prevented the accumulation of PGE2 (Figure 3A). In comparison, Figure 3B shows the results after incubation of
these cells, treated as above, with exogenous arachidonic
acid and calcium ionophore for 30 minutes. In this short
time frame, the pattern of PGE2 release was similar to
that for endogenous production and accumulation of
PGE2 in culture medium after 42 hours (Figure 3A).
Metabolic profiling was also performed by incubation of
SFMCs with exogenous 14C-labeled arachidonic acid,
followed by RP-HPLC analyses with online radioactivity
detection. In LPS-treated cells, the predominant radio-
mPGES-1 AND ANTIRHEUMATIC TREATMENT
3443
Figure 2. Coordinate expression of mPGES-1 and COX-2 in the monocyte/macrophage population of SFMCs in patients with rheumatoid arthritis. A, Flow cytometric analysis revealed the
expression of mPGES-1 in CD14⫹ monocytes a, without stimulation and b–f, after stimulation
with lipopolysaccharide (LPS) and treatment with dexamethasone (c), infliximab (d), or etanercept (e), compared with irrelevant antibody control (f). Results are expressed as the percentage of
the total number of gated monocytes expressing CD14 and producing mPGES-1. B, Flow
cytometric analysis results indicating the effects of treatment with dexamethasone (Dex),
infliximab (Inf), and etanercept (Eta) on enzyme expression in the LPS-induced monocyte/
macrophage population of SFMCs. Results are expressed as the mean and SEM percentage of the
total number of gated monocytes expressing CD14 and producing the respective enzymes:
a, mPGES-1 (n ⫽ 8), b, COX-2 (n ⫽ 4), and c, COX-1 (n ⫽ 4). See Figure 1 for other
definitions.
active product (Figure 3B, part C) eluted at the time
corresponding to PGE2 standard (Figure 3B, part A),
while control cells produced significantly less PGE2
(Figure 3B, part B). These data are consistent with the
EIA results.
Expression of mPGES-1 and COX in synovial
tissue. COX-1–positive staining was observed in the
synovial lining layer cells and in the synovial sublining
mononuclear and fibroblast-like cells (Figure 4A).
Strong mPGES-1 staining was detected in synovial lining
cells and in scattered macrophage- and fibroblast-like
cells in the synovial sublining (Figure 4B). The distribution pattern of COX-2–positive cells in the synovial
lining layer and in the synovial sublining (Figure 4C) was
similar to that of mPGES-1–positive cells (Figure 4D).
However, mPGES-1 staining was observed in vessel
endothelial cells in only some biopsy specimens (5 of
16), while COX-2 staining was detected in endothelial
cells in the majority of synovial specimens (15 of 16). As
demonstrated with double immunofluorescence, intensive mPGES-1 and COX-2 staining colocalized in a
considerable number of cells in the synovial lining layer
(Figure 5A) and in the synovial sublining (Figure 5B). In
contrast, double immunofluorescence showed no increased staining for COX-1 in synovial cells expressing
mPGES-1 (Figures 5C and D).
3444
Figure 3. Prostaglandin E2 (PGE2) generation in synovial fluid mononuclear cell (SFMC) cultures stimulated with lipopolysaccharide (LPS)
and treated with dexamethasone (Dex), infliximab (Inf), or etanercept
(Eta) compared with unstimulated cells. A, Accumulation of PGE2
over the entire treatment period. Values are the mean and SEM (n ⫽
8). B, Conversion of exogenous arachidonic acid to PGE2 by SFMCs,
cultured under the conditions described above, washed, and incubated
with arachidonic acid and calcium ionophore A23187 for 30 minutes.
Values are the mean and SEM (n ⫽ 4). The boxed area shows the
prostaglandin profile for the control and LPS-stimulated SFMCs
incubated with 14C-labeled arachidonic acid. The chromatogram at 195
nm represents the retention times for PGF2␣, PGE2, and PGD2
standards (marked I, II, and III, respectively) (A) and the detection of
radioactive prostaglandin products formed in control and LPSstimulated SFMCs (B and C).
Effects of anti-TNF therapy on mPGES-1 and
COX-2 expression in synovial tissue. COX-2– and
mPGES-1–positive cells were present in the synovial
tissue biopsy samples from all patients, although signif-
Figure 4. Brown immunoperoxidase staining of A, COX-1, B and
D, mPGES–positive cells, and C, COX-2 in representative synovial tissue sections from patients with rheumatoid arthritis. See
Figure 1 for definitions. (Hematoxylin counterstained; original
magnification ⫻ 250.)
KOROTKOVA ET AL
Figure 5. Coexpression of mPGES-1 and COX-2 in synovial tissue
sections from patients with rheumatoid arthritis. A and B, Double
immunofluorescence staining shows mPGES-1–positive (green),
COX-2–positive (red), and double-stained (yellow) cells in A, the
synovial lining layer and B, the synovial sublining. C and D, There was
no increase in COX-1 staining (red) in mPGES-1–positive (green) cells
in C, the synovial lining layer or D, the synovial sublining. See Figure 1
for definitions. (Original magnification ⫻ 250 in A and C; ⫻ 500 in B
and D.)
icant interindividual variations in the extent of the
staining were observed. After therapy with etanercept,
the mPGES-1 staining was higher in 2 patients, lower in
4 patients, and unchanged in 2 patients. The COX-2–
positive area was increased in 2 patients, decreased in 4
patients, and unchanged in 2 patients (Figure 6A). After
treatment with infliximab, the expression of mPGES in
synovial tissue was unchanged in 2 patients, increased in
4 patients, and decreased in 4 patients. COX-2 expression in synovial tissue after treatment with infliximab
was lower in 1 patient, higher in 5 patients, and unchanged in 4 patients (Figure 6B). The differences in the
expression of mPGES-1 as well as COX-2 in synovial
tissue biopsy specimens from RA patients before and
after treatment with TNF blockers were not significant.
Effects of intraarticular steroid therapy on the
expression of mPGES-1 and related enzymes in synovial tissue. After treatment with locally administered
steroids, the expression of mPGES-1 in synovial tissue
was significantly reduced (P ⬍ 0.005), while the expression of housekeeping cPGES was not significantly
changed (Figure 6C). The area stained positive for
COX-1 and COX-2 in synovial tissue was significantly
lower after treatment with intraarticular steroids (P ⬍
0.05) (Figure 6C).
mPGES-1 AND ANTIRHEUMATIC TREATMENT
Figure 6. Expression of microsomal prostaglandin synthase 1
(mPGES-1) and related enzymes in synovial tissue of patients with
rheumatoid arthritis before and after therapy with A, etanercept, B,
infliximab, or C, local intraarticular steroids. Values are expressed as
the mean and SEM percentage of the total area of counterstained
tissue. ⴱ ⫽ P ⬍ 0.05; ⴱⴱ ⫽ P ⬍ 0.005. COX-2 ⫽ cyclooxygenase 2;
cPGES ⫽ cytosolic PGES.
DISCUSSION
In patients with RA, PGE2 levels in synovial fluid
are markedly elevated (32,33), and both activated synovial tissue cells and recruited synovial fluid cells might
contribute to the release of PGE2 into the synovial fluid.
We have previously reported that mPGES-1 is strongly
expressed in synovial tissue from patients with RA,
specifically in synovial macrophages and fibroblasts (15).
Here we analyzed the expression and codistribution of
mPGES-1 and cyclooxygenases (COX-1 and COX-2) in
both synovial fluid cells and synovial tissues from patients with RA. We also studied the effects of different
antirheumatic therapies (TNF blockade and locally administered steroids) on mPGES-1 and cyclooxygenase
expression in these patients in vitro and in vivo. Expression of mPGES-1 and COX-2 in SFMCs was low under
basal conditions and substantially increased after 42
hours of in vitro culture in the presence of LPS (Figures
1A and 2B).
Double immunofluorescence staining revealed
that mPGES-1 was expressed in the cells of the mononuclear phagocyte lineage (Figure 1B). Flow cytometric
analysis demonstrated that COX-2 was induced by LPS
3445
in ⬃35% of the monocytes, whereas 25% expressed
mPGES-1. Approximately 25–30% of the monocytes
under these culture conditions coexpressed COX-2 and
mPGES-1. In comparison, COX-1 was expressed in
15–20% of these cells with or without LPS activation.
Treatment of synovial fluid cells with either TNF blockers or dexamethasone significantly reduced the number
of cells expressing mPGES-1 and COX-2, but not
COX-1. In accordance with this, a strong correlation of
PGE2 formation with the expression profiles of COX-2
and mPGES-1 was observed. Under these experimental
conditions, the induction of PGE2 release by LPS seems
to be mediated through a TNF-dependent pathway.
LPS is known to increase the release of both
TNF␣ and IL-1␤ by monocytes (34), and both cytokines
are known to induce COX-2 and mPGES-1 in synovial
fibroblasts and chondrocytes (10,35). Thus, our results
suggest that TNF blockers also prevent the production
of IL-1␤, as has been previously demonstrated (36).
In RA synovial tissue biopsies, up-regulation of
COX-2 was demonstrated in infiltrating mononuclear
cells, vascular endothelial cells, synovial lining cells, and
sublining fibroblast-like cells (22,37). Although COX-1
expression has also been detected in synovial tissue from
patients with RA (22,37), the role of COX-1 in RA has
not been elucidated. Recently, we have shown strong
mPGES-1 staining in RA synovial lining cells, in sublining macrophage- and fibroblast-like cells, and, in a few
patients, in vascular endothelial cells (15). In the present
study, we found that the distribution pattern of mPGES1–positive cells in RA synovial tissue largely reflected
the distribution of COX-2–positive cells. Moreover,
using double immunofluorescence, we have directly
shown for the first time the colocalization of mPGES-1
and COX-2 in RA synovial lining layer and sublining
cells. The observed coexpression of these 2 enzymes in
RA synovial tissue cells could account for the significant
increase of PGE2 formation, which contributes to inflammatory and destructive processes.
Endothelial cells may also contribute to PGE2
biosynthesis under proinflammatory conditions. For example, endothelial cells of the blood–brain barrier in
vivo, as well as human umbilical vein endothelial cells
and human rheumatoid synovium microvessel endothelium in vitro, have been demonstrated to produce PGE2
after induction with IL-1␤ (38–40). Consistent with this,
we observed endothelial cell staining of mPGES-1 in
approximately one-third of the study patients, and this
finding might be important in, for example, inflammatory angiogenesis. However, the significance of endothelial mPGES-1 expression is presently unclear, and many
3446
factors could be involved in explaining the result, such as
treatments, disease stage, etc. Endothelial cells with the
capacity to produce PGE2 are likely to constitute an
additional cell target for treatment of inflammatory
diseases and fever.
Intraarticular treatment with glucocorticoids efficiently reduces clinical signs of active arthritis, such as
swelling and joint pain. Consistent with our in vitro
results discussed above, locally administered steroids
significantly decreased both mPGES-1 and COX-2 expression in synovial tissue from patients with inflammatory arthritis, including RA (Figure 6C). This suggests
that mPGES-1 might be an additional target for glucocorticoid antiinflammatory action in RA. Intriguingly,
COX-1 staining was also significantly down-regulated
after glucocorticoid treatment. Although COX-1 is generally considered a constitutively expressed enzyme involved in cell homeostasis, recent studies have shown
that it might be induced by different stimuli, for instance
by vascular endothelial growth factor (VEGF) in endothelial cells (41,42) and by IL-1␣ in synovial fibroblasts
(43). VEGF accumulates in SF and is expressed in RA
synovial tissue (44). A suppressive effect of glucocorticoids on VEGF expression has been shown in rheumatoid synovial cells (45). Thus, glucocorticoids might
affect COX-1 expression in RA synovial tissue indirectly
by suppression of VEGF or other unknown factors.
Although treatment of RA with TNF blockers
leads to strong relief of pain and inflammation by a
number of mechanisms, these drugs do not induce
complete remission. Interestingly, in contrast to our in
vitro data, analysis of synovial tissue from RA patients
before and after treatment with etanercept or infliximab
revealed that expression of COX-2 and mPGES-1 was
not significantly down-regulated by TNF blockade (Figures 6A and B). This suggests that either the TNF
blocking effects are not optimal in vivo or, more likely,
that other mechanisms operate in sustaining the inflammation independently of TNF. For instance, it has been
shown that short-term therapy with infliximab reduced
synovial TNF␣ expression in RA patients, while expression of IL-1␤ was not significantly changed (25). Moreover, in mice with experimental arthritis, synovial inflammation was reduced by anti-TNF treatment and
almost completely blocked by a combination of antiTNF and anti–IL-1 treatments (46). In primary cultures
of synovial cells, IL-1␤ seems to be more potent in
inducing mPGES-1 compared with TNF␣ (10). Thus,
TNF blocking agents may not prevent the IL-1␤–
dependent induction of mPGES-1 and PGE2 release,
KOROTKOVA ET AL
which explains the limited therapeutic effects of TNF
blockade in RA.
In conclusion, our results demonstrate that local
treatments with glucocorticoids led to a significant
down-regulation of mPGES-1 as well as COX-1 and
COX-2. In contrast, systemic anti-TNF therapy was
found to be insufficient to suppress the PGE2 pathway in
vivo, albeit not in vitro. Thus, the development of
specific mPGES-1 inhibitors is encouraging since they
might complement TNF blockers for optimal antiinflammatory results in RA.
ACKNOWLEDGMENT
The authors would like to thank Associate Professor
R. A. Harris for linguistic advice.
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